Lee Russell McDowell

Vitamins
inAnimaland
Human
Nutrition
SECOND EDITION

Iowa State University Press / AmesV I TA M I N SINANIMALAND
HUMAN
NUTRITION

Lee Russell McDowell

Vitamins
inAnimaland
Human
Nutrition
SECOND EDITION

Iowa State University Press / Ames

Lee Russell McDowell, PhD, is a professor of animal science in the Department of
Animal Science, University of Florida, Gainesville. His research interests center primarily on minerals for grazing livestock, vitamins for livestock, and feed composition.
Dr. McDowell also collaborates with numerous animal nutritionists in tropical countries of Latin America, Africa, and Southeast Asia.

© 2000 Iowa State University Press; 1989 Academic Press
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Printed on acid-free paper in the United States of America
First edition, 1989 (© Academic Press)
Second edition, 2000
Library of Congress Cataloging-in-Publication Data
McDowell, L. R.
Vitamins in animal and human nutrition/Lee Russell McDowell—2nd ed.
p. cm.
Includes bibliographical references and index.
ISBN 0-8138-2630-6
1. Vitamins in human nutrition. 2. Vitamins in animal nutrition.
3. Avitaminosis. I. Title.
QP771.M396 2000
613.2′86—dc21
The last digit is the print number: 900-027082
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6

5

4

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1

This book is dedicated, with appreciation, to myparents; my wife, Lorraine; my daughters, Suzannah,
Joanna, and Teresa and their husbands and children; and to
Tony J. Cunha (deceased), the former head of the Department of Animal Science at the University
of Florida, for his practical knowledge of the livestock industry and for his support and encouragement to write
books.

CONTENTS
Preface, xv
1. Introduction and Historical Considerations
Definition of Vitamins, 3Classification of Vitamins, 4Vitamin Nomenclature, 5
Vitamin Requirements, 6
Vitamin Occurrence, 7History of the Vitamins, 7References, 13
2. Vitamin A
Introduction, 15
History, 16
Chemical Structure and Properties, 18
Analytical Procedures, 21
Metabolism, 22
Functions, 28
Requirements, 39
Natural Sources, 43
Deficiency, 48
Supplementation, 69
β-Carotene Function Independent of Vitamin A, 75
Toxicity, 78
References, 82
3. Vitamin D
Introduction, 91
History, 91
Chemical Structure, Properties, and Antagonists, 94
Analytical Procedures, 95
vii

CONTENTS

Metabolism, 97
Functions, 103
Requirements, 111
Natural Sources, 115
Deficiency, 117
Supplementation, 134
Toxicity, 140
References, 147
4.

Vitamin E
Introduction, 155
History, 155
Chemical Structure and Properties, 157
Analytical Procedures, 160
Metabolism, 161
Functions, 164
Requirements, 170
Natural Sources, 174
Deficiency, 178
Supplementation, 207
Toxicity, 216
References, 217

5.

Vitamin K
Introduction, 227
History, 227
Chemical Structure, Properties, and Antagonists, 229
Analytical Procedures, 231
Metabolism, 232
Functions, 235
Requirements, 239
Natural Sources, 242
Deficiency, 244
Supplementation, 254
Toxicity, 258
References, 259

6.

ThiaminIntroduction, 265
History, 265
Chemical Structure, Properties, and Antagonists, 267
Analytical Procedures, 269

CONTENTS

Metabolism, 269
Functions, 271
Requirements, 274
Natural Sources, 278
Deficiency, 280
Supplementation, 301
Toxicity, 305
References, 305
7.

Riboflavin
Introduction, 311
History, 311
Chemical Structure, Properties, and Antagonists, 312
Analytical Procedures, 314
Metabolism, 315
Functions, 317
Requirements, 321
Natural Sources, 324
Deficiency, 326
Supplementation, 340
Toxicity, 342
References, 342

8.

NiacinIntroduction, 347
History, 347
Chemical Structure, Properties, and Antagonists, 350
Analytical Procedures, 350
Metabolism, 351
Functions, 355
Requirements, 357
Natural Sources, 360
Deficiency, 362
Supplementation, 376
Toxicity, 379
References, 380

9.

Vitamin B6
Introduction, 385
History, 385
Chemical Structure, Properties, and Antagonists, 386
Analytical Procedures, 388
ix

CONTENTS

Metabolism, 389
Functions, 391
Requirements, 394
Natural Sources, 397
Deficiency, 399
Supplementation, 409
Toxicity, 412
References, 413
10. Pantothenic Acid
Introduction, 419
History, 419
Chemical Structure, Properties, and Antagonists, 420
Analytical Procedures, 422
Metabolism, 423
Functions, 424
Requirements, 426
Natural Sources, 429
Deficiency, 430
Supplementation, 439
Toxicity, 441
References, 441
11. Biotin
Introduction, 445
History, 445
Chemical Structure, Properties, and Antagonists, 446
Analytical Procedures, 448
Metabolism, 448
Functions, 450
Requirements, 452
Natural Sources, 454
Deficiency, 455
Supplementation, 471
Toxicity, 474
References, 475
12. Folacin
Introduction, 479
History, 480
Chemical Structure, Properties, and Antagonists, 481
Analytical Procedures, 484
x

CONTENTS

Metabolism, 485
Functions, 487
Requirements, 490
Natural Sources, 494
Deficiency, 495
Supplementation, 512
Toxicity, 515
References, 516
13. Vitamin B12Introduction, 523
History, 523
Chemical Structure, Properties, and Antagonists, 526
Analytical Procedures, 528
Metabolism, 529
Functions, 532
Requirements, 535
Natural Sources, 538
Deficiency, 540
Supplementation, 555
Toxicity, 558
References, 559
14. Choline
Introduction, 565
History, 565
Chemical Structure and Properties, 566
Analytical Procedures, 567
Metabolism, 568
Functions, 571
Requirements, 574
Natural Sources, 576
Deficiency, 578
Supplementation, 588
Toxicity, 592
References, 592
15. Vitamin C
Introduction, 597
History, 597
Chemical Structure, Properties, and Antagonists, 600
Analytical Procedures, 601
xi

CONTENTS

Metabolism, 602
Functions, 604
Requirements, 609
Natural Sources, 611
Deficiency, 613
Supplementation, 627
Toxicity, 633
References, 634
16. Carnitine
Introduction, 641
History, 641
Chemical Structure and Properties, 641
Analytical Procedures, 642
Metabolism, 642
Functions, 644
Requirements, 645
Natural Sources, 646
Deficiency, 647
Supplementation, 653
Toxicity, 655
References, 655
17. Vitamin-like Substances
Introduction, 659
Myo-Inositol (Inositol), 660
Pyrroloquinoline Quinone (PQQ), 666
p-Aminobenzoic Acid (PABA), 667
Flavonoids (Polyphenols), 668
Lipoic Acid (Thioctic Acid), 669
Coenzyme Q (Ubiquinones), 669
Vitamin B13 (Orotic Acid), 670
Vitamin B15 (Pangamic Acid), 670
Vitamin B17 (Laetrile), 670
Vitamin H3 (Gerovital), 671
Vitamin U (Cabagin), 671
Glucose Tolerance Factor, 671
Other Vitamin-like Factors, 671
References, 673

xii

CONTENTS

18. Essential Fatty Acids
Introduction, 675
History, 675
Chemical Structure and Properties, 676
Analytical Procedures, 678
Metabolism and Functions, 679
Requirements, 689
Natural Sources, 693
Deficiency, 695
Supplementation, 704
Toxicity, 708
References, 708
19. Vitamin Supplementation
Introduction, 713
Factors Resulting in Inadequate Dietary Intakes of Vitamins, 714
Factors Affecting Vitamin Requirements and Utilization, 720
Optimum Vitamin Allowances, 723
Vitamin Supplementation Most Needed by Livestock, 727
Vitamin Supplementation for Humans, 732
Providing Vitamin Supplements, 735
Formulating Vitamin Premixes, 741
References, 742
Appendix: Vitamin Requirements of Livestock and Humans; Feed
Composition; Metric Conversions
A1a: Vitamin Requirements for Ruminants and Horses, 748
A1b: Vitamin Requirements for Monogastric Animals, 750
A1c: Vitamin Requirements for Humans, 756
A2: Composition of Important Feeds, 758
A3: Metric Weights and Measures with Customary Equivalents,
762
Index, 763

xiii

P R E FAC E
Vitamins in Animal and Human Nutrition contains 19 chapters of concise,
up-to-date information on vitamin nutrition for both animals and humans. The
first chapter deals with the definition of vitamins, general considerations, andthe fascinating history of these nutrients. Chapters 2 through 16 discuss the 15
established vitamins in relation to history; chemical structure, properties, andantagonists; analytical procedures; metabolism; functions; requirements;
sources; deficiency; supplementation; and toxicity. Chapter 17 deals with othervitamin-like substances, and Chapter 18 reviews the importance of essential
fatty acids. The final chapter discusses vitamin supplementation considerations.
An earlier edition of this book with a somewhat similar title was published by Academic Press in 1989. The present book has been completely andvigorously revised with one additional chapter. In the last 10 years, a great
deal of new information has been generated in the field of vitamins; this is reflected by the fact that more than half of all the references have been published since the first edition. It is hoped that this book will be of worldwide
use and will continue, as the first edition, to be used as a textbook and as an
authoritative reference book for use by research and extension specialists,
feed manufacturers, teachers, students, and others. An attempt has been madeto provide a balance between animal nutrition and clinical human nutrition.
Likewise, a comparison between the balance of chemical, metabolic, andfunctional aspects of vitamins and their practical and applied considerations
has been made.
A unique feature is the description of the practical implications of vitamin deficiencies and excesses and the conditions that might occur with various animal species and humans. A large number of photographs illustrate vitamin deficiencies in farm livestock, laboratory animals, and humans. Unlike
other textbooks, this one places strong emphasis on vitamin supplementationin each chapter and devotes the last chapter to this subject.
In preparing this book, I have obtained numerous suggestions from eminent scientists both in the United States and in other countries. I wish to express my sincere appreciation to them and to those who supplied photographs
and other material used. I am especially grateful to the following: L.B. Bailey, R.B. Becker, B.J. Bock, H.L. Chapman, J.H. Conrad, G.L. Ellis (dexvP R E FA C E

ceased), R.H. Harms, J.F. Hentges, J.K. Loosli, R. Miles, R.L. Shirley, R.R
Streiff, and W.B. Weaver (Florida); R.T. Lovell and H.E. Sauberlich (Alabama); O. Balbuena, B.J. Carillo, and B. Ruksan (Argentina); H. Heitman(California); J.M. Bell, M. Hidiroglou, and N. Hidiroglou (Canada); N. Ruiz
(Colombia); N. Comben (England); M. Sandholm (Finland); L.S. Jensen(Georgia); T.B. Keith (deceased) (Idaho); A.H. Jensen (Illinois); V. Ramadas
Murthy (India); A. Prabowo (Indonesia); V. Catron (deceased) and V.C. Speer(Iowa); G.L. Cromwell (Kentucky); G.F. Combs (Maryland); F.J. Stare(Massachusetts); D.K. Beede, R.W. Luecke, E.R. Miller, R.C. Piper, J.W.
Thomas, and D.E. Ullrey (Michigan); R.T. Holman and T.W. Sullivan (Minnesota); V. Herbert, L.E. Kook, M.C. Latham, and M.L. Scott (deceased)(New York); A. Helgebostad and H. Rimeslatten (Norway); D.E. Becker(Ohio); P.R. Cheeke, D.C. Church, O.H. Muth, and J.E. Oldfield (Oregon);
D.S. McLaren (Scotland); J.R. Couch and T.M. Ferguson (deceased) (Texas);
D.C. Dobson (Utah); J.P. Fontenot, M.D. Lindemann, and L.M. Potter (Virginia); J.R. Carlson, J.A. Froseth, and L.L. Madsen (deceased) (Washington);and M.L. Sunde (Wisconsin). Appreciation is expressed to company representatives, including G. Patterson (Chas. Pfizer Co); J.C. Bauernfeind, T.M.
Fry, E.L. MacDonald, L.A. Peterson, William Seymour, and S.N. Williams(Hoffmann-LaRoche, Inc.); C.H. McGinnis (Rhône-Poulenc, Inc.); A.T. Forrester (The Upjohn Co.); and M.B. Coelho (BASF Co.). Special thanks go to
J.P. Fontenot for the preliminary planning of the first edition, and to P.R.
Cheeke and J.E. Oldfield for editing and providing useful suggestions for the
first edition of this book.
I am particularly grateful to Nancy Wilkinson and Lorraine M. McDowellfor their useful suggestions and assistance in the editing of the entire book.
Likewise, I wish to acknowledge with thanks and appreciation the skill andcare of Patricia French for overseeing the typing and proofreading of chapters,
and also thank Vanessa Carbia and Mary Schemear for their valuable assistance. Also, I am indebted to the Animal Science Department of the University
of Florida for providing the opportunity and support for this undertaking. Finally, I thank Tony J. Cunha (deceased) for encouraging me to undertake theresponsibility of writing nutrition books and for his expertise on vitamins.

xvi

V I TA M I N SINANIMALAND
HUMAN
NUTRITION

chapter one

INTRODUCTION
AND HISTORICALC O N S I D E R AT I O N S
DEFINITION OF VITAMINS
Vitamins are defined as a group of complex organic compounds
present in minute amounts in natural foodstuffs that are essential to normal metabolism and lack of which in the diet causes deficiency diseases.
Vitamins consist of a mixed group of chemical compounds and are notrelated to each other as are proteins, carbohydrates, and fats. Their classification together depends not on chemical characteristics but on function. Vitamins are differentiated from the trace elements, also present in
the diet in small quantities, by their organic nature.
Vitamins are required in trace amounts (micrograms to milligrams
per day) in the diet for health, growth, and reproduction. Omission of asingle vitamin from the diet of a species that requires it will produce
deficiency signs and symptoms. Many of the vitamins function as coenzymes (metabolic catalysts); others have no such role, but perform certain essential functions.
Some vitamins deviate from the preceding definition in that they do
not always need to be constituents of food. Certain substances that areconsidered to be vitamins are synthesized by intestinal tract bacteria inquantities that are often adequate for body needs. However, a clear distinction is made between vitamins and substances that are synthesized intissues of the body. Ascorbic acid, for example, can be synthesized bymost species of animals, except when they are young or under stress conditions. Likewise, in most species, niacin can be synthesized from theamino acid tryptophan and vitamin D from action of ultraviolet light onprecursor compounds in the skin. Thus, under certain conditions and forspecific species, vitamin C, niacin, and vitamin D would not always fit
the classic definition of a vitamin.
31

CHAPTER ONE

CLASSIFICATION OF VITAMINSClassically, vitamins have been divided into two groups based on
their solubilities in fat solvents or in water. Thus, fat-soluble vitamins
include A, D, E, and K, while vitamins of the B-complex and C are classified water soluble. Fat-soluble vitamins are found in foodstuffs in association with lipids. The fat-soluble vitamins are absorbed along withdietary fats, apparently by mechanisms similar to those involved in fat
absorption. Conditions favorable to fat absorption, such as adequate
bile flow and good micelle formation, also favor absorption of the fatsoluble vitamins (Scott et al., 1982). Water-soluble vitamins are notassociated with fats, and alterations in fat absorption do not affect theirabsorption. Three of the four fat-soluble vitamins (vitamins A, D, andE) are well stored in appreciable amounts in the animal body. Except forvitamin B12, water-soluble vitamins are not well stored, and excesses are
rapidly excreted. A continual dietary supply of the water-soluble vitamins and vitamin K is needed to avoid deficiencies. Fat-soluble vitaminsare excreted primarily in the feces via the bile, whereas water-soluble
vitamins are excreted mainly in the urine. Water-soluble vitamins are
relatively nontoxic, but excesses of fat-soluble vitamins A and D cancause serious problems. Fat-soluble vitamins consist only of carbon,hydrogen, and oxygen, whereas some of the water-soluble vitamins also
contain nitrogen, sulfur, or cobalt.
Table 1.1 lists 14 vitamins classified as either fat or water soluble.
The number of compounds justifiably classified as vitamins is controversial. The term vitamin has been applied to many substances that do
not meet the definition or criteria for vitamin status. Of the 14 vitamins
listed, choline is only tentatively classified as one of the B-complex vitamins. Unlike other B vitamins, choline can be synthesized in the body, isrequired in larger amounts, and apparently functions as a structural
constituent rather than as a coenzyme. Myo-inositol and carnitine are
not listed in Table 1.1 even though they could fit the vitamin category
but apparently for only several species. Chapters 2 through 15 in thisbook concern the 14 vitamins listed in Table 1.1; Chapter 16 is aboutcarnitine; Chapter 17 concerns vitamin-like substances; and Chapter 18
considers essential fatty acids. The essential fatty acids are not vitamins,
but a deficiency disease does result that is similar to vitamin deficiency.
The final chapter deals with vitamin supplementation considerations.

4

INTRODUCTION■ Table 1.1

Fat- and Water-Soluble Vitamins with Synonym Names

Vitamin

Synonym

Fat soluble
Vitamin A1
Vitamin A2
Vitamin D2
Vitamin D3
Vitamin E
Vitamin K1
Vitamin K2
Vitamin K3

Retinol, retinal, retinoic acid
Dehydroretinol
Ergocalciferol
Cholecalciferol
Tocopherol, tocotrienols
Phylloquinone
Menaquinone
Menadionea

Water soluble
Thiamin
Riboflavin
NiacinVitamin B6
Pantothenic acid
BiotinFolacin
Vitamin B12
CholineVitamin C

Vitamin B1
Vitamin B2
Vitamin pp, vitamin B3
Pyridoxol, pyridoxal, pyridoxamine
Vitamin B5
Vitamin H
Folic acid, folate, vitamin M, vitamin Bc
Cobalamin
Gossypine
Ascorbic acidaThe synthetic form is water soluble.

VITAMIN NOMENCLATURE
When the vitamins were originally discovered, they were isolated
from certain foods. During these early years, the chemical compositionof the essential factors was unknown; therefore, these factors wereassigned letters of the alphabet. The system of alphabetizing became
complicated when it was discovered that activity attributed to a single
vitamin was instead the result of several of the essential factors. In thisway, the designation of groups of vitamins appeared (e.g., the vitamin“B” group). Additional chemical studies showed that variations inchemical structure occurred within compounds having the same vitamin
activity but in different species. To overcome this, a system of suffixes
was adopted (e.g., vitamin D2 and D3). The original letter system of designation thus became excessively complicated.
With the determination of the chemical structure of the individual
vitamins, letter designations were sometimes replaced with chemicalstructure names (e.g., thiamin, riboflavin, and niacin). Vitamins havealso been identified by describing a function or its source. The term vitamin H (biotin) was used because the factor protected the haut, the5CHAPTER ONE

German word for skin. Likewise, vitamin K was derived from theDanish word koagulation (coagulation). The vitamin pantothenic acidrefers to its source, as it is derived from the Greek word pantos, meaning “found everywhere.”
The Committee on Nomenclature of the American Institute of
Nutrition (CNAIN, 1981) has provided definite rules for the nomenclature of the vitamins. This nomenclature is used in this book. The official
and major synonym names of vitamins are given in Table 1.1 and in therespective vitamin chapters.

VITAMIN REQUIREMENTS
Vitamin requirements for animals and humans are listed in the
Appendix tables at the end of this book and in the appropriate chapter.
While metabolic needs are similar, dietary needs for vitamins differ
widely among species. Some vitamins are metabolic essentials, but notdietary essentials, for certain species, because they can be synthesized
readily from other food or metabolic constituents.
Poultry, swine, and other monogastric animals are dependent on
their diet for vitamins to a much greater degree than are ruminants.
Tradition has it that ruminants in which the rumen is fully functioning
cannot suffer from a deficiency of B vitamins. It is generally assumed
that ruminants can always satisfy their needs from the B vitamins naturally present in their feed, plus that synthesized by symbiotic microorganisms. However, under specific conditions relating to stress and highproductivity, ruminants have more recently been shown to have requirements, particularly for the B vitamins thiamin (see Chapter 6) and niacin(see Chapter 8). Likewise, vitamin B12 cannot be synthesized in therumen if the essential building block cobalt is lacking in the diet.
The rumen does not become functional with respect to vitamin synthesis for some time after birth. For the first few days of life, the young
ruminant resembles a nonruminant in requiring dietary sources of the B
vitamins. Beginning as early as 8 days, and certainly by 2 months of age,ruminal flora have developed to the point of contributing significant
amounts of the B vitamins (Smith, 1970). Production of these vitaminsat the proximal end of the gastrointestinal tract is indeed fortunate forthey become available to the host as they pass down the tract throughareas of efficient digestion and absorption.
In monogastric animals, including humans, intestinal synthesis ofmany B vitamins is considerable (Mickelsen, 1956) though not as extensive or as efficiently utilized as in ruminants. Low efficiency of utiliza6

INTRODUCTION

tion is probably related to several factors. Intestinal synthesis in nonruminants occurs in the lower intestinal tract, an area of poor absorption. The horse, with a large production of B vitamins in the large intestine, is apparently able to meet most of its requirements for these vitamins in spite of the poor absorption from this area. Intestinally produced vitamins are more available to those animals (rabbit, rat, and others) that habitually practice coprophagy and thus recycle products of thelower gut. This behavior yields significant amounts of B vitamins to thehost animal.

VITAMIN OCCURRENCE
Vitamins originate primarily in plant tissues and are present in animal tissue only as a consequence of consumption of plants, or because
the animal harbors microorganisms that synthesize them. Vitamin B12 isunique in that it occurs in plant tissues as a result of microbial synthesis. Two of the four fat-soluble vitamins, A and D, differ from the watersoluble B vitamins in that they occur in plant tissue in the form of aprovitamin (a precursor of the vitamin), which can be converted into avitamin in the animal body. No provitamins are known for any watersoluble vitamin. However, the amino acid tryptophan can be converted
to niacin for most species. In addition, fat- and water-soluble vitamins
differ in that water-soluble B vitamins are universally distributed in allliving tissues, whereas fat-soluble vitamins are completely absent fromsome tissues.

HISTORY OF THE VITAMINSThe history of the discovery of the vitamins is an inspirational andexciting reflection of the ingenuity, dedication, and self-sacrifice of many
individuals. Excellent reviews of vitamin history with appropriate references include Funk (1922), McCollum (1957), Wagner and Folkers(1962), Maynard et al. (1979), Scott et al. (1982), Widdowson (1986),
and Loosli (1991). Important books that describe the historical discovery of three specific vitamin deficiency diseases include the following:
Eijkman, 1890–1896; Funk, 1911; Williams, 1961 (beriberi); Hess,
1920; Carpenter, 1986 (scurvy); Harris, 1919; and Carpenter, 1981 (pellagra). A brief sketch of important events emphasizing early history ofvitamins is outlined in Table 1.2.
The existence of nutritive factors, such as vitamins, was not recognized until about the start of the twentieth century. The word “vitamin”
7

■ Table 1.22697 B.C.

Brief History of Vitamins (Ancient History–1951)

Beriberi was recognized in China and is probably the earliest documented
deficiency disorder.
1500 B.C.
Scurvy, night blindness, and xerophthalmia were described in ancient
Egypt. Liver consumption was found to be curative for night blindness andxerophthalmia.
130–200 A.D Soranus Ephesius provided classical descriptions of rickets.
1492-1600
World exploration threatened by scurvy:
Magellan lost four-fifths of his crew.
Vasco da Gama lost 100 of his 160 men.
1747
Lind performed controlled shipboard experiments on the preventive effectof oranges and lemons on scurvy. He also developed a method of preserving citrus juice by evaporation and conserving in acid form.
1768–1771
Captain Cook demonstrated that prolonged sea voyages were possible
without ravages of scurvy.
1816
Magendie described xerophthalmia in dogs fed carbohydrate and olive oil.
1824
Combe described a fatal anemia (pernicious anemia) and suggested that it
could be related to a disorder of the digestive tract.
1849
Choline was isolated by Streker from the bile of pigs.
1881
Lunin reported that animals did not survive on diets composed solely of
purified fat, protein, carbohydrate, salts, and water.
1880sJapanese physician Takaki prevented beriberi in the Japanese Navy by substituting other foods for polished rice.
1897
Eijkman showed that beriberi (thiamin deficiency) from polished rice consumption could be cured by adding rice polishings back into the diet.
1901
Grijins concluded that beriberi was caused by a vitally important food constituent.
1906
Hopkins suggested that substances in natural foods, termed “accessory
food factors,” were indispensable and did not fall into the categories ofcarbohydrate, fat, protein, or mineral.
1907
Holst and Frolich produced experimental scurvy in guinea pigs by feedinga deficient diet, with pathological changes resembling those in humans.
1909
Hopkins reported a rat growth factor in some fats.
1910
Vedder was convinced that beriberi was caused by a nutritional deficiency
and saved many lives in the Philippines by feeding rice polishings.
1911
The term “vitamine” was first used by the Polish biochemist Funk to
describe an accessory food factor.
1913
McCollum and Davis discovered fat-soluble A in butter that was associated with growth.
1919
Steenbock reported that the yellow color (carotene) of vegetables was vitamin A.
1919
Mellanby produced rickets in dogs, which responded to a fat-soluble vitamin in cod liver oil.
1920
Goldberger reported that pellagra was not caused by bacterial infection,
but rather was an ill-balanced diet high in corn.
1922
McCollum established vitamin D as independent of vitamin A by preventing rickets after destroying vitamin A activity when bubbling oxygen
through cod liver oil.
1923
Evans and Bishop discovered vitamin E. The deficiency caused female rats
to abort, while male rats became sterile.
1926
Jansen and Donath isolated thiamin in crystalline form from rice bran.
1926
Minot and Murphy showed that large amounts of raw liver given bymouth daily would alleviate pernicious anemia.

8■ Table 1.2

continued

1926

Steenbock showed that irradiation of foods as well as animals produced
vitamin D.
Goldberger and Lillie described a rat syndrome, later shown to be
riboflavin deficiency.
Bechtel and coworkers established that rumen bacteria of cattle synthesized
B vitamins.
Szent-Györgyi isolated hexuronic acid (ascorbic acid, vitamin C) fromorange juice, cabbage juice, and cattle adrenal glands.
Moore proved that the animal body converts carotene to vitamin A.
Norris and coworkers reported a curled-toe paralysis (riboflavin deficiency) in chicks.
Castle showed that pernicious anemia resulted from the interaction of a
dietary (extrinsic) factor and an intrinsic factor produced by the stomach.
Norris and Ringrose described a pellagra-like dermatitis in the chick, later
established as a pantothenic acid deficiency.
Pappenheimer and Goettsch showed that vitamin E is required for prevention of encephalomalacia of chicks and nutritional muscular dystrophy in
rabbits and guinea pigs.
Willis demonstrated that a factor from yeast was active in treating a tropical macrocytic anemia seen in women of India.
Choline was discovered to be the active component of pure lecithin previously shown to prevent fatty livers in rats.
Williams and associates fractionated a growth factor from yeast andnamed it pantothenic acid.
György named a factor that would cure dermatitis in young rats, vitamin
B6.
Dam and Schönheyder described a nutritional disease of chickens characterized by bleeding, thus a new fat-soluble vitamin was discovered (vitamin
K).
Wald demonstrated the relation of vitamin A to night blindness and vision.
Kuhn in Germany and Karrer in Switzerland synthesized riboflavin.
Warburg and coworkers first demonstrated a biochemical function for
nicotinic acid when they isolated it from an enzyme (NADP).
Cobalt, the central ion in vitamin B12, was shown to be a dietary essential
for cattle and sheep by Underwood and coworkers in Australia and in
Florida by Becker and associates.
Biotin was the name given to a substance isolated from egg yolk by Kogl
and Tonnis that was necessary for the growth of yeast.
Williams and colleagues determined the structure of thiamin and synthesized the vitamin.
Elvehjem and associates found that nicotinic acid cured black tongue indogs. It was quickly shown to be effective for pellagra in humans.
Vitamin K was isolated by Dam and Karrer of Europe and a few months
later in the U.S. from three different laboratories.
Harris and associates completed the first synthesis of biotin.
Baxter and Robeson crystallized vitamin A.
Chemists from the Lederle group crystallized and later synthesized folacin.
Rickes and coworkers in the U.S. and Smith in England isolated vitamin
B12.
Smith and coworkers showed that cobalt deficiency in sheep could be prevented by vitamin B12 injection.

1926
1928
1928
1929
1929
1929
1930
1931
1931
1932
1933
1934
1934
1935
1935
1935
1935–1937
1936
1936
1937
1939
1940
1942
1943–1946
1948
1951

9

CHAPTER ONE

had not been coined yet. However, what were to be later known as vitamin-deficiency diseases, such as scurvy, beriberi, night blindness,
xerophthalmia, and pellagra, had plagued the world at least since theexistence of written records. Records of medical science from antiquity
attesting to human association of certain foods with either the cause or
prevention of disease and infirmity are considered the nebulous beginnings of the concept of essential nutrients (Wagner and Folkers, 1962).
Even so, at the beginning of the twentieth century, the value of food inhuman nutrition was expressed solely in terms of its ability to provide
energy and basic building units necessary for life.
In the late 1800s and early 1900s, some scientists believed that life
could be supported with chemically defined diets. In 1860, Louis Pasteur
reported that yeast could grow on a medium of sugar, ammonium salts,
and ash of yeast. Justus von Liebig observed that certain yeasts wereunable to grow at all under these conditions, while others grew only very
slowly. The ensuing arguments between Liebig and Pasteur did not solve
the question. Pasteur’s (1822–1895) research showing that bacteriacaused disease led scientists trained in medicine to be reluctant to believe
the “vitamin theory” that certain diseases resulted from a shortage of
specific nutrients in foods (Loosli, 1991). Guggenheim (1995) suggested
that the immensely successful germ theory of disease, with the related
toxin theory and success of using antisepsis and vaccination, occupied
the thoughts of scientists at that time with the idea that only a positive
agent could cause a disease.
The first phase leading to the “vitamin hypothesis” began with
gradual recognition that the cause of diseases such as night blindness,
scurvy, beriberi, and rickets could be related to diet. Although the truecause, nutritional deficiency, was not suspected, these results marked thefirst uncertainty in the germ and infection theories of origin for these diseases. Finally, in the early 1900s, many scientists in the field of nutrition
almost simultaneously began to realize that a diet could not be adequately defined in terms of carbohydrate, fat, protein, and salts. At thattime, it became evident that other organic compounds had to be present
in the diet if health was to be maintained.
Beriberi was probably the earliest documented deficiency disorder,
being recognized in China as early as 2600 B.C. Scurvy, night blindness,and xerophthalmia were described in the ancient Egyptian literaturearound 1500 B.C. Substances rich in vitamin A as remedies for night
blindness were used very early by the Chinese, and livers were recommended as curative agents for night blindness and xerophthalmia by

10

INTRODUCTION

Hippocrates around 400 B.C. In 1536, Canadian Indians cured Jacques
Cartier’s men of scurvy with a broth of evergreen needles. In 1747,
James Lind, a British naval surgeon, showed that the juice of citrus fruits
was a cure for scurvy, but its routine use was not started in the British
Navy until 1795. Cod liver oil was used as a specific treatment for rickets long before anything was known about the cause of this disease, andwas fed to farm animals as early as 1824. In the 1880s, the Japanesephysician Takaki recognized the cause of beriberi in the Japanese Navyas stemming from an unbalanced white rice diet, and virtually eliminated this condition by increasing the consumption of vegetables, fish, andmeat and by substituting barley for rice.
The period before the close of the nineteenth century was characterized by the discovery of diseases of nutritional origin in animals,
which opened the way for controlled experimental studies of nutritional causes and cures for diseases that were common to both humans andthe lower animals. The rat undoubtedly contributed most to the discovery of vitamins from 1900 through the 1920s, although chickens,
pigeons, guinea pigs, mice, and dogs also played their part (Widdowson,1986). In 1890, Christiaan Eijkman, a Dutch physician working in a
military hospital in Java, found that chickens fed almost exclusively on
polished rice developed polyneuritic signs bearing a marked resemblance
to those of beriberi in humans. A new head cook at the hospital discontinued the supply of “military” rice (polished), and thereafter the birds
were fed on whole-grain “civilian” rice, with the result being that they
recovered. He also noted that beriberi in prisoners eating polished ricetended to disappear when a less highly milled product was fed. Many
great advances in science have started from such chance observations
pursued by men and women of inspiration.
Beginning in the middle 1850s, German scientists were recognized
as leaders of nutrition. In the late 1800s, Professor C. von Bunge(Dorpat, Estonia, Germany, and then at Basel) had graduate students
experimenting with purified diets for small animals (Wolf andCarpenter, 1997). In 1881, N. Lunin, a Russian student studying in vonBunge’s laboratory, observed that mice died (16–36 days) when fed adiet composed solely of purified fat, protein, carbohydrate, salts, andwater. Lunin proposed that natural foods such as milk contain small
quantities of as yet “unknown substances essential to life.” Other
researchers from von Bunge’s school and under his influence had essentially the same results as Lunin; these researchers included C.A. Socin(1891), W.S. Hall (1896), W. Falta (1906), and C.T. Noeggerath (1906).

11

CHAPTER ONE

Von Bunge explained away results of these experiments as he wasinclined to disbelieve the existence of unknown nutritional factors. Von
Bunge believed that iron and phosphorus must be present in preformed
organic combinations, which was his explanation for the deaths of laboratory animals consuming purified diets (Wolf and Carpenter, 1997).
In 1906, Frederick Hopkins in England suggested that unknown
nutrients were essential for animal life and used the term “accessory
growth factors.” Hopkins was responsible for opening up a new field ofdiscovery that largely depended on the use of the rat. When Hopkins
later discovered that he was not the first to suggest that unknown nutrients were essential, or to conduct animal experiments, he was anxious
to share his Nobel prize with Eijkman in 1929.
In 1911, Casimir Funk proposed the “vitamine theory.” He hadreviewed the literature and made the important conclusion that beriberi
could be prevented or cured by a protective factor present in natural
food, which he had isolated from rice by-products (Funk, 1911). Funk
named the distinct factor that prevented beriberi a “vitamine.” This
word was derived from “vital amine.” Later, when it became evident
that not all “vitamines” contained nitrogen (amine), the term became“vitamin.” Funk had not believed that all “vitamines” were amines;
rather, the name was chosen as a catchword to create interest in the new
emerging field of nutrition.
After reviewing the literature between 1873 and 1906, in whichsmall animals had been fed restricted diets of isolated proteins, fats, andcarbohydrates, E.V. McCollum of the United States noted that the animals rapidly failed in health and concluded that the most important
problem in nutrition was to discover what was lacking in such diets. By
1915, McCollum and M. Davis of Wisconsin discovered that the rat
required at least two essential growth factors: a “fat-soluble A” factor
and a “water-soluble B” factor. In addition to being required as factors
for normal growth, the “fat-soluble A” factor was found to cure
xeropththalmia, and the “water-soluble B” factor cured beriberi. At thesame time as their work in Wisconsin, T.B. Osborne and L.B. Mendel ofConnecticut also established the importance of what was later named
vitamin A.
With the pioneer work of Eijkman, Hopkins, Funk, McCollum, andothers, scientists began to seriously consider the new class of essential
nutrients. The brilliant research of scientists in the first half of the twentieth century led to the isolation of more than a dozen vitamins as purechemical substances. The golden age of vitamin research was mainly in

12

INTRODUCTION

the 1930s and 1940s. For vitamin discovery, the general procedure
employed was first to study the effects of a deficient diet on a laboratory animal and then to find a food that would prevent the deficiency.
Using a variety of chemical manipulations, the particular nutrient
involved was gradually concentrated from the food, and its potency wastested at each stage of concentration on further groups of animals(Wagner and Folkers, 1962). This laborious procedure has been simplified in recent years by the discovery that several vitamins are also
growth factors for microorganisms that can therefore replace animals
for potency testing. By such methods, it is now possible to isolate vitamins and subsequently to identify them chemically. A remarkableachievement has been the direct synthesis by chemists of at least ten vitamins identified in this way. The last vitamin to be discovered was vitamin B12 in 1948, which brought the period of vitamin discovery to aclose. On the other hand, the possibility that there are still undiscovered
vitamins must be recognized (see Chapter 17). More detailed historicalconsiderations for each vitamin are presented in the respective chapters(Chapters 2 through 16).

■ REFERENCES
Carpenter, K.J. (1981). In Pellagra, Hutchinson Ross Publishing Company,
Stroudsburg, Pennsylvania.
Carpenter, K.J. (1986). In The History of Scurvy and Vitamin C. Cambridge
University Press, Cambridge, London.
CNAIN. (1981). (Committee on Nomenclature of the American Institute of
Nutrition). J. Nutr. 111, 8.
Eijkman, C. (1890–1896). Polyneuritis in Chickens, or the Origin of Vitamin
Research. Roche, Basel.
Funk, C. (1911). J. Physiol. 43, 395.
Funk, C. (1922). The Vitamins, Williams and Wilkins Co., Baltimore.
Guggenheim, K.Y. (1995). In Basic Issues of the History of Nutrition, The
Magnes Press, Hebrew University, Jerusalem, Israel.
Harris, H.F. (1919). Pellagra, The Macmillan Co., New York.
Hess, A.F. (1920). Scurvy Past and Present, J.B. Lippincott Co., London.
Loosli, J.K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.)Academic Press, San Diego.
Maynard, L.A., Loosli, J.K., Hintz, H.F., and Warner, R.G. (1979). Animal
Nutrition, 7th Ed., p. 283. McGraw-Hill, New York.
McCollum, E.V. (1957). In A History of Nutrition, p. 217. Houghton Mifflin,
Boston.
Mickelsen, O. (1956). Vitam. Horm. 14, 1.

13

CHAPTER ONE

Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Smith, S.E. (1970). In Duke’s Physiology of Domestic Animals, 8th Ed. p. 634(M.J. Swenson, ed.), Cornell University Press, Ithaca, New York.
Wagner, A.F., and Folkers, K. (1962). Vitamins and Coenzymes, Wiley(Interscience), New York.
Widdowson, E.M. (1986). Nutr. Rev. 44, 221.
Williams, R.R. (1961). Toward the Conquest of Beriberi, Harvard University
Press, Cambridge.
Wolf, G., and Carpenter, K.J. (1997). J. Nutr. 127, 1255.

14

chapter two

V I TA M I N A

INTRODUCTION
Although all vitamins are equally important in supporting animal
life, vitamin A may be considered the most important vitamin from apractical standpoint. It is important as a dietary supplement for all animals, including ruminants. Vitamin A itself does not occur in plants;
however, its precursors (carotenoids) are found in plants, and these canbe converted to true vitamin A by a specific enzyme located in the intestinal walls of animals. Prior to the discovery of vitamin A, farmers complained that hogs in dry lot or barns did poorly when fed a ration consisting largely of white corn instead of yellow corn. Agriculturalchemists would disagree and explain to farmers that chemical analysis
showed that white corn and yellow corn were the same with the exception of color. Then came the vitamin era, which explained what thefarmers already knew, that white corn has no carotene, the precursor ofvitamin A (Ensminger and Olentine, 1978).
In human nutrition, vitamin A is one of the few vitamins of which
both deficiency and excess constitute a serious health hazard. Deficiency
occurs in endemic proportions in many developing countries and is considered to be the most common cause of blindness in young children
throughout the world. McLaren (1986) lists 73 countries and territories
that are considered to have potentially serious vitamin A deficiency
problems. Vitamin A toxicity usually arises from abuse of vitamin supplementation.

15

2

CHAPTER TWOHISTORY
For thousands of years humans and animals have suffered from vitamin A deficiency, typified by night blindness and xerophthalmia (a condition named for the Latin words for dry eye, a manifestation of vitamin
A deficiency in which the conjunctiva [covering of the eye] dries out, thecornea becomes inflamed, and the eye becomes ulcerated). The cause
was unknown, but it was recognized that consumption of animal andfish livers had curative powers according to records and folklore fromearly civilizations. One of the earliest known reports was from Eber’s
Papyrus, an ancient Egyptian medical treatise of about 1500 B.C., which
recommended the livers of cattle or poultry as curative agents(Arykroyd, 1958). An early reference to vitamin A deficiency in livestock is the Bible (Jeremiah 14:6): “and the asses did stand in highplaces, their eyes did fail, because there was no grass.” Also from the
Bible was the cure of the blind Tobias by means of fish bile.
The observation that experimental animals lose weight and die on
purified diets was noted by many investigators toward the end of thenineteenth century. However, it was not until early in the twentieth century that vitamin A was discovered. Its history has been reviewed by anumber of authorities (Funk, 1922; McCollum, 1957; Sebrell andHarris, 1967; Loosli, 1991). From 1906 through 1912, Hopkins ofGreat Britain found that a growth-stimulating principle from milk waspresent in an alcoholic extract of milk rather than in the ash. In 1909,
Hopkins and Stepp found that certain fat-soluble substances were necessary for growth of mice and rats. In the years 1913 through 1915,
McCollum and Davis described “fat-soluble A,” a factor isolated fromanimal fats (unsaponifiable fraction of milk fat) or fish oils, which they
associated with a growth-promoting activity. In their experiments, thegrowth of rats ceased prematurely when lard was used as the source of
fat in the diet, whereas adequate growth was obtained when the dietary
fat was either butter or fat extracted from egg yolk. At the same time,
Osborne and Mendel also reported that something in butter appeared tobe essential for life and growth in rats. Later, Drummond suggested thatthe “fat-soluble factor A” should be named vitamin A. In 1919,
Steenbock called attention to the fact that among vegetable foods, vitamin A potency was associated with yellow color. He suggested thatcarotene was the source of the vitamin, but later recognized that thevitamin was not carotene itself because certain potent sources of thevitamin were colorless. Ten years later, Von Euler and associates in16

V I TA M I N A

Stockholm obtained a definite growth response when carotene wasadded to vitamin A-deficient diets. In 1929, Moore produced proof thatthe animal body transformed carotene into vitamin A. Animals fed
carotene had vitamin A in livers, whereas controls did not.
Research in the 1920s and 1930s demonstrated that most animal
species need dietary vitamin A. The simultaneous use of chemical methods and experimental rats to test metabolic products resulted in the successful demonstration of vitamin A activity, making it the first confirmed
vitamin rather than vitamin B or C, which had received earlier attention.
Similar testing methods were used to identify most of the other vitamins.
Only a few years after vitamin A was discovered, it was thought thatrickets was also a vitamin A deficiency. Proof that rickets was not causedby vitamin A deficiency was provided by McCollum and associates in1922. This proof was obtained by oxidizing cod liver oil until vitamin A
was destroyed, as shown by the inability of the oil to cure xerophthalmia, and then by demonstrating that the oxidized oil was still effective in curing rickets.
Vitamin A deficiency was shown to be responsible forxeropththalmia and certain forms of night blindness. A link between
vitamin A and the visual process was demonstrated in 1935 when Wald,in a series of experiments, obtained a specific form of vitamin A (retinal)from bleached retinas. Wagner and coworkers suggested in 1939 thatthe conversion of β-carotene into vitamin A occurs within the intestinal
mucosa. In 1944, Morton suggested that retinal from bleached visual
purple (rhodopsin) might be identical with vitamin A aldehyde; he wasable to prove this by synthesis.
The isolation of pure vitamin A became possible when a relationshipwas found between its growth-promoting activity and the intensity of
the Carr-Price antimony trichloride color at 620 nm or the light absorption at 328 nm. Karrer and his group were thus able to obtain a pureoily retinol from vitamin A-rich concentrates. From 1930 to 1931,
Karrer and coworkers proposed the exact structural formulas for vitamin A and β-carotene. Six years later, the first crystals of vitamin A wereobtained, and still another growth-promoting factor-vitamin A2 was isolated from freshwater fish liver oils.
In 1942, Baxter and Robeson crystallized pure vitamin A and several of its esters; five years later, they also succeeded in isolation andcrystallization of the 13-cis-vitamin A isomer. Isler and coworkers synthesized the first pure vitamin A in 1947. In 1950, Karrer and Inhoffen
reported the synthesis of β−carotene. In the early 1980s, β−carotene andother carotenoids began to be recognized as important factors (inde17

CHAPTER TWO

pendent of provitamin A activity) in potentially reducing the risk of certain cancers and other disease conditions.

CHEMICAL STRUCTURE AND PROPERTIES
Vitamin A itself does not occur in plant products, but its precursor,
carotene (Fig. 2.1) occurs in several forms. These compounds are commonly referred to as provitamin A because the body can transform theminto the active vitamin. This is how the vitamin A needs of farm animals
are met, for the most part, because their rations consist mainly or entirely of foods of plant origin. The combined potency of a feed, represented
by its vitamin A and carotene content, is referred to as its vitamin A
value. Retinol is the alcohol form of vitamin A (Fig. 2.1). Replacementof the alcohol group by an aldehyde group gives retinal, and replacement by an acid group gives retinoic acid. Esters of retinol are called
retinyl esters. Vitamin A in animal products exists in several forms, but
principally as long-chain fatty acid esters (e.g., retinyl palmitate).
In addition to retinol, there is another form that is isolated fromfish. It was originally distinguished on the basis of a different maximum
spectral absorption and named A2 to differentiate it from the previously isolated form. Vitamin A2 is closely related to vitamin A1 but contains
an additional double bond in the β−ionone ring (Fig. 2.1). Liver oils ofmarine fish origin usually average less than 10% vitamin A2 of the total
vitamin A content. The relative biological activity of vitamin A2 is 40 to
50% that of A1.
Vitamin A is a nearly colorless, fat-soluble, long-chain, unsaturated
alcohol with five double bonds. The vitamin is made up of isoprene units
with alternate double bonds, starting with one in the β−ionone ring thatis in conjugation with those in the side chain (Fig. 2.1). Since it contains
double bonds, vitamin A can exist in different isomeric forms. Morecommon isomeric forms of vitamin A and their relative biological activities are presented in Fig. 2.2.
The most active vitamin A form and that most usually found in
mammalian tissues is the all-trans-vitamin A. cis-Forms can arise fromthe all-trans-forms, and a marked loss of vitamin A potency results.
These structural changes in the molecule are promoted by moisture,heat, light, and catalysts. Therefore, conditions present during hay making and ensiling, dehydrating, and storage of crops are detrimental to the
biological activity of any carotenoids present.
Precursors of vitamin A, the carotenes, occur as orange-yellow pigments mainly in green leaves and to a lesser extent in corn. Of more than18

V I TA M I N A

Fig. 2.1 Chemical structure of vitamin A1, β-carotene, and vitamin A2.

500 carotenoids that have been isolated from nature, only 50 to 60 possess biological activity. Structures of some of the important carotenoid
pigments and their distribution and relative biological activity are presented in Fig. 2.3. Four of these carotenoids—α-carotene, β−carotene, γcarotene, and cryptoxanthine (the main carotenoid of corn)—are of particular importance because of their provitamin A activity. Vitamin A
activity of β−carotene is substantially greater than that of othercarotenoids. Lycopene is an important carotenoid for its antioxidant
function but does not possess the β−ionone ring structure, and therefore
is not a precursor of vitamin A. In humans, β−carotene and lycopene are19CHAPTER TWO

Fig. 2.2 Isomers of vitamin A (retinol). (Adapted from Ullrey, 1972.)

the predominant carotenoids in tissue (Ribaya-Mercado et al., 1995).
Theoretically, 1 mol of β−carotene could be converted (cleavage of
the C15=C15´ bond) to yield 2 mol of retinal. However, biological tests
have consistently shown that pure vitamin A has twice the potency of β−carotene on a weight-to-weight basis. Thus, only one molecule of vitamin A is formed from one molecule of β−carotene. Loss of potential
activity results from inefficient cleavage and intestinal absorption.
Vitamin A activity is expressed in international units (IU) or, less frequently, in United States Pharmacopeia (USP) Units, both of which areof equal value. An IU is defined as the biological activity of 0.300 μg ofvitamin A alcohol (retinol) or 0.550 μg of vitamin A palmitate. One IUof provitamin A activity is equal in activity to 0.6 μg of β−carotene, thereference compound. Vitamin A may be expressed as retinol equivalents(RE) instead of IU. By definition, 1 retinol equivalent is equal to 1 μg of20

V I TA M I N A

Fig. 2.3 The yellow carotenoids. (Adapted from Ullrey, 1972.)

retinol, 6 μg of β−carotene, or 12 μg of other provitamin A carotenoids.
In terms of international units, 1 RE is equal to 3.33 IU of retinol or 10
IU of β−carotene.

ANALYTICAL PROCEDURES
A number of methods are available for carotene and vitamin A
determination (Pit, 1985). Biological methods include growth responsesof rats or chicks, the storage test (liver), and quantitative evaluations ofcell changes in vaginal smears (rats). Physicochemical methods include

21

CHAPTER TWO

color reactions with antimony trichloride (Carr-Price method), gas chromatography, thin-layer chromatography, and spectrophotometric procedures. A number of reports (Grace and Bernhard, 1984; Hidiroglou etal., 1986; Horst et al., 1995) indicate excellent results and high recovery
rates from high-pressure liquid chromatography (HPLC). The HPLC
procedure is the most common method for analyzing carotenoids, vitamin A and its analogs in pharmaceutical preparations, feedstuffs, andtissues combined with an ultraviolet (UV) detector. A procedure forretinol-binding protein is radioimmunoassay (Vallet, 1994).

METABOLISM
Digestion
Vitamin A in animal products and carotenoids are released fromproteins by the action of pepsin in the stomach and proteolytic enzymes
in the small intestine (Ong, 1993; Ross, 1993). In the duodenum, bilesalts break up fatty globules of carotenoids and retinyl esters to smaller
lipid congregates, which can be more easily digested by pancreatic
lipase, retinyl ester hydrolase, and cholesteryl ester hydrolase.
A number of factors influence digestibility of carotene and vitamin
A. Working with lambs, Donoghue et al. (1983) reported that dietarylevels of vitamin A ranging from mildly deficient to toxic levels affectdigestion and uptake. Percentage transfer from the digestive tract fromsupplemental dietary levels of 0, 100, and 12,000 μg of retinol per kilogram were 91, 58, and 14%, respectively. Wing (1969) reported that theapparent digestibility of carotene in various forages fed to dairy cattle
averaged about 78%. Variables that influenced carotene digestibility
included month of forage harvest, type of forage (hay, silage, greenchop, or pasture), species of plant, and plant dry matter. In general,carotene digestibility was higher than average during warmer months
and lower than average during winter.
Several reports indicate that appreciable amounts of carotene orvitamin A may be degraded in the rumen. Various studies with different
diets have resulted in preintestinal vitamin A disappearance values ranging from 40 to 70% (Ullrey, 1972). Rode et al. (1990) compared microbial degradation of vitamin A (retinyl acetate) from steers fed concentrate, hay, or straw diets. Estimated effective rumen degradation of biologically active vitamin A was 67% for cattle fed concentrates compared
to 16 and 19% for animals fed hay and straw diets, respectively.

22

V I TA M I N A

Absorption and Transport
Much of the conversion of β−carotene to vitamin A takes place in
the intestinal mucosa. Provitamin A carotenoids must contain one
unsubstituted β−ionone ring to be active. This conversion of β−carotene
into vitamin A involves two enzymes. β−Carotene-15,15´-dioxygenasecatalyzes the cleavage of β−carotene at the central double bond to yield
two molecules of retinal for one molecule of β−carotene. However,extensive evidence exists also for random (eccentric) cleavage, resulting
in retinoic acid and retinal, with a preponderance of apocarotenals
formed as intermediates (Wolf, 1995). The cleavage enzyme has beenfound in many vertebrates but is not present in the cat or mink.
Therefore, these species cannot utilize carotene as a source of vitamin A.
The second enzyme, retinaldehyde reductase, reduces the retinal toretinol.
In most mammals, the product ultimately absorbed from the intestinal tract as a result of feeding carotenoids is mainly vitamin A itself.
There is considerable species specificity regarding the ability to absorb
dietary carotenoids. In some species, such as the rat, pig, goat, sheep,
rabbit, buffalo, and dog, almost all of the carotene is cleaved in theintestine. In humans, cattle, horses, and carp, significant amounts ofcarotene can be absorbed. Absorbed carotene can be stored in the liverand fatty tissues. Hence, these latter animals have yellow body and milkfat, whereas animals that do not absorb carotene have white fat.
In the case of cattle, there is a strong breed difference in absorptionof carotene. The Holstein is an efficient converter, having white adipose
tissue and milk fat. The Guernsey and Jersey breeds, however, readily
absorb carotene, resulting in yellow fat. The chick, on the other hand,absorbs only hydroxy carotenoids in the unchanged form and storesthem in tissues. Hydrocarbon carotenoids with provitamin A activity areconverted by the chick intestine and absorbed as vitamin A. Species
specificity in vitamin A conversion may be due to presence or absence ofsuitable receptor proteins or the ability to form suitable micellar solutions in the intestinal lumen.
A number of factors affect absorption of carotenoids. cis-trans-isomerism of the carotenoids is important in determining their absorbability, with the trans-forms being more efficiently absorbed (Stahl et al.,1995). Among carotenoids there is a differential uptake and clearance ofspecific carotenoids (Bierer et al., 1995; Johnson et al., 1997; Erdman etal., 1998; Van den Berg, 1999). Dietary fat is important (Roels et al.,23

CHAPTER TWO

1958; Fichter and Mitchell, 1997; Takyi, 1999). When small supplements of fat were given to vitamin A-deficient boys in a region ofCentral Africa, the absorption of dietary carotenoids increased remarkably (from less than 5% to about 50%) (Roels et al., 1958). Dietary
antioxidants (e.g., vitamin E) also appear to have an important effect on
the utilization and perhaps absorption of carotenoids. It is uncertain
whether the antioxidants contribute directly to efficient absorption orwhether they protect both carotene and vitamin A from oxidative breakdown. Protein deficiency reduces absorption of carotene from the intestine.
Almost no absorption of vitamin A occurs in the stomach. The mainsite of vitamin A and carotenoid absorption is the mucosa of the proximal jejunum. The absorption of vitamin A in the intestine is believed to
be 80 to 90%, while that of β−carotene is about 50 to 60% (Olson,
1991). Vitamin A efficiency of absorption decreases somewhat with very
high doses. Intestinal β−carotene cleavage activity was shown to behigher in rats deficient in vitamin A than in rats with a high intake ofeither vitamin A or β−carotene (Van Vliet et al., 1996). Carotenoids arenormally converted to retinol in the intestinal mucosa but may also be
converted in the liver and other organs, especially in yellow fat species
such as poultry (McGinnis, 1988). Lipid micelles in the intestinal lumen
serve as carriers by taking up vitamin A and carotene from emulsified
dietary lipid and bringing these lipids into contact with the mucosal cell,
where they diffuse from the micelle through the lipid portion of the
microvillar membrane.
Vitamin A occurs in food primarily as the palmitate ester. This ishydrolyzed in the small intestine by retinyl ester hydrolase, which issecreted by the pancreas. Bile salts are required both for the activationof this enzyme and for the formation of the lipid micelle, which carries
vitamin A from the emulsified dietary lipid to the microvillus. Normally,
vitamin A is absorbed almost exclusively as the free alcohol, retinol.
Within the mucosal cells, retinol is reesterified mostly to palmitate, isincorporated into the chylomicra of the mucosa, and is secreted into thelymph. A small amount of retinol may be oxidized first to retinal andthen to retinoic acid, which may form a glucuronide and pass into theportal blood. A specific transporter in the intestinal brush border for alltrans- and 3-dehydroretinol, constituting a process of carrier-facilitateddiffusion, was reported, while other retinoids studied were taken up bypassive diffusion at a slower rate (Wolf, 1995). At normal intakes,
uptake of β−carotene by the rat was linear suggesting passive uptake(Moore et al., 1996).
24

V I TA M I N A

Vitamin A is transported through the lymphatic system with a lowdensity lipoprotein in lymph acting as a carrier to the liver, where it isdeposited mainly in hepatocytes and stellate and parenchymal cells.
When liver stores of vitamin A are adequate, it can be transferred fromparenchymal cells to stellate cells, where it is reesterified (Blomhoff etal., 1991).
When vitamin A is mobilized from the liver, stored vitamin A esteris hydrolyzed (retinyl ester hydrolase) prior to its release into the bloodstream, and vitamin A alcohol (retinol) then travels via the bloodstream
to the tissues. Retinol is transported by a specific transport and binding
protein, retinol-binding protein (RBP). The RBP is synthesized andsecreted by hepatic parenchymal cells. Human RBP has a molecularweight of about 21,000 and contains one binding site for one moleculeof retinol. Intercellular and intracellular binding proteins belong to aclosely related family of retinoid-binding proteins of low molecularweight (Table 2.1).
Retinol is secreted from liver in association with RBP and circulates
to peripheral tissues complexed to a thyroxine-binding protein,transthyretin (Blomhoff et al., 1991; Ross, 1993). The retinoltransthyretin complex is transported to target tissues, where the complex binds to a cell-surface receptor and the retinol is transported into
cells of target tissue. Metabolism, storage, and release of vitamin A by
the liver are under several forms of homeostatic control, with circulating RBP maintained over a wide range of total liver reserves (Blomhoff
et al., 1991; Ross, 1993). One factor that specifically regulates RBP
secretion from liver is the nutritional vitamin A status of the animal. The
secretion of RBP from the liver is regulated by estrogen and nutritional
status of vitamin A, protein, and zinc. Retinol deficiency specifically
blocks secretion of RBP from the liver so that plasma RBP levels fall andliver RBP levels rise.
Contrary to current knowledge for all other species, vitamin A
serum levels in dogs were affected by the daily vitamin A supply as wellas the type of food (Schweigert et al., 1990). This was explained by thefact that dogs and other carnivores transport most of their vitamin A inblood as retinyl esters (mainly retinyl stearate) bound to lipoproteins.
The differences in the retinyl ester pattern between vitamin A in liverand in blood indicate that the mobilization of vitamin A from the liver
involves a selective mechanism.
Once the retinoids are transferred into the cell, they are quickly
bound by specific binding proteins in the cell cytosol. The intracellularretinoid-binding proteins bind retinol, retinal, and retinoic acid for pur25

CHAPTER TWO■ Table 2.1

Principal Retinoid-Binding Proteins

Name

Abbreviation

Retinoid

Principal Location

INTERCELLULAR TRANSPORT

Chylomicron
retinol-binding protein
Interphotoreceptor
retinol-binding protein

RBP
IRBP

Retinyl esters
Retinol

Intestine, liver
Liver, extrahepaticorgansAll-trans-retinol/ll-cis-retinol

RetinaINTRACELLULAR TRANSPORT

Cellular
CRBP (I)retinol-binding protein (I)Retinol

Most tissues except
adrenal, heart, ileum,muscle, serum

Cellular retinoic
acid-binding protein (I)CRABP (I)Retinoic acid

Seminal vesicle, vasdeferens, skin, eye

Cellular retinoic
acid-binding protein (II)CRABP (II)Retinoic acid

Skin

Epididymal retinoic acid

EBP 1,2Retinoic acid

Lumen of epididymis

Cellular retinaldehydebinding protein

CRALBPll-cis-retinal/
ll-cis-retinol

RetinaNuclear retinoic acidreceptors

RAR α, β, γAll-trans-retinoic
acid

Most tissues except
adult liver

Nuclear retinoic acid X
receptors

RXR α, β, γ9-cis-retinoic acid

Most tissues

poses of protection against decomposition, solubilize them in aqueous
medium, render them nontoxic, and transport them within cells to theirsite of action. These binding proteins also function by presenting theretinoids to the appropriate enzymes for metabolism (Wolf, 1991). Someof the principal forms of intracellular (cytoplasmic) retinoid-binding
proteins (Table 2.1) are cellular retinol-binding proteins (CRBP I and II),
cellular retinoic acid-binding proteins (CRABP I and II), cellular retinaldehyde-binding protein (CRALBP), and six nuclear retinoic acidreceptors (RAR α, β, γ and RXR α, β, γ).
The cellular retinol and retinoic acid-binding proteins—CRBP (I, II)
and CRABP (I, II)—function in transport and metabolism of retinoids
within parenchymal, intestinal, reproductive, and fetal cells and acrossblood-organ barriers. A different group of retinoid-binding proteins,more related to serum retinol-binding protein, functions in epididymis
26

V I TA M I N A

and uterus. Retinaldehyde-binding protein aids in the oxidation-reduction reaction of 11-cis-retinol–11-cis-retinaldehyde in the retina, where
the interphotoreceptor retinol-binding protein transports retinol
between pigment epithelium and photoreceptors (Wolf, 1998). Finally,
there are two classes of nuclear receptors: all-trans-retinoic acid is theligand for RAR, and 9-cis-retinoic acid is the ligand for RXR.
Receptors for 1,25-dihydroxyvitamin D3, all-trans retinoic acid, and9-cis-retinoic acid are members of the nuclear hormone receptor superfamily, a gene family having at least 30 members, including receptors forthe classic steroid hormones (estrogen, progesterone, glucocorticoids,
androgens, thyroid hormone, and several others). The feature common
to this class of molecules is that they control gene expression by interacting with specific DNA sequences or regulatory elements in control
regions of target genes.
Retinol is readily transferred to the egg in birds, but the transfer ofretinol across the placenta is marginal, and mammals are born with very
low liver stores of vitamin A. Uterine RBP has been identified in the pig
uterus, with the function of delivering retinol to the fetus (Clawitter etal., 1990).

Excretion
Derivatives of vitamin A with an intact carbon chain are generally
excreted in feces, whereas acidic chain-shortened products tend to be
excreted in urine (Olson, 1991). In the steady state, approximately equalamounts of metabolites are excreted in the feces and urine.
Some vitamin A derivatives are reexcreted into the intestinal lumen
via the bile. This is true for much of the retinoic acid and some retinol.
The major vitamin A components of the bile are vitamin A glucuronides.
An appreciable portion of these glucuronides are reabsorbed, thus creating an enterohepatic circulation for vitamin A derivatives and providing an opportunity for vitamin A conservation (Barua, 1997). Kinetic
studies have demonstrated that the retinol molecule cycles several times
between liver and extrahepatic tissues before it is degraded (Blomhoff etal., 1991).

StorageLiver normally contains about 90% of total-body vitamin A. The
remainder is stored in the kidneys, lungs, adrenals, and blood, withsmall amounts also found in other organs and tissues. A large quantityof vitamin A is stored in the kidney as well as the liver in cats and dogs.
This high level of vitamin A in the kidney is unique to cats and dogs, and27

CHAPTER TWO

the reasons for the storage are not fully understood (Ralston Purina,
1987; Raila et al., 1997). Vitamin A is highly concentrated in stomach
oils of certain seabirds and in the intestinal wall of some fish. The entire
vitamin A reserve of certain shrimp is in the eyes. Carotenoids are more
evenly distributed in species that have the ability to absorb and store
these precursors. Grass-fed cattle have large stores of carotene in theirbody fat, which is evidenced by a deep yellow color.
The liver can store large amounts of vitamin A; in humans approximately 50 to 80% of the total body vitamin A is stored in the stellate
cells in the form of retinyl esters (Blomhoff et al., 1991). Bardos (1991)
compared vitamin A content in liver lobes between dogs and otherspecies. Uniformity of liver vitamin A was less for dogs than cattle andchickens. In all species, there was a low correlation between liver andblood retinol. Several studies have shown that liver can store enoughvitamin A to protect the animal from long periods of dietary scarcity(McDowell, 1985). This large storage capacity must be considered instudies of vitamin requirements to ensure that intakes that appear adequate for a given function are not being supplemented by reserves storedprior to the period of observation. Measurement of the liver store ofvitamin A at slaughter or in samples obtained from a biopsy is a usefultechnique in the study of vitamin A status and requirements. Fig. 2.4
illustrates a liver biopsy procedure that was adapted from human medicine.

FUNCTIONS
Vitamin A is necessary for support of growth, health, and life of
higher animals. In the absence of vitamin A, animals will cease to growand eventually die. The classic biological assay method is based on
measurement of growth responses of weanling rats to graded doses ofvitamin A. It is of primary importance in development of young, growing animals.
The metabolic function of vitamin A, explained in biochemicalterms, is only now beginning to be understood. Vitamin A deficiency
causes at least four different and probably physiologically distinct
lesions: loss of vision due to a failure of rhodopsin formation in the retina; defects in bone growth; defects in reproduction (e.g., failure of spermatogenesis in the male and resorption of the fetus in the female); anddefects in growth and differentiation of epithelial tissues, frequently
resulting in keratinization. Keratinization of these tissues results in lossof function; this occurs in the alimentary, genital, reproductive, respira28

A

B

C
Fig. 2.4 Illustration of liver biopsy sample taken for both vitamin A and mineral analysis. Sample can be taken quickly with a trocar and cannula: (A)Trocar; (B) Insertion of trocar for sampling; (C) Release of sample from thecannula. (Courtesy of H.L. Chapman and L.R. McDowell, University of29

CHAPTER TWO

tory, and urinary tracts. Such altered characteristics make the affected
tissue more susceptible to infections. Thus, diarrhea and pneumonia are
typical secondary effects of vitamin A deficiency. Retinoic acid is theform of vitamin A that has been shown to perform as a hormone (e.g.,
all-trans-retinoic acid and 9-cis-retinoic acid). However, retinoic acidhas been found to support growth and tissue differentiation but notvision (Fig. 2.5) or reproduction (Scott et al., 1982). Vitamin A-deficient
rats fed retinoic acid were healthy in every respect, with normal estrus
and conception, but failed to give birth and resorbed their fetuses. When
retinol was given even at a late stage of pregnancy, fetuses were saved.
Male rats on retinoic acid were healthy but produced no sperm, andwithout vitamin A, both sexes were blind (Anonymous, 1977).
Although retinol is needed for normal vision and some aspects ofreproduction, discoveries have revealed that most, if not all, actions ofvitamin A in development, differentiation, and metabolism are mediated by nuclear receptor proteins that bind retinoic acid, the active formof vitamin A (Anonymous, 1993). A group of retinoic acid binding proteins (receptors) function in the nucleus by attaching to promoter
regions in a number of specific genes to stimulate their transcription andthus affect growth, development, and differentiation. Six high-affinity
receptor proteins for retinoic acid (RAR α, β, γ and RXR α, β, γ) havebeen identified. Apparently RAR nuclear receptors bind to all-transretinoic acid, while RXR receptors bind with 9-cis-retinoic acid (Kasner
et al., 1994; Kliewer et al., 1994). Retinoic acid receptors in cell nuclei
are structurally homologous and functionally analogous to the known
receptors for steroid hormones, thyroid hormone (triiodothyronine),and vitamin D [1,25-(OH)2D]. Thus, retinoic acid is now recognized tofunction as a hormone to regulate the transcription activity of a large
number of genes (Ross, 1993). As an example, the well-known connection between vitamin A deficiency and hepatic glycogen depletioncaused by reduced gluconeogenesis can now be explained at the molecular level by the dependence of phosphoenolpyruvate carboxykinase
gene expression on adequate vitamin A (Shin and McGrane, 1997).
Actions of vitamin A in development, differentiation, and metabolism are mediated by nuclear receptor proteins (RARs and RXRs) thatbind retinoic acid with steroid and thyroid hormone receptors. The
super family of nuclear proteins interact with specific genes and regulate
their transcription. Retinoic acid has been found to stimulate, synergistically with thyroid hormone, the production of growth hormone in cultured pituitary cells. The RARs have been found to bind both the geneelement responsive to RAR, in addition to the one responsive to tri30

V I TA M I N A

ABFig. 2.5 The appearance of (A) the eye of a blind hen fed retinoic acid compared with (B) the normal eye of a hen fed retinol. (Courtesy of M.L. Scott,Cornell University.)

iodothyronine, suggesting that retinoic acid and the thyroid hormone
control overlapping networks of genes. Many proteins appear during
retinoic acid-induced cell differentiation.
Retinoids have a wide spectrum of biological activities. Retinoic
acid plays an important role in growth and differentiation of embryonic tissues. It also regulates the differentiation of epithelial, connective,
and hematopoietic tissues (Safonova et al., 1994). The nature of thegrowth and differentiation response elicited by retinoic acid depends
upon cell type. Retinoic acid can be an inhibitor of many cell types witha potential to reduce adipose tissues in meat-producing animals(Suryawan and Hu, 1997).
Evidence indicates a morphogenic role for retinoic acid and one ofits metabolites, 3,4-didehydroretinoic acid (Anonymous, 1991).
Morphogens form concentration gradients or morphogenic fields
through developing tissues that specify the eventual three-dimensional
structure at maturity. Cell differentiation in the developing chick limb
bud has been studied where vitamin A morphogens are operative. Cellsof the limb bud can differentiate into muscle, cartilage, and bone cells.

VisionThe physiological function of vitamin A that has been most clearly
defined on a biochemical basis is its role in vision (Wald, 1968). Vitamin
A is an essential component of vision. Retinol is utilized in the aldehyde
form (trans- form to 11-cis-retinal) in the retina of the eye as the prosthetic group in rhodopsin for dim light vision (rods) and as the prosthetic group in iodopsin for bright light and color vision (cones). When
11-cis-retinal (aldehyde form of vitamin A) is combined with the protein
31

CHAPTER TWO

opsin, rhodopsin (visual purple) is produced. Chemical reactions
involved in vision and the roles that trans-retinal and 11-cis-retinal playin this function are presented in Fig. 2.6. Rhodopsin breaks down in thephysiological process of sight as a result of photochemical reaction. The
all-trans- retinaldehyde cannot form a stable complex with the opsin.
Opsin opens through a series of changes that expose reactive groups.
Finally, retinaldehyde is hydrolyzed off the opsin. The energy derived
from this reaction is transported to the brain via the optic nerve andrecorded in various intensities depending on the amount of light entering the eye.
Vitamin A deficiency, in terms of the need for the resynthesis ofrhodopsin, results in night blindness (nyctalopia), which is a clinical signin both animals and humans. During the reactions in the retina, some ofthe vitamin A is lost and is replaced by vitamin A from blood. If vitamin
A blood level is too low, a functional night blindness will result. The
deficiency first manifests itself as a slow adaptation to the dark, and progresses to total night blindness. At dusk or in moonlight, livestock withnight blindness will bump into obstacles (Fig. 2.7) intentionally placedin their path, or into logs or stumps when driven at night.
In vitamin A deficiency, the outer segments of the rods lose theirFig. 2.6 The role of vitamin A in vision. (Adapted from Wald, 1968.)
32

V I TA M I N A

Fig. 2.7 A blind steer with vitamin A deficiency walks into a fence. (Courtesyof T.B. Keith, University of Idaho.)

opsin, leading to their eventual degeneration. The entire structure
becomes filled with tubules and vesicles. On a molecular basis, collagenase activity is increased in vitamin A deficiency. Retinoic acid has beenshown to inhibit the enzyme collagenase by forming an inactive protein
complex with the liganded nuclear retinoic acid receptors (Wolf, 1992).
Even at a late stage, it is possible to regenerate rods, but continued deficiency results in disintegration of cones and total blindness. Vitamin A
is needed for integrity of the visual cells as well as their normal regeneration.
Other eye clinical signs vary markedly among species, some ofwhich represent secondary infections. Vision also can be impaired inxerophthalmia. Xerophthalmia is an advanced stage of vitamin A deficiency seen in all species. In children, dogs, foxes, and rats, xerophthalmia is characterized by a dry condition of the cornea and conjunctiva, cloudiness, and ulceration. Copious lacrimation is a more prominenteye sign in cows (Fig. 2.8) and horses. In chickens, on the other hand,the secretions of the tear glands dry up and an infection may then occur,
resulting in a discharge that causes the lids to stick together. Some ofthese conditions develop as a result of basic epithelial changes caused bya deficiency of vitamin A (Maynard et al., 1979).
33

CHAPTER TWO

Fig. 2.8 Calf in the Philippines shows vitamin A deficiency characterized bycopious lacrimation and blindness. The 6-month-old animal had been fed
reconstituted skim milk powder and poor-quality bleached hay (practicallydevoid of carotene). (Courtesy of J.K. Loosli, University of Florida.)

Maintenance of Normal Epithelium
Vitamin A is required for maintenance of epithelial cells, which
form protective linings on many of the body’s organs. The respiratory,
gastrointestinal, and urogenital tracts, as well as the eye, are protectedfrom environmental influences by mucous membranes. If, however,there is a deficiency of vitamin A, epithelial cells that make up the membrane will change their characteristic structure. It is postulated thatvitamin A plays an important role in altering permeability of lipoprotein
membranes of cells and of intracellular particles. Vitamin A penetrates
lipoprotein membranes and, at optimum levels, may act as a cross-linkage agent between the lipid and protein, thus stabilizing the membrane(Scott et al., 1982).
The normal mucus-secreting cells of epithelium in various locations
throughout the body become replaced by a stratified, keratinized epithelium when vitamin A is deficient. Keratinized epithelium allows
pathogen entry through the skin, lung, gastrointestinal tract, and uro-34

V I TA M I N A

genital tract surface. Vitamin A deficiency can impair regeneration of
normal mucosal epithelium damaged by infection or inflammation(Ahmed et al., 1990; Stephensen et al., 1993) and thus could increase theseverity of an infectious episode and/or prolong recovery from that
episode. Adequate dietary vitamin A is necessary to help maintain normal resistance to stress and disease.
Studies have shown that the epithelial cells from vitamin A-deficientanimals fail to differentiate to mucus-secreting cells, and mesenchymal
cells fail to differentiate beyond the blast stage. This occurs in the alimentary, genital, reproductive, respiratory, and urinary tracts. Suchaltered characteristics make affected tissues more susceptible to infection. Thus, colds and pneumonia are typical secondary effects of vitamin
A deficiency. Adequate dietary vitamin A is necessary to help maintainnormal resistance to stress and disease. However, greater than optimal
intakes of vitamin A will not aid in preventing infections.
There are many noninfective problems due to keratinization ofepithelium, such as diarrhea. The formation of kidney and bladderstones is favored when damaged epithelium interferes with normal
secretion and elimination of urine, and sloughed keratinized cells may
form foci for the formation of stones. There is a specific interferencewith reproduction caused by altered epithelium that is of great importance. Squamous metaplasia in the parotid gland is an early change invitamin A-deficient calves and proves useful in diagnosing deficiency.
Elevated cerebrospinal fluid pressure observed in vitamin A-deficientanimals, a very sensitive measure of the onset of vitamin A deficiency, isthe result of cell changes. Increased ground substance in the dura matersurrounding the arachnoid villus and altered epithelial cells cause adecreased absorption of the fluid.
There is evidence that vitamin A is necessary for the formation oflarge molecules containing glucosamine (Goodman, 1980). These are
the mucopolysaccharides occurring in almost all tissues of mammalianorganisms but principally in the mucus-secreting epithelia and in theextracellular matrix of cartilage, mainly as chondroitin sulfate. The intimate involvement of vitamin A in the biosynthesis of glycoproteins,which are constitutents of membrane systems in cells, helps explain
many biological effects of this vitamin.
In severe vitamin A deficiency, abnormalities in both RNA metabolism and protein synthesis have been reported. These changes in nucleic
acid metabolism and protein synthesis may, however, reflect secondary
effects of deficiency rather than the primary function of the vitamin.

35

CHAPTER TWO

Reproduction
In most livestock, the absence of vitamin A in the ration will dramatically reduce reproductive ability. Hatchability is significantly
reduced when hens are fed a vitamin A-deficient ration. One of the firstsigns of vitamin A deficiency in rabbits is a reduction in fertility and anincreased incidence of abortion in pregnant does.
In a number of species, vitamin A deficiency in the male results in a
decline in sexual activity and failure of spermatogenesis, and in thefemale results in the resorption of the fetus, abortion, or birth of deadoffspring. Retained placenta may be a characteristic of vitamin A deficiency in some species. Often the reproductive problems associated withvitamin A deficiency are actually the result of failure to maintain healthyepithelium.
Degeneration of germinal epithelium and seminiferous tubules andcessation of spermatogenesis in vitamin A-deficient rats are not prevented by retinoic acid. However, vitamin A-deficient rats had lowered
testosterone levels that were restored by retinoic acid. Since retinoic acidcannot restore the germinal epithelium in vitamin A-deficient rats, theseresults indicate a role for retinoic acid in testosterone synthesis(Anonymous, 1977).

Bone Development
Vitamin A has a role in the normal development of bone throughcontrol exercised over the activity of osteoclasts of the epithelial cartilage. In vitamin A deficiency, osteoclast (reabsorbing bone) activity is
reduced, resulting in excessive deposition of periosteal bone by theapparently unchecked function of osteoblasts (depositing bone).
Disorganized bone growth and irritation of the joints are two manifestations of vitamin A deficiencies. In some cases, there is a constriction of
the openings through which the optic and auditory nerves pass, thereby
resulting in blindness and/or deafness.
Bone changes may also be responsible for the muscle incoordinationand other nervous symptoms shown by vitamin A-deficient cattle, sheep,and swine. These changes may be involved in the increase in cerebrospinal fluid pressure shown to be characteristic of the deficiency.
While the pathological basis is unknown, several studies have shownthat a lack of vitamin A causes congenital malformation in certain soft
tissues (Maynard et al., 1979). Examples are the birth of pigs without
eyeballs and hydrocephalus in rabbits.

36

V I TA M I N A

Relationship to Immunological Response and Disease
Conditions
Animals deficient in vitamin A will show increased frequency andseverity of bacterial, protozoal, and viral infections as well as other disease conditions. Part of disease resistance, as a function of vitamin A, isrelated to maintenance of mucous membranes and normal functioningof the adrenal gland for production of corticosteroids needed to combat
disease. An animal’s ability to resist disease depends on a responsive
immune system, with a vitamin A deficiency causing a reduced immune
response (Semba, 1998).
Vitamin A deficiency affects immune function, particularly the antibody response to T-cell–dependent antigens (Ross, 1992). The RAR-αmRNA expression and antigen-specific proliferative responses of T lymphocytes are influenced by vitamin A status in vivo and are directly
modulated by retinoic acid (Halevy et al., 1994). Vitamin A deficiency
affects a number of cells of the immune system, and that repletion withretinoic acid effectively reestablishes the number of circulating lymphocytes (Zhao and Ross, 1995).
A diminished primary antibody response could also increase theseverity and/or duration of an episode of infection, whereas a diminished secondary response could increase the risk of developing a second
episode of infection. Vitamin A deficiency causes decreased phagocytic
activity in macrophages and neutrophils. The secretory immunoglobulin(Ig) A system is an important first line of defense against infections ofmucosal surfaces (McGhee et al., 1992). Several studies in animal models have shown that the intestinal IgA response is impaired by vitamin A
deficiency (Davis and Sell, 1989; Wiedermann et al., 1993; Stephensen
et al., 1996).
An optimal vitamin A range exists for enhancement of vitamin A
responses, because both deficient status and excessive status suppress
immune function. In many experiments with laboratory and domestic
animals, the effects of both clinical and subclinical deficiencies of vitamin A on the production of antibodies and on the resistance of the different tissues against microbial infection or parasitic infestation havefrequently been demonstrated (Kelley and Easter, 1987). Supplemental
vitamin A improved the health of animals infected with roundworms, ofhens infected with Capillaria organisms, and of rats with hookworms(Herrick, 1972). Vitamin A is valuable in treating ringworm(Trichophyton verrucosum) infection in cattle.
Vitamin A-deficient chicks showed rapid loss of lymphocytes, and37

CHAPTER TWO

deficient rats showed atrophy of the thymus and spleen and reduced
response to diphtheria and tetanus toxoids (Krishnan et al., 1974).
Mortality from fowl typhoid (Salmonella gallinarum) was reduced inchicks fed vitamin A levels greater than the normal levels in a high-protein diet. Serum antibody levels in chicks were increased twofold to fivefold by high dietary vitamin A concentrations. The immune response to
introduced Newcastle disease virus was higher in broiler chicks receiving 2,500 IU of vitamin A than in controls, and was best in those receiving 20,000 IU (Serman and Mazija, 1985). Harmon et al. (1963) studied the effect of a vitamin A deficiency on antibody production by baby
pigs and found a high correlation coefficient between serum vitamin A
and antibody titer. Baby pigs infected with Trichuris suis responded tosupplemental vitamin A with an enhanced immunological response compared with controls (Bebravicius et al., 1987).
Vitamin A and β−carotene have important roles in protecting animals against numerous infections, including mastitis. A protective effectof dietary vitamin A supplementation against experimental
Staphylococcus aureus mastitis in mice has been reported (Chew et al.,1984). Potential pathogens are regularly present in the teat orifice, andunder suitable circumstances can invade and initiate clinical mastitis.
Any unhealthy state of the epithelium would increase susceptibility of a
mammary gland to invasion by pathogens. There are reports ofimproved mammary health in dairy cows supplemented with β−carotene
and vitamin A during the dry (Dahlquist and Chew, 1985) and lactating(Chew and Johnston, 1985) periods.
In vitamin A-deficient animals, incidence of cancer has been shown
to be higher than in animals receiving normal vitamin A intakes. In somespecial cases, administration of toxic doses of vitamin A caused a regression of tumors. Extreme toxicity of natural vitamin A makes it an ineffective drug for treatment of certain types of neoplasms. Synthetic
analogs (retinoids) of vitamin A, however, have been successfully used toprevent cancer of the skin, lung, bladder, and breast in experimental animals. This is a pharmacological approach to prevention of cancer byenhancement of intrinsic epithelial defense mechanisms. Synthetic
retinoids are superior for this purpose (Goodman, 1980). In humans,studies suggest that diets rich in β−carotene provide a blocking or inhibition of certain types of cancer, including lung cancer (even in people
who smoke). Also, skin diseases, including psoriasis, cystic acne, androsacea have responded exceptionally well to synthetic retinoids.

38

V I TA M I N A

REQUIREMENTS
Extensive research has been conducted to determine the vitamin A
requirements of various species. Requirements have been published in
the United States by the Committee on Animal Nutrition of the National
Academy of Science-National Research Council. Vitamin A requirements can be expressed on the basis of IU per kilogram of body weight,
on a daily basis, or as a unit of diet. In agreement with general practice,
the requirements are normally expressed per unit of diet rather than per
kilogram of body weight.
Minimum requirements have been determined by various methods,
including amounts required to prevent night blindness, amounts
required for storage and reproduction, and maintenance of normal pressure in the cerebrospinal fluid. The minimum vitamin A requirement fornormal growth may be lower than the amount required for higher
growth rates, resistance to various diseases, and normal bone development. It was suggested that calves born with low vitamin A liver storesshould receive a minimum of 16,500 IU/100 kg body weight, and thatthree to five times this level are necessary for adequate vitamin A liver
stores in calves during the critical first few months of life.
Table 2.2 summarizes the vitamin A requirements for various
species; a more complete listing is provided in Appendix Table A1a.
These requirements are deemed sufficient to provide optimal growth;satisfactory reproduction and milk, egg, or wool production; and prevention of deficiency signs. The requirements presented are designed tobe adequate for these purposes under practical conditions of feeding andmanagement and to allow for a certain amount of storage. The decision
as to the minimum vitamin A requirement of young swine depends upon
whether the criterion to determine the requirements is based upon
growth and feed utilization or also considers liver vitamin A storage.
Storage of vitamin A in the liver certainly appears to be desirable since
under conditions of little or no liver storage, stresses and diseases may
precipitate vitamin A deficiency. The vitamin A reserves of the sow makeit difficult to establish requirements. Braude et al. (1941) reported thatmature sows fed diets without supplemental vitamin A completed threepregnancies normally; only in the fourth pregnancy did deficiency signs
appear. Gilts receiving adequate vitamin A levels until 9 months of age
completed two reproductive cycles without signs of vitamin A deficiency (Selke et al., 1967).

39

CHAPTER TWO■ Table 2.2

Vitamin A Requirements for Various Animals and for Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle

Feedlot cattle
Pregnant heifers and cows
Lactating cows and bulls
Growing
Lactating cows and bulls
Calf milk replacer
All classes

2,200 IU/kg
2,800 IU/kg
3,900 IU/kg
2,200 IU/kg
3,200 IU/kg
3,800 IU/kg
5,000 IU/kg
1,500 IU/kg
2,000 IU/kg
1,500 IU/kg
1,500 IU/kg
4,000 IU/kg
5,000 IU/kg
1,567 IU/kg
3,306 IU/kg
2,380 IU/kg
1,976 IU/kg
2,200 IU/kg
1,300 IU/kg
4,000 IU/kg
2,000 IU/kg
2,000 IU/kg

NRC (1996)
NRC (1996)
NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1989a)
Morand-Fehr(1981)NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1985b)
NRC (1985b)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)

3,000 IU/kg
5,930 IU/kg
2,440 IU/kg
6,000 IU/kg
3,336 IU/kg
580 IU/kg
1,160 IU/kg
1,000–2,000 IU/kg
1,000–2,000 IU/kg
2,500 IU/kg
2,300 IU/kg
2,400 IU/kg
10,000 IU/kg
400–700 μg REb
800–1,000 μg/RE
1,200–1,300 μg/RE

NRC (1989b)
NRC (1982a)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1977)
NRC (1977)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
NRC (1978)
RDA (1989)
RDA (1989)
RDA (1989)

Dairy cattle
Goat
Chicken

Leghorn, 0–18 weeks
Laying (100-g intake)
Broilers
Geese
Growing
Breeding
TurkeyGrowing and breeding
Sheep
Replacement ewes, 60 kg
Pregnancy, 70 kg
Lactation, 70 kg
Replacement rams, 80–100 kg
Swine
Growing 5–10 kg
Growing, 20–120 kg
Pregnant swine and boars
LactatingHorse
Growing, maintenanceand working
Pregnancy and lactation
Mink
Growing
Fox
GrowingCat
Gestation
Dog
Growing
RabbitGrowing
Gestation
Fish
CatfishCommon carp
Rainbow trout
Rat
Growing and reproduction
Mouse
Growing
Nonhuman primate All classes
HumanChildren
AdultsLactatingaExpressed as per unit of animal feed either on as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as μg/day.
bRetinol equivalent (RE): 1 RE = 1 μg retinol or 6 μg β-carotene.

In establishing a satisfactory vitamin A level for practical diets, it isnecessary to consider a number of factors that may alter the vitamin A
requirement. Type and level of production are important as greater production rates increase requirements. Pregnancy, lactation, and egg production also lead to higher requirements. Other practical factors are listed in Table 2.3.
40

V I TA M I N A■ Table 2.3

Factors Influencing Vitamin A Requirements

Genetic differences (species, breed, strain)
Carryover effect of stored vitamin A (principally in the liver but the kidney is also important for cats)
Conversion efficiency of carotenes to vitamin A
Variations in level, type, and isomerization of carotenoid vitamin A precursors in feedstuffs
Presence of adequate bile in vivo
Destruction of vitamin A in feeds through oxidation, prolonged storage, high pelleting
temperatures, catalytic effects of trace minerals, and peroxidizing effects of rancid fats
Presence of disease and/or parasites
Environmental stress and temperature
Adequacy of dietary fat, protein, zinc, phosphorus, and antioxidants (including vitamin E,
vitamin C, and selenium)
Pelleting and subsequent storage of feed

Different species of animals convert β−carotene to vitamin A withvarying degrees of efficiency. The conversion rate of the rat has beenused as the standard value, with 1 μg of β−carotene equal to 1,667 IU
of vitamin A. The comparative efficiencies of various species are shownin Table 2.4, based on this standard. Of the species studied, only poultry are equal to rats in vitamin conversion; cattle are only 24% as efficient.
Some factors that influence the rate at which carotenoids are converted to vitamin A are type of carotenoid, class and production level ofanimal, individual genetic differences in animals, and level of carotene
intake (NRC, 1996). Efficiency of vitamin A conversion from β−carotene is decreased with higher levels of intake (Van Vliet et al., 1996)(Table 2.5). As β−carotene level is increased, conversion efficiency drops
from a ratio of 2:1 to 5:1 for the chicken and from 8:1 to 16:1 for thecalf (Bauernfeind, 1972).
Stress conditions, such as extremely hot weather, viral infection, andaltered thyroid function, have also been suggested as causes for reduced
conversion efficiency of carotene to vitamin A. Vitamin A requirements
are higher under stressful conditions such as abnormal temperatures or
exposure to disease. For example, with poultry, coccidiosis not only
causes destruction of vitamin A in the gut but also injures the microvilli of the intestinal wall, thereby decreasing absorption of vitamin A andat the same time causing the chickens to stop eating for several days
(Scott et al., 1982).
41

CHAPTER TWO■ Table 2.4

AnimalStandard
Beef cattle
Dairy cattle
Sheep
Swine
Horse
Growing
Pregnant
Poultry
Dog
Rat
Fox
Cat
Mink
Human

Conversion of β-Carotene to Vitamin A by Different Animals
Conversion of mg ofβ-Carotene to IU of
Vitamin A (mg) (IU)
1 = 1,667
1 = 400
1 = 400
1 = 400–450
1 = 500
1 = 555
1 = 333
1 = 1,667
1 = 833
1 = 1,667
1 = 278
Carotene not utilized
Carotene not utilized
1 = 556

IU of vitamin A Activity(Calculated from Carotene)
(%)
100
24
24
24–30
30
33.3
20
100
50
100
16.7
—
—
33.3

Sources: Adapted from Beeson (1965) and United States-Canadian Tables of Feed
Composition (NRC, 1982b).

Likewise, other factors may affect metabolism and increase vitamin
A requirements. These include free nitrates in feeds, inadequate protein,zinc deficiency, and low dietary phosphorus (Harris, 1975).
Considerable work and controversy have been reported on the relationship between nitrates and vitamin A nutrition. In a review of this subject, Rumsey (1975) concluded that although nitrates can be shown tohave an adverse effect on vitamin A in vitro, this does not appear totranslate into a significant effect under most feeding conditions.
The efficiency of β−carotene in meeting the vitamin A requirementof trout and salmon apparently is dependent on water temperature.
Cold-water fish utilize precursors of vitamin A at 12.4 to 14oC but not
at 9oC (Poston et al., 1977). Activity of β−carotene-15,15´-dioxygenase,which oxidizes β−carotene to retinal in the intestinal mucosa, may be
restricted at cold temperatures.
In humans the recommended allowance for adult females is set at
80% of that for males or 800 retinol equivalents (4,000 IU) (RDA,
1989). The allowance during lactation is increased to 1,200 retinol
equivalents to provide for vitamin A secreted in milk. Daily vitamin A
requirements for children (10 years or younger) vary between 400 and
700 μg retinol equivalents.

42

V I TA M I N A■ Table 2.5

Equivalence of β-Carotene and Retinol at Different Dietary Concentrationsin Rats

Dose ofβ-Carotene(μmol/kg BW)

Relative
Molar
Biopotency (%)

Moles
β-Carotene
Equivalent to 1
mol Retinol

μg β-Carotene
Equivalent to 1
mol Retinol

< 0.3
1
2
6
12
48

100.0
50.0
30.0
15.4
9.5
4.3

1.0
2.0
3.3
6.5
10.5
23.3

1.84
3.75
6.25
12.20
19.70
43.60

Source: Adapted from Brubacher and Weiser (1985).

NATURAL SOURCES
The richest sources of vitamin A are fish oils. Some swordfish liver
oils contain as many as 250,000 IU of vitamin A per gram. Halibut liver
oil may contain even more. Thus, both are many times more potent thancod liver oils. Products from the same species, however, may be highlyvariable in potency; thus, in their manufacture for use as a vitamin A
supplement, they are subjected to a biological assay so that the user may
be assured of a certain minimum potency. Among the common foods ofanimal origin, milk fat, egg yolk, and liver are rated as rich sources, but
this is not the case if the animal from which they came has been receiving a vitamin A-deficient diet for an extended period. Since the vitaminis present in the fat, skim milk contains very little. The effect of dietary
vitamin A on egg yolk concentration was reported by Hill (1961).
Vitamin A concentrations of 1,760, 4,400, and 22,000 IU/kg feed resulted in vitamin A yolk levels of 0.9, 6.3, and 16.3 IU/g, respectively.
Sources of supplemental vitamin A are derived primarily from fish
liver oils, in which the vitamin occurs largely in esterified form, andfrom industrial chemical synthesis. Before the era of the chemical production of vitamin A, the principal source of vitamin A concentrates was
the liver and/or body oils of marine fish. Since industrial synthesis wasdeveloped in 1949, the synthetic form has become the major source ofthe vitamin to meet the requirements of domestic animals and humans.
The synthetic vitamin usually is produced as the all-trans-retinyl palmitate or acetate.
Provitamin A carotenoids, mainly β−carotene in green feeds, are the43CHAPTER TWO

principal source of vitamin A for grazing livestock. All green parts ofgrowing plants are rich in carotene and therefore have a high vitamin A
value. In fact, the degree of green color in a roughage is a good index ofits carotene content. Although the yellow color of carotenoids is masked
by chlorophyll, all green parts of growing plants are rich in carotene andthus have a high vitamin A value. Good pasture always provides a liberal supply, and type of pasture plant—whether grass or legume—
appears to be of minor importance. At maturity, however, leaves containmuch more than stems, and thus legume hay is richer in vitamin content
than grass hay (Maynard et al., 1979). With all hays and other forage,
vitamin A value decreases after the bloom stage. Plants at maturity canhave 50% or less of the maximum carotenoid value of immature plants.
Both carotene and vitamin A are destroyed by oxidation, and this is
the most common cause of any depreciation that may occur in thepotency of sources. The process is accelerated at high temperatures, butheat without oxygen has a minor effect. Butter exposed in thin layers in
air at 50oC loses all its vitamin A potency in 6 hours, but in the absenceof air there is little destruction at 120oC over the same period. Cod liver
oil in a tightly corked bottle has shown activity after 31 years, but it may
lose all its potency in a few weeks when incorporated in a feed mixture
stored under usual conditions (Maynard et al., 1979).
Much of carotene content is destroyed by oxidation in the processof field curing. Russell (1929) found that there may be a loss of morethan 80% of the carotene in alfalfa during the first 24 hours of the curing process. This loss occurs chiefly during daylight hours, partly
because of photochemical activation of the destructive process. In alfalfa leaves, sunlight-sensitized destruction comprises 7 to 8% of the total
pigment destroyed, while enzymatic destruction comprises 27 to 28%(Bauernfeind, 1972). Enzymatic destruction requires oxygen, is greatest
at high temperatures, and ceases after complete dehydration.
Hays that are cut in the bloom stage or earlier and cured without
exposure to rain or too much sun retain a considerable proportion of
their carotene content, while those cut in the seed stage and exposed torain and sun for extended periods lose most of it. Green hay curing inthe swath may lose half its vitamin A activity in one day’s exposure to
sunlight and almost all of it if left exposed to rain as well as sunlight.
Thus, hay usually has only a small proportion of the carotene content offresh grass. Under similar conditions of curing, alfalfa and other legume
hays are much richer than grass hays because of their leafy nature, but
a poor grade of alfalfa may have less than a good grade of grass hay(Maynard et al., 1979).
44

V I TA M I N A

The carotene content of dried or sun-cured forages decreases in storage with the rate of destruction depending on factors such as temperature, exposure to air and sunlight, and length of storage. Under average
conditions, carotene content of hay can be expected to decrease by
about 6 to 7% per month. In artificial curing of hay with a “hay drier,”
there is only a slight loss of carotene because of the rapidity of theprocess and protection against exposure to oxygen, with the final product having 2 to10 times the value of field-cured hay. Severe heating of
hay in the mow or stack reduces vitamin content, and there is a gradualloss in storage, so old hay is poorer than new. Aside from yellow corn
and its by-products, practically all the concentrates used in feeding animals are devoid of vitamin A value, or nearly so. In addition, yellow
corn contains a high proportion of non-β−carotenoids (e.g., cryptoxanthin, lutein, and zeacarotene) that contain much less or no vitamin A
value compared to β−carotene.
The potency of yellow corn is only about one-eighth that of goodroughage. Roots and tubers as a class supply practically no vitamin A,
but carrots are a very rich source, as are sweet potatoes, as might be
expected from their yellow color. Pumpkins and squash also supply considerable amounts, and green leafy vegetables used in human nutrition
are rich in carotene (Maynard et al., 1979). Tankage, meat scraps, andsimilar animal by-products have little if any vitamin A potency. Certain
fish meals are fair sources, but variation in the raw material and inmethods of processing may entirely destroy any potency originally present.
There is evidence that yellow corn may lose carotene rapidly duringstorage. For instance, a hybrid corn high in carotene lost about half of
its carotene in 8 months of storage at 25oC and about three-quarters in
3 years. Less carotene was lost during storage at 7oC (Quackenbush,1963). Bioavailability of natural β−carotene is less than chemically synthesized forms (Hussein and El-Tohamy, 1990; White et al., 1993).
Using ferrets, all-trans β−carotene was less bioavailable from carrot juice
than from β−carotene beadlet-fortified beverages (White et al., 1993). In
calves, a small enhancing effect of mild heat treatment of carrots onserum and tissue accumulation of carotenoids has been reported (Pooret al., 1993).
In forages, carotene in alfalfa hay may be more bioavailable thanthat in grass hay (NRC, 1989b). A marked discrepancy exists between
the carotene content of corn silage and the vitamin A status of ruminants
fed corn silage. On the average, corn silage carotenes were found to beabout two-thirds as effective as β−carotene for maintaining liver stores
45

CHAPTER TWO

in rats (Miller et al., 1969; Rumsey, 1975). Martin et al. (1971) reported five times less carotenes in October and November corn silage thanin September corn silage. More mature silages were not able to sustain
liver vitamin A stores in beef steers, particularly if the ensiled corn plant
was finely chopped. Diets high in corn silage harvested after a killing
frost and fed to cattle would be marginal in their supply of both vitamins A and E. Miller et al. (1969) have reported that ethanol, sometimes
found in corn silage as a product of fermentation, may reduce liver vitamin A stores as much as 26% by increasing mobilization of vitamin A
from liver.
Wing (1969) reported carotene digestibility in plants to be greater
during the warmer months. Variations were found in the digestibility ofcarotenes in plants according to year, species of plant, dry matter content, and form of forage; carotene digestiblity was somewhat lower insilages than in pastures or hay. Table 2.6 presents typical vitamin A values of foods and carotene concentration of feeds. As noted earlier,degree of greenness in a roughage is a good index of its carotene content. The data in Table 2.6 are useful for indicating the order of the differences found among various roughages differing as to color, kind, andother factors. Average published values of carotene content can serve
only as approximate guides in feeding practice because of many factors
affecting actual potency of individual samples as fed (NRC, 1982b).
Cooking processes commonly used in human food preparation do
not cause much destruction to the vitamin potency. The blanching andfreezing process generally causes little loss of carotenoid content in vegetables and fruits. Heat, however, does isomerize the all-transcarotenoids to cis-forms. In a report from Indonesia, isomerization during traditional cooking caused a loss of up to 9% of vitamin A potency(Van der Pol et al., 1988). Hydrogenation of fats lessens their vitamin A
value, while saponification does not destroy the vitamin if oxidation is
avoided.
Several factors can influence the loss of vitamin A from feedstuffs
during storage. The trace minerals in feeds and supplements, particularly copper, are detrimental to vitamin A stability. Dash and Mitchell(1976) reported the vitamin A content of 1,293 commercial feeds over a
3-year period. The loss of vitamin A was over 50% in 1 year’s time.
Vitamin A loss in commercial feeds was evident even if the commercial
feeds contained stabilized vitamin A supplements. The stability of vitamin A in feeds and premixes has been improved tremendously in recent
years by chemical stabilization as an ester and by physical protection
using antioxidants, emulsifying agents, gelatin, and sugar in spray-dried,
46

V I TA M I N A■ Table 2.6

Vitamin A (Retinol) and β-Carotene Content of Feeds

Vitamin A Source
Whale liver oil
Swordfish liver oil
Halibut liver oil
Herring liver oil
Tuna liver oil
Shark liver oil
Bonito liver oil
White sea bass liver oil
Barracuda liver oil
Dogfish liver oil
Seal liver oil
Cod liver oil
Sardine body oil
Pilchard body oil
Menhaden body oil
ButterCheeseEggs
Milk

Vitamin A (IU/g)
400,000
250,000
240,000
211,000
150,000
150,000
120,000
50,00040,00012,00010,000
4,000
750
500
340
35
14
10
1.5

Carotene Source
Carotene (mg/kg)
Fresh green legumes and grasses, immature (wet basis)
33–88
Dehydrated alfalfa meal, fresh, dehydrated without field curing,very bright green
242–297
Dehydrated alfalfa meal after considerable time in storage, brightgreen
110–154
Alfalfa leaf meal, bright green
120–176
Legume hays, including alfalfa, very quickly cured with minimum
sun exposure, bright green, leafy
77–88
Legume hays, including alfalfa, good green color, leafy
40–59
Legume hays, including alfalfa, partly bleached, moderate amountof green color
20–31
Legume hays, including alfalfa, badly bleached, or discolored, tracesof green color
9–18
Non-legume hays, including timothy, cereal, and prairie hays, well cured,
good green color
20–31
Non-legume hays, average quality, bleached, some green color
9–18
Legume silage (wet basis)
11–44
Corn and sorghum silages, medium to good green color (wet basis)
4–22
Grains, mill feeds, protein concentrates, and by-product concentrates,
except yellow corn and its by-products
0.02–0.44
Sources: Adapted from Scott et al. (1982) and Maynard et al. (1979).

beaded, or prilled products (Shields et al., 1982). Nevertheless, vitamin
A supplements should not be stored for prolonged periods prior to feeding.
Vitamin A and carotene destruction also occurs from processing offeeds with steam and pressure. Pelleting effects on vitamin A in feed arecaused by die thickness and hole size, which produce frictional heat anda shearing effect that can break supplemental vitamin A beadlets and47

CHAPTER TWO

expose the vitamin. In addition, steam application exposes feed to heatand moisture. Running fines back through the pellet mill exposes vitamin A to the same factors a second time. Between 30 and 40% of vitamin A present at mixing may be destroyed during pelleting (Shields etal., 1982).

DEFICIENCY
Effects of Deficiency
Vitamin A is necessary for normal vision in animals and humans,
maintenance of healthy epithelial or surface tissues, and normal bonedevelopment. The vitamin A deficiency signs observed in ruminants vary
somewhat, but most relate to these three changes in tissues. Numerous
studies have also demonstrated increased frequency and severity of
infection in vitamin A-deficient animals. Lack of vitamin A results indecreased antibody production and impaired cell-mediated immune
processes against infective agents (Davis and Sell, 1983). Clinical signs
may be specific for vitamin A deficiency, or only general signs may beobserved, including loss of appetite, loss of weight, unthrifty appearance, thick nasal discharge, and reduced fertility. The normal epitheliumin various locations throughout the body becomes replaced by a stratified, keratinized epithelium when vitamin A is deficient. This effect has
been noted in the respiratory, alimentary, reproductive, and genitourinary tract as well as in the eye. A wide range of signs of vitamin A deficiency are noted in Table 2.7.

Ruminants
Ruminants lacking vitamin A may be more susceptible to pinkeye orother diseases related to the mucous membranes. Keratinization lowers
the resistance of the epithelial tissues to the entrance of infectious organisms. Thus respiratory diseases, such as colds and sinus infections, tend
to be more severe when vitamin A is deficient.
Vitamin A deficiency could indirectly result from zinc deficiency.
Zinc deficiency, interferes with the synthesis of retinol-binding protein(RBP) in liver, which carries vitamin A (retinol) in plasma. Thus, in zinc
deficiency, decreased liver RBP levels may cause low concentrations ofplasma vitamin A. Zinc-deficient goats have been observed to have low
serum vitamin A despite adequate dietary vitamin A (Chhabra et al.,1980). In calves, serum vitamin A was significantly higher for animals

48

V I TA M I N A■ Table 2.7

Signs of Vitamin A Deficiency

General
Cessation of growth
Cystic pituitary glands
Death
Decline in body weight
Diarrhea (scours)
Failure of appetite
General edema
Reduced resistance to parasite infections
Untidy hair or feathers
Xerosis of membranes
Bone Formation
Cancellous bone
Defective modeling
Narrowing of foramina
Restriction of brain cavity
Congential Abnormalities
Anophthalmia
Aortic arch deformities
Cleft palate
Hydrocephalus
Kidney deformities
Microophthalmia
Defective Reproduction
Abnormal estrous cycle
Dead, weak, or blind offspring
Degeneration of testes
Reduced egg production and hatchability
Resorption of fetuses
Eyes
Keratomalacia
Lacrimation
Night blindness (nyctalopia)
Xerophthalmia
Constriction of optic nerve
Loss of lensOpacity of cornea
Papilledema

Liver
Degeneration of Kupffer’s cells
Metaplasia of bile ducts
Nervous System
Constriction at foramina
Convulsive seizure
Hydrocephalus
Incoordination and staggering gait
ParesisRaised cerebrospinal fluid pressure
Twisting of nerve
Respiratory System
Lung abscesses
Metaplasia of nasal passages
Nasal discharge
Pneumonia
Urinary System
Cystitis
Nephrosis
Pus in ureters
PyelitisThickening of bladder wall
Urolithiasis

Source: Modified from Bauernfeind and DeRitter (1972).

supplemented with 50 mg/kg zinc (Chhabra and Arora, 1987). Cattle
from tropical northern Australia showed a 12% annual mortality in part
because of a slow release of liver vitamin A (Guerin, 1981). Apparently,
high calcium and low forage zinc concentrations contributed to thisslow liver vitamin A release. Since tropical forages have often beenshown to be low in zinc (McDowell, 1997), conditioned vitamin A deficiency may result even though liver vitamin A values indicate adequate
concentrations of this vitamin.

49CHAPTER TWO

VITAMIN A DEFICIENCY IN CATTLE
In cattle, signs of vitamin A deficiency (Figs. 2.9–2.11) include
reduced feed intake, rough hair coat, edema of the joints and brisket,
lacrimation, xerophthalmia, night blindness, slow growth, blindness,
low conception rates, abortion, stillbirths, blind calves, abnormal
semen, reduced libido, and susceptibility to respiratory and other infections (NRC, 1996). Animals in advanced stages of deficiency may exhibit a staggering gait, convulsive seizures, and papilledema, resulting fromelevated cerebrospinal fluid pressure.
The greatest need for vitamin A is during calving and breeding. If it
is inadequate, calves may get pinkeye, pneumonia, or other diseases
related to the mucous membranes. Vitamin A deficiency lowers reproductive efficiency in both males and females. Reduced libido and sterility in bulls with degeneration of seminiferous tubules has been reported(Larkin and Yates, 1964). Spermatozoa decrease in number and motility, and abnormal forms markedly increase. In cows, key indications of
the deficiency are lowered conception rates; shortened pregnancies,
either as abortions or reduced gestation length; high incidence ofFig. 2.9 Vitamin A-deficient calf. Note the emaciated appearance and evidenceof diarrhea. The calf also shows excessive lacrimation and nasal discharge
characteristic of vitamin A deficiency. (Courtesy of G. Patterson, Chas. Pfizer
Inc.)
50

V I TA M I N A

Fig. 2.10 Advanced
stage of anasarca inhindquarters of vitamin
A-deficient steer.
(Courtesy of L.L.
Madsen, Washington
State University.)

retained placenta; and birth of dead, weak, incoordinated, or permanently blind calves caused by bone abnormalities in the optic foramenthat constricts the optic nerve (Miller, 1979). If born alive, weak calves
have trouble getting to their feet and lack the instinct to nurse. Vitamin
A-deficient newborn calves may have very severe diarrhea, which may
soon be followed by death. In young calves, signs of vitamin A deficiency also include watery eyes and nasal discharge, and sometimes muscular incoordination, staggering gait, and convulsive seizures. However, inthe calf, elevated cerebrospinal fluid pressure is the earliest change specific for vitamin A deficiency.
The classic sign of vitamin A deficiency in cattle is night blindness.
As vitamin A deficiency develops, the animal’s adaptation to darkness is
reduced, and night blindness occurs; this condition is readily detected
when animals encounter obstacles in dim light. Blindness or night blindness may be the first noticeable sign of vitamin A deficiency in rapidly
growing cattle fed high-concentrate diets. In severe vitamin A deficiency, characteristic changes occur in the eye, including excessive watering,keratitis, softening and cloudiness of the cornea, and development ofxerophthalmia characterized by drying of the conjunctiva. However, inthe eyes of cattle, copious lacrimation (rather than xerophthalmia) is themost prominent clinical sign of vitamin A deficiency (Maynard et al.,51

CHAPTER TWO

Fig. 2.11 Vitamin A-deficient calf shows incoordination and weakness.
(Courtesy of J.W. Thomas, Michigan State University.)1979). Blindness may follow eye infection caused by the deficiency.
Vitamin A is involved in normal bone development; when it is deficient, the bones are altered in shape during growth, and the teeth areaffected. Failure of the spine and some other bones to develop normally
results in pressure and degeneration of the nerves. For example, blindness in calves results from constriction of the optic nerve caused by anarrowing of the bone canal through which it passes (Maynard et al.,1979). Bone changes may also be responsible for muscular incoordination and other nervous signs shown by vitamin A-deficient cattle.
In finishing cattle, generalized edema may occur, with signs of lameness in the hock and knee joints and swelling in the brisket area (NRC,
1996). Booth et al. (1987) reported that feedlot cattle had low concentration of serum vitamin A; appeared blind; and had fixed dilated pupils,
severe ataxia, and poor weight gains. Feedlot cattle with mild vitamin A
deficiency reduce their feed intake and do not make satisfactory weightgains. Lowered feed intake may result in deficiencies of other nutrients
when the diet is borderline in these nutrients.
As part of the major role of maintaining a healthy epithelium, vitamin A and β−carotene have been shown to have an important role in
reducing the incidence and severity of mastitis in dairy cows (Oldham et
52

V I TA M I N A

al., 1986; Chew, 1987). A diet adequate in vitamin A and β−carotene arenecessary for nutritional protection against this disease.
Vitamin A-deficient cattle have depressed activity of natural killer
cells, decreased antibody production, decreased responsiveness of lymphocytes to mitogenic stimulation, and increased susceptibility to infection (Ross, 1992; Nonnecke et al., 1993; Michal et al., 1994; Rajaraman
et al., 1998).
Vitamin A helps maintain normal immune resistance (immunostimulation), while the antioxidant activity of β−carotene increases the bactericidal activity of blood and milk polymorphonuclear neutrophils
against S. aureus (Daniel et al., 1991). In addition, some studies haveshown that β−carotene has a favorable effect on fertility of heifers andlactating cows (Bonsembiante et al., 1980). However, other studies(Akordor et al., 1986; Oldham et al., 1991) have shown no effect of β−carotene on reproductive performance or incidence of mastitis.
Additional studies are needed to clarify the physiological role of β−carotene as differences in results may relate to different body stores ofboth β−carotene and vitamin A in experimental animals.
In the 1950s, it was discovered that cattle developed signs of vitamin A deficiency that was originally referred to as X disease (hyperkeratosis). This disease was due to feeds that contained highly chlorinatednaphthalene found in lubricating oil. Because of the depressed vitamin
A levels in blood plasma, it was concluded that the toxic substance interfered with the conversion of carotene to vitamin A (Maynard et al.,1979). Removal of naphthalenes from oils eliminated X disease.
Studies have shown that vitamin A-deficient cattle lack heat tolerance. Deficient cattle stood and panted considerably, and their daily feed
consumption decreased (Perry, 1980), while vitamin A-supplemented
cattle showed improved hot-weather tolerance and spent much of thetime chewing their cud.

VITAMIN A DEFICIENCY IN SHEEP
Clinical signs of vitamin A deficiency in sheep (Fig. 2.12) are similar to those in cattle; night blindness is the common means of determining the deficiency (NRC, 1985). Vitamin A deficiency results in keratinization of the respiratory, alimentary, reproductive, and urinary
tracts, and ocular epithelia. Keratinization causes lowered resistance toinfections. The immune response is decreased in lambs with low vitamin
A status (Bruns and Webb, 1990).
Additional clinical signs of vitamin A deficiency in sheep include
growth retardation, bone malformation, degeneration of the reproduc53

CHAPTER TWO

Fig. 2.12 Typical appearance of a vitamin A-deficient lamb. Note the extreme
weakness and swayed back. This was followed by the inability to stand.
(Courtesy of T.J. Cunha and Washington State University.)

tive organs, and elevated pressure in cerebrospinal fluid. Deficiency
interferes with normal development of bone, which may relate to muscular incoordination and nervous signs. Vitamin A deficiency can also
result in lambs born weak, malformed, or dead. In addition, retainedplacenta occurs in vitamin A-deficient ewes.
Vitamin A deficiency has resulted in low semen quality in rams
(Lindley et al., 1949). Vitamin A deficiency has detrimental effects onwool production and characteristics, including shortened wool fibers
and decreased fiber thickness, strength, and elongation (Farid andGhanem, 1982).

VITAMIN A DEFICIENCY IN GOATS
In goats, vitamin A deficiency often appears first as a rough, dullhair coat condition. Goats deficient in vitamin A show keratinization of
the epithelia of the respiratory, alimentary, reproductive, and urinary
tracts, and of the eye (NRC, 1981). Signs include multiple infections,poor bone development, birth of abnormal offspring, and vision
impairment. Night blindness is the classic sign of deficiency. Experimentally produced signs of vitamin A deficiency in goats include loss of54

V I TA M I N A

appetite, loss of weight, unthrifty appearance, night blindness, and thicknasal discharge (Schmidt, 1941).
In India, limited work with vitamin A-deficient goats suggests thatthe deficiency may lead to urinary calculi (Majumdar and Gupta, 1960).
Lack of vitamin A may produce temporarily impaired fertility or permanent infertility and, after kidding, metritis may be caused by damage
to the integrity of the uterine mucosa (Guss, 1977). In fully grown goats,
reduced fertility of the female and male is the more common clinicalsign; the animals show poor conception rates during short or delayedheat periods, and reduced semen quality.

Swine
In pigs, the absence of vitamin A results principally in nervous signs
such as unsteady gait, incoordination, trembling of the legs, spasms, andparalysis (Hentges et al., 1952) (Fig. 2.13). Eye lesions are less common.
In pigs, the effect of vitamin A deficiency on appetite or rate of gain does
not occur until eventual paralysis and weakness prohibit movement to
the feeder (Cunha, 1977).
During reproduction and lactation, a vitamin A deficiency in thesow produces the following clinical signs: failure of estrus, resorption ofyoung, wobbly gait, weaving and crossing of the hind legs while walking, dropping of the ears, curving with head down to one side, impaired
vision, spasms, and loss of control of hindquarters and forequarters andthus inability to stand up (Cunha, 1977). Depending on degree of severity of vitamin A deficiency, fetuses were resorbed, born dead, or carried
to term. Fetuses carried to term had a variety of defects, including various stages of arrested formation of the eyes as well as complete lack ofeyeballs. Other defects included harelip, cleft palate, misplaced kidneys,
accessory earlike growths, having only one eye, having one large andone small eye, and bilateral cryptorchidism (Cunha, 1977). Vitamin A
appears to improve reproductive performance of gilts by decreasing
embryonic mortality, resulting in more pigs per litter (Brief and Chew,
1985).
Field trials have indicated that litter size could be increased by 0.6
to 1.5 pigs per litter by injecting sows with vitamin A at weaning(Whaley et al., 1997). The correlation between size of the embryo andthe retinol content in uterine flushings is high (r = 0.97) (Trout et al.,
1992; Seijas, 1995). Retinol in the uterine secretions may function byprotecting the conceptus against the oxidizing activity of uteroferrin(Vallet, 1995). Consequently, RBP may have several functions in the
reproductive tract of pigs: (1) transfer of retinol to the fetus, (2) protec55

A

B

C

Fig. 2.13 Vitamin A deficiency in growing pigs: (A) partial paralysis and seborrhea, (B) initial stage of spasm, (C) lordosis and weakness of hind legs.
(Courtesy of J.F. Hentges, R.H. Grummer, and the University of Wisconsin.)56

V I TA M I N Ation of tissues against oxidative reactions, and (3) provision of a substrate to generate retinoic acid and other biologically active metabolites
of retinol.
The effect of administering vitamin A on subsequent reproductive
performance has had mixed responses (Pusateri et al., 1999). Coffey and
Britt (1993) demonstrated that after the injection of 200 mg of provitamin A (β−carotene) at weaning, litter size was increased in multiparous
but not in primiparous sows. In contrast, Tokach et al. (1994) conducted a study on a commercial swine farm with 956 sows and demonstrated that a single injection of β−carotene, vitamin A, or a combination of
the two at weaning did not affect subsequent reproductive performance.

Poultry
Clinical signs of vitamin A deficiency are similar among chickens,turkeys, and other poultry species. On the same low vitamin A diet,turkeys and Japanese quail would exhibit clinical signs earlier thanchickens due to their high requirement for the vitamin. In poultry, lackof vitamin A in the diet causes slower growth (Fig. 2.14), lowered resistance to disease, eye lesions, muscular incoordination (Fig. 2.15), andother signs (Scott et al., 1982). As deficiency progresses in adult poultry,
the chickens become emaciated and weak, and their feathers are ruffled.
A marked decrease in egg production occurs, and the length of time
between clutches increases greatly. Hatchability is decreased, andembryonic mortality in eggs from affected birds increases. A watery discharge from the nostrils and eyes is noted, and eyelids are often stuck
together. Average survival time of the progeny fed a diet with no vitamin
A increased in a linear fashion with the increasing levels of vitamin A in
the maternal turkey diet (Jensen, 1965).
Vitamin A level of the hen’s diet is positively correlated with thegrowth of chicks and poults from that hen, and the level of vitamin A inthe chick’s and poult’s diet is positively correlated with growth. Clearly,
a deficiency of vitamin A will produce loss of appetite and reduction in
growth. Severe deficiency will produce ataxia and death if not corrected(Hill, 1961).
When day-old chicks are given a diet with no vitamin A, clinicalsigns may appear at the end of the first week if the chicks are progenyof hens receiving a diet low in vitamin A. If chicks are progeny of hens
receiving high levels of vitamin A, signs of deficiency may not appear
until chicks are 6 or 7 weeks of age even though they are receiving a dietcompletely devoid of vitamin A (Scott et al., 1982). Gross signs of vitamin A deficiency in chicks are characterized by anorexia, cessation of57

CHAPTER TWO

Fig. 2.14 Six-week-old turkeys: Left: fed a vitamin A-deficient diet; Right: feda control diet. (Courtesy of L.M. Potter, Virginia Polytechnic Institute.)

growth, drowsiness, weakness, incoordination, emaciation, and ruffled
plumage. The mucous epithelium is replaced by a stratified squamous,
keratinizing epithelium. As a result of mucous membrane breakdown,
bacteria and other pathogenic microorganisms may invade these tissues
and enter the body, thereby producing infections that are secondary to
original signs of vitamin A deficiency. The mucous membranes of thenasal passage, mouth, esophagus, and pharynx of poultry are affected
and develop white pustules (Scott et al., 1982). The kidneys may be distended with uric acid deposits, and the epithelium of the eye is affected,
producing exudates and eventually resulting in xerophthalmia. Clearly,
there is a breakdown of the epithelium of many systems in the body due
to vitamin A deficiency. Loss of membrane integrity, in turn, alters water
retention (Lopen et al., 1973) and impairs the ability to withstand infection (Sijtsma et al., 1989). Inadequate vitamin A also reduces theimmune system’s response to challenge and further contributes to disease susceptibility (Davis and Sell, 1989). Viral infections have beenfound to impair the vitamin A status of chickens (West et al., 1992).
Vitamin A deficiency can cause alterations in bone growth, which
create several areas of compression on the central nervous system, causing a loss in mobility (Howell and Thompson, 1967). Nockels (1988)
58

V I TA M I N A

Fig. 2.15 Ataxia in chicks due to dietary deficiency of vitamin A. (Courtesy of
M.L. Scott, Cornell University.)

reported that hypothyroidism is an early indication of vitamin A deficiency in chicks. Reductions in testes size, circulating testosterone, andfertility have been reported in vitamin A-deficient cockerels (Hall et al.,1980).

Horses
The importance of vitamin A to the overall health and well-being ofhorses has been well documented. Night blindness, lacrimation, keratinization of the cornea and respiratory system, reproductive difficulties,
capricious appetite, progressive weakness, and death occur in horses
deficient in vitamin A (NRC, 1989b).
During the winter, vitamin A depletion has been observed in grazing
horses (Greiwe-Crandell et al., 1997) and in stabled horses fed conserved feeds (Mäenpää et al., 1988). Vitamin A depletion had adverse
effects on growth and reproduction in horses (McDowell et al., 1995).
In a lengthy experiment with vitamin A-deficient horses, all horses
developed night blindness that abated when dietary vitamin A was provided (Howell et al., 1941). Upon withdrawal of vitamin A supplements,night blindness again appeared in the animals.
Histological changes in the retinas of vitamin A-deficient horses
59

CHAPTER TWO

have been shown (NRC, 1989b). Copious lacrimation is characteristically associated with vitamin A deficiency along with varying degrees ofcorneal keratinization.
Greiwe-Crandell et al. (1997) reported that vitamin A status indepleted mares was improved by supplementation with retinyl palmitate. However, in mares with no access to pasture, supplementation withtwice the currently recommended amount of vitamin A barely matchedthe vitamin A status of unsupplemented mares on green pasture.

Other Animal Species
CATS
Vitamin A deficiency in cats is characterized by lack of growth,loss of weight, and poor appetite after 2 to 3 months of feeding a diet
deficient in vitamin A. Muscle wasting is severe, while stored body fat
may be nearly normal. Epithelial tissues and mucous glands forming
the membrane lining of the alimentary, respiratory, urinary, andreproductive tracts as well as the membrane covering the eye and eyelids are markedly impaired by vitamin A deficiency (Gershoff et al.,1957).
Night blindness can be detected early in vitamin A-deficient kittens.
As the deficiency progresses, the pupillary reflex to light is delayed from
1 to 2 seconds in the normal state to 5 to 10 seconds in the deficientstate. Irreversible degeneration occurs in the rods and cones (visual cells)of the retina. Unfortunately, assessment of night blindness in cats has
not been adequately distinguished from retinal degeneration due to taurine deficiency (Knopf et al., 1978), since all experimental A-deficient
diets to date have used casein as the source of protein, a procedure
which depletes taurine in cats.
With vitamin A deficiency, bones become thickened and constrict
the central nervous system. Ataxia, stiff gait, and hydrocephalus in newborn kittens from deficient dams suggest increased cerebrospinal fluidpressure, whereas the description of cleft palate suggests deranged
intrauterine mitosis and cell differentiation (Ralston Purina, 1987).
Male cats become sterile from testicular hypoplasia; females withsevere vitamin A deficiency do not ovulate. With a lesser degree of deficiency, queens conceive and implantation occurs, but resorption or abortion of the fetus results at approximately the 49th day of pregnancy(Gershoff et al., 1957).
Bartsch et al. (1975) described ataxia, star gazing, blindness, andintermittent convulsions in African lion cubs presumed to be vitamin A60

V I TA M I N A

deficient. There was severe thickening of the cranium, compression of
the brain, and partial herniation of the cerebellum.

DOGS
Vitamin A deficiency is usually confined to young animals, andadvanced deficiency is always associated with cessation of growth oreven weight loss and eye problems. Signs of xerophthalmia (excessive
dryness of the eye), night blindness, conjunctivitis, corneal opacity andulceration, and skin lesions are associated with rough hair coat, anorexia, growth depression, and muscle weakness (Singh et al., 1965).
Respiratory infection in prolonged deficiency often results in death.
Deafness and facial paralysis occurred in puppies with severe chronic deficiency when bone growth of the skull was altered by inadequate
remodeling and excessive periosteal growth to constrict cranial nerves.
Blood and liver concentrations of vitamin A were negligible in thesecases. Mellanby (1938) established that such damage to the cochlear andvestibular divisions of the eighth cranial nerve, plus serious labyrinthitis, may induce deafness. Similar damage may also affect function of the
optic nerve, although this only has been observed after prolonged experimental deficiency (NRC, 1985a).
Vitamin A deficiency is generally a disease in growing pups, as
adults seldom develop signs due to their extremely slow depletion ofliver stores. In fact, several litters of pups have been produced while thedam was maintained on a vitamin A-deficient diet (Ralston Purina,1987).

FISH
General signs of vitamin A deficiency in fish are impaired growth,
exophthalmos, eye lens displacement, corneal thinning and expansion,degeneration of retina, edema, and depigmentation (NRC, 1993).
Vitamin A deficiency in rainbow trout causes anemia, twisted gill opercula, and hemorrhages in the eyes and base of fins (Kitamura et al.,1967). Brook trout exhibit poor growth, high mortality, and eye lesions,such as edematous eyes, displaced lens, and degeneration of the retina
when fed a vitamin A-deficient diet (Poston et al., 1977). Anorexia, pale
body color, hemorrhagic skin and fins, exophthalmia, and twisted gill
opercula occur in vitamin A-deficient common carp (Aoe et al., 1968).
Rapidly growing yellowtail fingerlings fed a vitamin A-deficient dietdevelop deficiency signs in 20 days, including arrested growth of gill
opercula, dark pigmentation, anemia, and hemorrhage in the eyes andliver, accompanied by high mortality (Hosokawa, 1989).
61

CHAPTER TWO

FOXESFoxes fed a diet deficient in vitamin A develop a series of nervous
derangements, usually manifested in this order: first, trembling or cocking of the head; next, unsteadiness (resulting from a disturbed sense ofbalance); then a tendency to run in circles (NRC, 1982a).

LABORATORY ANIMALS
Signs of vitamin A deficiency can be divided into six categories(Smith, 1990; NRC, 1995): (1) defect in vision, (2) bone defects, (3)increase in cerebral spinal fluid pressure, (4) reproductive failure (cessation of spermatogenesis in males and cornification of the reproductive
tract in females), (5) epithelial metaplasia and keratinization and (6)growth failure. Insufficient vitamin A in rats leads to reduced efficiencyof the urea synthesis pathway, thus accounting for the increased amino
nitrogen excretion that is seen with the deficiency.
In mice, one of the early and significant consequences of vitamin A
deficiency is impairment of the functional immune system. As the deficiency progresses, many epithelial tissues become keratinized, including
those of the seminal vesicles, testes, bladder, kidney, trachea, esophagus,
salivary glands, and lungs (Van Pelt and De Rooij, 1990; NRC, 1995).
In the guinea pig, the first evidence of vitamin A deficiency is poor
growth, then weight loss, followed by incrustations of eyelids and severedermatitis resulting from bacterial infection (NRC, 1995). Often, animals develop pneumonia prior to death. Vitamin A-deficient hamsters
develop abnormally and have coarse and sparse hair, xerophthalmia,
and keratinized stratified tracheal lining (NRC, 1995).

NONHUMAN PRIMATES
In rhesus and cebus monkeys, the first manifestations of deficiency
are weakness, diarrhea, loss of appetite, and an apparent increase in susceptibility to respiratory infection (NRC, 1978). With more severe deficiency, pathological changes in the eye become apparent. Two of four
pregnancies carried to term by rhesus monkeys, which were maintainedon marginal intakes of vitamin A, produced infants with congenital
xerophthalmia, and a third infant developed xerophthalmia after receiving a vitamin A-deficient diet for 2 years (NRC, 1978).

RABBITS
Signs of vitamin A deficiency in rabbits are similar to thosedescribed for other animals and include retarded growth, neural lesions,
62

V I TA M I N A

ataxia, spastic paralysis, xerophthalmia, and impaired reproduction(NRC, 1977).

HumansThroughout history, night blindness and xerophthalmia have been
the major signs of vitamin A deficiency in humans. Xerophthalmia
includes various stages of conjunctival xerosis, Bitot’s spots, and corneal
involvement and ulceration. Bitot’s spots—foamy white accumulationsof sloughed cells that usually appear on the temporal quadrant of the
conjunctiva—are used as a clinical indicator of vitamin A deficiency inyoung children (Olson, 1991). Skin changes have also been observed,
most specifically follicular hyperkeratosis (Olson, 1991).
Xerophthalmia is the most common cause of blindness in young
children throughout the world (Fig. 2.16), with as many as 5 million
Asian children developing xerophthalmia each year (Sommer et al.,1981). One-tenth of children with xerophthalmia have severe corneal
involvement, and half of them become blind. Xerophthalmia is recog-

Fig. 2.16
Keratomalacia in a
Jordanian infant withsevere vitamin A deficiency. The central
area of the cornea isundergoing typicalcolliquative necrosis.
Sight is irrevocably
lost at this advanced
stage. (Courtesy of
D.S. McLauren, Royal
Infirmary, Edinburgh,
Scotland.)63

CHAPTER TWO

nized as endemic in many countries in southern and eastern Asia, partsof Latin America, and in many countries in Africa and the Middle East.
Vitamin A deficiency remains the second major nutritional problem
among children in many countries, including Bangladesh, India,
Indonesia, Brazil, and other developing countries. An estimated 20 to 40
million children worldwide have at least mild vitamin A deficiency, nearly half of whom are said to reside in India (United Nations
Administrative Committee, 1987).
Vitamin A deficiency affects about 100 million young children
worldwide and has been long known to cause blindness. It has now
become increasingly clear that even mild vitamin A deficiency also
impairs the immune system, reducing children’s resistance to diarrhea,
which kills 2.2 million children a year, and measles, which kills nearly 1
million annually (UNICEF, 1998).
The group most vulnerable to vitamin A deficiency is children frombirth to 6 years of age because of their low liver reserves; low protein,lipid, and vegetable intakes; and increased requirements due to rapid
growth rate. Likewise, vitamin A deficiency for which night blindness isa marker is a problem for pregnant lactating women in rural Nepal, withhealth consequences for women and their infants (Katz et al., 1995). The
mortality rate among children with mild xerophthalmia (night blindness
and/or Bitot’s spots) is on average four times the rate, and in some agegroups, 8 to 12 times the rate, of children without xerophthalmia(Sommer et al., 1986). In Indonesia, a 34% difference in mortality wasobserved between preschool children who received vitamin A supplements for 1 year and those who received none (Sommer et al., 1986).
An early manifestation of vitamin A deficiency is premature growth(West et al., 1997) followed by infection of a wide variety of organs,overwhelming sepsis, and death. Death may frequently precede the onsetof eye manifestations of deficiency. Many studies have shown that evenmild vitamin A deficiency leads to patchy keratinization of the epithelial
lining of the gastrointestinal tract, respiratory tract, urogenital tract, andskin. Five to six times more children in the developing world are likely
to be subclinically than clinically deficient in vitamin A (Underwood,1994).
Lack of vitamin A has been associated with an increased susceptibility to infections, especially of the respiratory and digestive tracts(Milton et al., 1987). Immune mechanisms become impaired and antibody synthesis is diminished. Large-dose vitamin A supplementation has
improved vitamin A status and reduced childhood mortality(Rahmatullah et al., 1991; West et al., 1991; Fawzi et al., 1994). Regular64

V I TA M I N A

intake of food rich in vitamin A is associated with reduced incidence of
diarrhea and measles (Fawzi et al., 1995).
Vitamin A deficiency in humans occurs in endemic proportions inmany developing countries and is seen only occasionally in technologically developed societies in patients with severe malabsorption, transport disorders, or liver disease (McLaren, 1984). It has been reported,
however, that alcohol consumption results in significant hepatic vitamin
A depletion, due perhaps to associated failure to consume an adequate
diet. Also, drugs such as phenobarbital or food additives such as butylated hydroxytoluene (BHT), when combined with ethanol, result in a
striking depletion of hepatic vitamin A concentrations in rats (Leo et al.,1987).
Vitamin A deficiency in humans under natural conditions is notwholly comparable to the human volunteers or experimental animals onan otherwise good diet lacking only vitamin A. Under field conditions,vitamin A deficiency is nearly always accompanied by protein-energy
malnutrition, is exacerbated by parasitic infections, and is frequently
complicated by intercurrent infections. Diarrheal diseases are a major
cause of childhood morbidity and mortality in developing countries,
causing approximately 4 million deaths per year in children younger
than 5 years of age (Claeson and Merson, 1990). In areas where vitamin
A status is suboptimal, children with even mild vitamin A deficiency
appear to be at greater risk of developing diarrheal diseases than thosewith adequate vitamin A status (Bhandari et al., 1997). Chronic diarrhea is associated with reduced absorption of vitamin A (Liuzzi et al.,1998). It has been postulated that between 90 and 100 million children(predominantly in developing countries) who have some level of vitamin
A deficiency are at risk of poor health and death (Underwood, 1994).

Deficiency Incidence and Relation to Vitamin A
Body Stores
The circumstances most conducive to vitamin A deficiencies in livestock are (1) extended periods of drought, resulting in pastures becoming
dry and bleached with no green color; (2) diets composed primarily ofconcentrates and no green pasture; (3) feeding mainly corn silage and aconcentrate mixture low in vitamin A activity; (4) young animals fed milk
from mothers on low intake of vitamin A or carotene, or poultry hatchedwhere previous diets were low in vitamin A value; (5) young animals fed
relatively little whole milk or colostrum; and (6) substituting yellow corn
for grains containing no carotene. Mild vitamin A deficiency, especiallyin winter (or during the dry season) and early spring, is fairly common.
65

CHAPTER TWOWhen grazing livestock receive a modest amount of fresh green pasture forage, there is little likelihood of deficiency. Likewise, with a substantial amount of good silage made from green forage, or with liberal
feeding of fresh hay that has a good green color, deficiency will notoccur. However, low-quality forages and weathered and leached hay
may contain very little available carotene. Amounts of carotene in fresh
green forages are very high relative to dietary requirements. Thus, asmall quantity of fresh pasture forage will fully supply the need for vitamin A.
One of the most frequent stress conditions for grazing livestock inmany parts of the world is low intake of carotene under conditions of
winter feeding or protracted drought. When normal seasonal cycles prevail, and rainfall is sufficient to provide 4 months or more of green grazing, mature animals store sufficient vitamin A to carry them for many
months on dry feed, including grain and roughage that is low incarotene. However, when severe and prolonged drought conditions prevail, especially during the normal green grazing period, signs of vitamin
A deficiency may occur in adult animals. Geographical locations where
vitamin A deficiency might be expected would therefore be in regionswith long dry seasons. Many tropical regions in Africa, Asia, and Latin
America routinely have dry seasons lasting 6 months or longer. In India,
conditions reportedly permit fair grazing for only 3 months of the year
(Ray, 1963). With the cessation of the monsoon rains, grasses mature
rapidly, with a large drop in their carotene content. The value has beenfound to decrease from 100 to 200 mg per kilogram on a dry-matterbasis in the middle of the rainy season to as low as 0.5 to 1 mg per kilogram during the dry season. As a result, clinical signs such as night
blindness, blindness in newborn calves, and birth of weak calves havebeen reported in all parts of India.
In the various vitamin A supplementation trials to improve ruminant livestock production, contradictions on the benefits of supplementation have been reported. Overall, it has been shown beneficial to supplement grazing livestock with vitamin A, especially in middle to late dry
season, and to feedlot cattle not receiving green forages. A general ruleof thumb is that forages with a bright green color are always adequate
in vitamin A value (carotene).
Deficiencies of other nutrients interfere with vitamin A metabolism
and can result in vitamin A deficiency. Both protein and zinc have beenshown to be required for mobilization of vitamin A from liver (seeDeficiency, Ruminants). Vitamin A is required for intestinal absorptionof zinc (Berzin and Bauman, 1987) in poultry, while zinc influences vita66

V I TA M I N A

min A by affecting RBP synthesis and release from the liver (Brown etal., 1976).
Liver is the main vitamin A storage site for humans as well as forfarm species. Although most of the vitamin A reserves are in the liver,when carotene intake is high, some is stored in fat. During periods oflow dietary carotene, this stored vitamin A can be mobilized and utilized
without signs of vitamin A deficiency. At birth, the ruminant usually
does not have sufficient vitamin A reserves to provide for its needs forany substantial time. Accordingly, it is important that young ruminants
receive colostrum, which generally is rich in vitamin A, within a few
days after birth. If the cow has received a diet low in vitamin A, thenewborn calf is likely to be susceptible to a vitamin A deficiency because
body reserves are low and colostrum will have a subnormal content(Miller et al., 1969). Likewise, if the hen received a diet deficient in vitamin A, chicks will develop the deficiency.
Grazing livestock with access to green, high-quality pastures canstore sufficient vitamin A in the liver to be adequate for periods of low
intake during the winter or dry season, perhaps as long as 4 to 6 months.
Cattle grazing good pasture will have 30 to 80 ppm of liver vitamin A(Rumsey, 1975), and cattle entering the feedlot with 20 to 40 ppm will
have adequate liver stores for 3 to 4 months (Perry et al., 1967).
Intramuscular injection of 1 million IU of emulsified vitamin A apparently provides sufficient vitamin A to prevent deficiency signs for 2 to 4
months in growing or breeding beef cattle (NRC, 1996).
Because of vitamin A storage, sheep that graze on green forage during the normal growing season may be able to do reasonably well on a
low-carotene diet of dry feed for periods of 4 to 6 months (NRC,
1985b). Goats that have had access to good-quality green feed can probably depend on vitamin A stores for a minimum of 3 months without
detrimental effects (NRC, 1981). The tendency of goats to search out
palatable green plant parts ensures it an advantage over other ruminant
species. However, goats that are forced to consume more conventional
cattle or sheep diets because of the unavailability of browse would nothave such an advantage (NRC, 1981).
Florida beef cattle finished on Roselawn St. Augustine grass during
summer did not require supplemental vitamin A for production; however, 25,000 IU per animal daily increased weight gains by approximately
10% when cattle were pastured during the winter (Chapman et al.,1964). In a study of nine cattle ranches from four regions in Florida, forage carotene and consequently liver vitamin A were lower during thewinter than the summer (Kiatoko et al., 1982). In this study, cattle in the67CHAPTER TWO

northern-most region, with fewer total grazing days, had significantly
lower liver vitamin A than cattle in the other three regions andapproached critical levels.
Storage of vitamin A is considerable for humans and farm species
that consume diets with high vitamin A value. If sows have built up
stores of vitamin A until they are 8 to 9 months of age and are thenplaced on a diet practically devoid of carotene or vitamin A, they still
can produce at least two normal litters (Selke et al., 1967). Well-nourished humans have at least several months’ supply that the body can utilize (RDA, 1989).

Assessment of Status
A number of criteria are available to evaluate the status of livestock,
including production response, liver vitamin A stores, plasma vitamin A,
and cerebrospinal fluid pressure. Vitamin A level of blood may also
reflect the nutritional status with respect to vitamin A (Green et al.,1987); however, blood vitamin A concentrations are governed by extent
of liver stores as well as by current intake. In fact, in some species, bloodlevel tends to be maintained until liver stores are exhausted when thediet is devoid of the vitamin. For this reason, a normal blood level cannot be interpreted to ensure that current intake is adequate, but a low
level indicates a deficiency.
A plasma vitamin A level less than 20 μg/100 ml in Holstein calves
suggests a deficiency (Eaton et al., 1970), and a plasma level of 10
μg/100 ml indicates an advanced deficiency. For beef cattle, Perry et al.
(1967) suggested that plasma vitamin A is probably an accurate indicator of a borderline deficiency, with a level of less than 40 μg of vitamin
A per 100 ml of blood serum indicating deficiency.
In dairy cattle, liver vitamin A values below 1 IU/kg were indicativeof a critical deficiency (NRC, 1989a). Feedlot performance of beef cattle was good as long as liver vitamin A stores were above 2 IU/kg(Kohlmeier and Burroughs, 1970). In dairy calves, cerebrospinal fluidpressure rises at a rapid rate after liver concentrations of vitamin A fall
below 1 to 2 ppm. Cattle showed a positive response to supplemental
vitamin A when their liver concentrations were as low as 3 to 4 ppm(Perry et al., 1967).
One cannot determine the exact status of vitamin A storage in the
pig by analyzing the blood for vitamin A. When vitamin A values drop
below 10 μg/g in the liver and below 10 μg/100 ml in the plasma, the pig
is quite deficient in vitamin A (Cunha, 1977). Liver biopsies for deter-

68

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mination of liver vitamin A values would give the best indication of vitamin A storage or status in the pig as well as for most species.
The vitamin A content of liver of newly hatched chicks is an excellent measure of the vitamin A nutrition of breeding hens. Vitamin A
deficiency will not occur in chickens having storage levels above 2 to 5
IU vitamin A per gram of liver. Thus, throughout the life of the chicken,
vitamin A content of liver serves as a good measure for assessing vitamin A status (Scott et al., 1982). Liver vitamin A concentration of calvesis associated with the carotene intake of their mothers (Flachowsky etal., 1993).
A simple test for night blindness evaluation in many animals (e.g.,dogs and cats) can be made by darkening an unfamiliar room andobserving the animals’ movements in dim red light (Ralston Purina,1987). More detailed analysis is possible with the electroretinogram in
the anesthetized animal.
The overall methods that are used to determine vitamin A status inhumans include dietary assessment, clinical evaluation, retinal function
tests, and biochemical assessment. These methods of evaluation havebeen discussed in a review article (Underwood, 1990).

SUPPLEMENTATION
Supplemental vitamin A may be administered (1) as part of a concentrate or liquid supplement, (2) as included with a free-choice mineral mixture, (3) as an injectable product, and (4) in drinking water preparations. The most convenient and often most effective way to provide
vitamin A to livestock is to include it with concentrate mixtures that will
provide uniform consumption of the vitamin. For grazing livestock,
however, providing vitamin A in concentrate mixtures generally is not
economically feasible. Most vitamin A preparations used today in animal applications are administered orally as an ingredient to blend uniformly in dry feed. With the advent of low-cost synthetic vitamin A, fishoils are used only to a limited extent in various world regions.
In feed manufacturing, liquids such as oils are sprayed onto the feedin the batch or continuous blending operation, the amount being controlled volumetrically. In some cases the use of synthetic vitamin A withantioxidants in oil dilutions has continued; the oil addition plays a secondary role of reducing dustiness of feed and/or supplying an addedsource of calories (Bauernfeind and DeRitter, 1972).
Because of the lack of stability of vitamin A, particularly regarding69

CHAPTER TWO

exposure to oxygen, trace minerals, pelleting, feed storage, and otherfactors, the feed industry has readily accepted the dry stabilized forms ofthe vitamin. Stabilized and protectively coated (or beaded) forms of vitamin A slow destruction of the vitamin, but for highest potency, fresh
supplies of the mixture should be available on a regular basis. Practicalconsiderations that affect vitamin A stability are listed in Table 2.8. The
gelatin beadlet in which the vitamin A ester (palmitate or acetate) is
emulsified into a gelatin-plasticizer-antioxidant viscous liquid formulation and spray-dried into discrete dry particles results in products withgood chemical stability, good physical stability, and excellent biologicalavailability (Bauernfeind and DeRitter, 1972).
Supplemental vitamin A may be needed when (1) feed consists ofpoor-quality forage with little or no green color, (2) diets are composed
primarily of concentrates and no green pasture, (3) feed consists mainlyof corn silage and a concentrate mixture low in vitamin A activity, (4)young ruminants are receiving milk from cows fed a diet low in vitamin
A or carotene, (5) calves are fed relatively little whole milk or colostrum,and (6) purchased cattle are of unknown background and in unthrifty
condition, and body stores of vitamin A might be suboptimal (Perry,1980).
Vitamin A assimilation was decreased, and its utilization wasincreased by infection (Nockels, 1988). Aflatoxin ingestion has also
been reported to reduce vitamin A stores in the liver and induce its deficiency. Chickens showed decreased tissue accumulation of oxycarotenoid pigments when fed aflatoxin. It is therefore important thatanimals that are sick or are receiving vitamin A antagonists receive dietsthat are fortified with the vitamin.
The use of concentrate feeds in place of forages is probably thelargest single factor that has increased the need for vitamin A supplements in ruminant diets. Inefficient utilization of carotene from corn andthe destruction of carotene and vitamin A in the rumen are reasons foradding supplemental vitamin A to high-concentrate diets, regardless ofthe vitamin A levels supplied by feedstuffs in these diets, to meet the
total daily requirement (Rumsey, 1975). Weiss (1998) suggests that therequirement for vitamin A should be increased by approximately 50%
for dairy cattle because of differences in bioavailability between βcarotene and retinyl esters.
Vitamin A is often included, along with vitamins D and E, in liquid
feed supplements in addition to molasses, fat, urea, and selected minerals. Since the viscosity, pH, and solids content of liquid feed supplements
vary considerably, the development of vitamin A forms that would blend
70

V I TA M I N A■ Table 2.8

Practical Factors That Affect the Stability of Vitamin A

Factors Detrimental to Stability
Long storage of the vitamin product before mixing or of the feed product after mixing
Vitamin premixes containing minerals
High environmental temperature and humidity
Pelleting, blocking, and extrusion
Hot feed bins that sweat inside upon cooling
Rancid fat in the feed
Rain leaks in feed bins or storage facilities
Factors Promoting Stability
Minimum time between manufacture of the vitamin product and consumption by
the animal
Storing vitamins in a cool, dark, dry area in closed containers
Not mixing vitamins and minerals in the same premix until ready to mix the feed
Not mixing deliquescent substances such as choline chloride with the vitamins
Using good-quality feed ingredients and vitamins
Good maintenance of storage bins and other equipment
Coordination between production and purchasing to obtain a quality product thatwill not be stored an excessively long time before use
Source: Modified by personal communication (1985) with Dr. Charles H. McGinnis
Jr., Manager of Nutritional Services, Rhône-Poulenc, Inc., Atlanta, Georgia.

uniformly and be stable in such a product of variable composition andcharacteristics represented a new challenge in the technology of vitamin
A application. Products of choice are liquid emulsions of fat-soluble
vitamins A, D, and E in a tested formulation of emulsifiers, antioxidants,
synergists, preservatives, and carriers.
For grazing livestock, vitamin A can be provided as part of a freechoice mineral mixture as an alternative to mixing it with the feed. The
greatest limitation of administering vitamin A with free-choice minerals
is unknown consumption by individual animals and destruction of thevitamin with time. Vitamin A stability in a mineral mix is affected byabrasion, moisture, and prooxidant action of trace metals, particularly
copper. Mitchell (1967) reported virtual destruction of gelatinized vitamin A when mixed with salt after 5 weeks. Apparently enough moisture
was picked up from humid air, and perhaps saliva from cattle, to induce
caking. This may have dissolved some of the gelatin coating, resulting inloss of vitamin A potency. Improved coatings of vitamin A are more
insoluble in water and prevent exposure to air and the catalyzing effectsof mineral contact.
Many livestock producers have followed the practice of intramuscular injection with a vitamin A concentrate. This has become a meansof administering vitamin A either as a therapeutic measure or, more frequently, as a prophylactic approach when oral or dietary administration
71

CHAPTER TWO

is inconvenient or impossible. Feedlot cattle with an unknown history
are often given an injection of vitamin A (e.g. 1,000,000 IU) as part of
the adaptation or preconditioning process before the animal adjusts tothe new environment and the high-energy fattening ration.
Recommended high doses of vitamin A for ruminants are presented inTable 2.9.
Very high doses of vitamin A are also important under intensive
housing conditions, where there are increased requirements in early life,
or under conditions of stress. The practice has been found particularly
valuable under the following circumstances (Hoffmann-La Roche,1967): for day-old chicks before transport; for laying hens when, forreasons unknown, laying performance suddenly drops; for piglets andcalves during weaning or changeover to milk substitutes; as an adjuvant
in treatment of attacks by intestinal parasites, lungworms, and otherparasites; and for newly arrived feedlot cattle.
Administration of vitamin A in very high levels in drinking water orby injection is recommended to support any specific measures used intreatment of diseased and convalescent animals. This is particularly true
for animals in which vitamin A stores may have been depleted due tofever or in animals suffering from intestinal disorders when vitamin A
absorption is seriously impaired. Also, very high levels of vitamin A may
be beneficial in reducing the incidence of mastitis in dairy cows (Chew,
1987).
The level of vitamin A supplementation used should be based primarily on the degree of the deficiency that could potentially be problematic. As with most nutrients, a borderline deficiency is more likely
than a severe deficiency. A marginal deficiency adversely affecting performance by a few percentage points is not easily detected (Miller,
1979). Based on the positive results that may be derived, and considering that vitamin A supplementation is inexpensive and (reportedly) nontoxic at recommended levels, it seems beneficial to administer supplemental vitamin A to livestock that are not grazing or receiving green pastures (roughages) and to those that are sick or under stress. Vitamin A
injection is one good way to determine if a deficiency exists by injecting
half the animals and using the others as controls. This can be done when
it would be difficult to conduct complicated experiments.
Supplementation of pet foods would be particularly essential if
organ meats rich in vitamin A (particularly liver and kidney) were notpart of the diets. Some commercial foods for dogs and cats containinsufficient vitamin A. In 89 Australian brands, 8% of the dog foods and
14% of the cat foods had vitamin A concentrations below the minimum72

V I TA M I N A■ Table 2.9

Recommendation for Periodic Administration of High Doses of Vitamin A
to Ruminants

AnimalCattleCalvesCattle (rearing)
Cattle (fattening)
Cattle (dairy)
Breeding bulls
Sheep
Suckling and weaned lambs
Rams
Ewes

IU per animal
200,000–500,000
1,000,000–1,500,000
1,000,000–2,000,000
2,000,000 (1 month before calving)
2,000,000–3,000,000 (2 months before
and at the beginning of service)
100,000–500,000
1,000,000 (2 months before and at the
beginning of service)
1,000,000 (during service and a month
before lambing)Source: Adapted from Hoffmann-La Roche (1967).

recommended for dogs and pregnant or lactating cats (Heanes, 1990).
In contrast, canned pet foods stated to contain liver or kidney showedfavorable vitamin A concentrations.
The importance of synthetic retinoids in veterinary dermatology is
now being explored. In dogs, the retinoids have been used primarily totreat skin diseases purported to be keratinization disorders. Various skindisorders, including sebaceous adenitis, canine ichthyosis, solar-induced
precancerous lesions, and squamous cell carcinoma have all responded
to retinoid therapy (Power and Ihrke, 1990; Power et al., 1992). In bothdogs and cats with mycosis fungoides, isotretinoin seems to be a welltolerated and relieving type of therapy. Cure is not always achieved, but
the length and quality of life in treated animals are improved. The recommended dosage of isotretinoin in dogs and cats is between 1 and 3
mg per kilogram per day. Isotretinoin is available by prescription as
Accutane in 10-, 20-, or 40-mg capsules and is sold prepackaged in 10capsule units. The recommended dosage of etretinate used for keratinization disorders in dogs is 0.75 to 1 mg per kilogram per day. Initial
response to retinoid therapy may not be apparent until 45 to 60 days oftreatment (Power and Ihrke, 1990).
The major source of supplemental vitamin A used in diets is transretinyl acetate (McGinnis, 1988). The acetate, propionate and palmitate
esters are chemically synthesized by vitamin A manufacturers. These areavailable in gelatin beadlet product forms for protection against oxidative destruction in premixes mash, and cubed and pelleted feeds. Carbo73

CHAPTER TWO

hydrates, gelatin, and antioxidants are also generally included inside thebeadlets to stabilize the vitamin A. These provide physical and chemical
protection against factors normally present in feed that are destructive
to vitamin A. The vitamin A acetate products most frequently used infeeds contain 650,000 IU or United States Pharmacopeia Units (USP) per
gram of product.
Additional research is needed to establish whether supplemental β−carotene helps reduce mastitis incidence and improve reproductive performance. Experiments conducted so far to determine if β−carotene
affects mastitis and/or reproduction are limited and conflicting.
Likewise, the effect of supplemental β−carotene on reproduction function in mares is inconclusive (Peltier et al., 1997). One reason for the
conflicting results is that the vitamin A and β−carotene status of livestock can vary greatly. Many feeds, especially green forages, containlarge amounts of β−carotene, and animals can store vast quantities ofvitamin A in the liver. Diets containing stored roughages such as wheat
straw, corn silage, and dried beet pulp are low in β−carotene(Bonsembiante et al., 1980). According to Adams and Zimmerman(1984), analysis of various feedstuffs from different regions of theUnited States showed that in the absence of fresh pasture or good quality forage, diets may not provide sufficient β−carotene. Analysis also
showed that samples of moldy corn averaged 98% less carotene thansound corn.
In industrialized countries, xerophthalmia due to primary vitamin A
deficiency is rarely seen because of the availability of adequate fruits,
vegetables, and animal food combined with fortified food such as margarine. Loss of animal products rich in vitamin A can quickly bringabout deficiency of the vitamin where previously there was no problem.
Many rural Sudanese have depended on milk from their livestock as
their main source of vitamin A. Droughts, however, have caused animal
losses, thus limiting vitamin A intake from milk products (Nestel et al.,1993).
The primary causes of hypovitaminosis A are ignorance, both natural and human-induced catastrophes, economic deprivation, and disease. Inexpensive foods that contain carotenoids and vitamin A are
nonetheless available in most areas where vitamin A deficiency is prevalent, but, for a variety of reasons, they are not consumed in adequate
amounts (Olson, 1994). Also, chronic alcohol consumption can reduce
retinoic acid concentrations and diminish retinoid signaling (Wang,
1999).
Elimination of vitamin A deficiency is a global priority. Periodic
74

V I TA M I N A

supplementation using a large dose of vitamin A is the most widely usedintervention in controlling deficiency. The current World HealthOrganization recommendation for universal prophylactic dosing inareas where vitamin A deficiency is a public health problem is 210 μmol
every 3 to 6 months for children 1 to 6 years of age (Humphrey et al.,1994).
In developing countries, both marginal vitamin A status and intestinal helminths (e.g., Ascaris lumbricoides) are common among children(Tanumihardjo et al., 1996). The combination of deworming and vitamin A supplementation in a single program should be considered.
For humans, a number of reports have illustrated the benefits ofsupplemental vitamin A for conditions not associated with night blindness and xerophthalmia (see Functions and Deficiency sections). The
results of a dozen field studies conducted in Brazil, Ghana, India,
Indonesia, Nepal, and elsewhere indicate that supplementing the diets ofchildren who are at risk of vitamin A deficiency can reduce deaths fromdiarrhea by 35 to 50%, and the vitamin can almost halve the number ofdeaths due to measles (UNICEF, 1998).
Supplemental retinoids have been successfully used as inhibitors ofcarcinogenesis and have important roles in optimizing immune response.
Numerous clinical studies have demonstrated beneficial effects ofretinoids on skin diseases such as acne, psoriasis, rosacea, ichthyoses,
keratodermas, and skin cancers and their precursors, as well as a reversal of the effects of photoaging (Futoryan and Gilchrest, 1994).
Retinoid-based creams applied to skin result in a younger appearance(e.g., reduced wrinkling). Retinoic acid has recently been shown as atherapy for emphysema because it regenerated lung alveoli in a rat study(Massaro and Massaro, 1997).

β-CAROTENE FUNCTION INDEPENDENTOF VITAMIN A
Carotenoids have been shown to have biological actions independent of vitamin A (Olson, 1989; Chew, 1995). Some animal studies indicate that certain carotenoids with antioxidant capacities, but without
vitamin A activity, can enhance many aspects of immune functions, canact directly as antimutagens and anticarcinogens, can protect against
radiation damage, and can block the damaging effects of photosensitizers. In animal models, β−carotene and canthaxanthin have protectedagainst UV-induced skin cancer as well as some chemically induced
tumors. In some of these models, enhancement of tumor immunity has
75CHAPTER TWO

been suggested as a possible mechanism of action of these carotenoids(Bendich, 1989). β−Carotene can function as a chain-breaking antioxidant; it deactivates reactive chemical species such as singlet oxygen,
triplet photochemical sensitizers, and free radicals, which would otherwise induce potentially harmful processes (e.g., lipid peroxidation).
Cows supplemented with 300 mg of β−carotene per day had a lower
incidence of intramammary infections than cows supplemented withpreformed vitamin A or the unsupplemented control group. It was also
demonstrated that dairy cows supplemented with 300 mg of β−carotene
per day during the dry period maintained blood concentrations of β−carotene and had a lower incidence of mastitis than those supplemented
with vitamin A alone (Chew and Johnston, 1985). β−Carotene enhances
blastogenic responses of lymphocytes and increases cytotoxic activities
of natural killer cells and cytokine production by macrophages (Chew,
1993).
Polymorphonuclear neutrophils (PMNs) are the major line ofdefense against bacteria in the mammary gland. β−Carotene supplementation seems to exert a stabilizing effect on PMN and lymphocyte function during the period around dry-off (Tjoelker et al., 1990). Daniel etal. (1991a,b) reported that β−carotene enhanced the bactericidal activity of blood and milk PMN against S. auereus but did not affect phagocytosis. Vitamin A either had no effect or suppressed bactericidal activity and phagocytosis. Control of free radicals is important for bactericidal activity but not for phagocytosis. The antioxidant activity of vitamin
A is not important; it does not quench or remove free radicals. β−
Carotene, on the other hand, has significant antioxidant properties andeffectively quenches singlet oxygen free radicals (Di Mascio et al., 1991;
Zamora et al., 1991).
A comprehensive review of the reproductive effects of carotene
and/or vitamin A in ruminants has been reported (Hoffmann-La Roche,1994). Since 1976, a number of studies have indicated that β−carotene
has a function independent of vitamin A in dairy cattle. Dairy cattle
receiving extra β−carotene have a higher intensity of estrus, increased
conception rates, and reduced frequency of follicular cysts than controls.
The corpus luteum of the cow has higher β−carotene concentrations
than any other organ, and it has been suggested that β−carotene has a
specific effect on reproduction in addition to its role as a precursor ofvitamin A. Graves-Hoagland et al. (1988) suggest a positive relationshipbetween β−carotene and luteal cell progesterone during the winter whenplasma β−carotene and vitamin A are decreased. Other studies with cattle have been reported in which pregnancy rates were lower with low
76

V I TA M I N A

dietary β−carotene but were adequate in vitamin A (Cooke andCombden, 1978; Bonsembiante et al., 1980). However, other studies ofcattle failed to detect differences between groups supplemented with β−carotene and control groups in the above-mentioned responses (Folman
et al., 1979; Wang et al., 1988; Aréchiga et al., 1998b). Aréchiga et al.
(1998a) reported that in cows fed β−carotene the pregnancy rate at 120
days postpartum was 14.3% higher, and milk yield was 6 to 11% higher than in controls.
Injectable β−carotene has increased conception rates in swine andimproved live births and live weights in pigs (Chew, 1993).
Contradictory results were obtained from studies that investigated theeffect of supplemental β−carotene on reproductive function in mares. β−
Carotene supplementation had a positive effect on the pregnancy rate ofmares in some studies (Ahlswede and Konermann, 1980; Peltier et al.,1997).
There are significant new views on the health benefits of megavitamin doses, particularly the antioxidant vitamins (C, E, and β−carotene).
Because damage to mammalian tissues induced by free radicals is
believed to contribute to the aging process and to the development ofsome degenerative diseases (Canfield et al., 1992), it has been proposedthat dietary carotenoids serve as antioxidants in tissues (Thurnham,
1994). This possibility is supported by numerous epidemiological studies that indicate an inverse association between the increased intake of
carotenoid-rich fruits and vegetables and the incidence of disease.
There has been much research regarding the potential protective
effects of carotenoids against chronic diseases (Cooper et al., 1999b).
Studies have shown inverse relationships between serum levels of one ormore carotenoids and a number of diseases, including cancer (Van
Poppel and Goldbohn, 1995), cardiovascular disease (Gaziano andHennekens, 1993; VERIS, 1996), eye diseases of age-related maculardegeneration (Snodderly, 1995), and nuclear sclerotic and cortical
cataracts (Mares-Perlman et al., 1995). In relation to cancer, in vitro
studies have demonstrated that carotenoids can inhibit chemicallyinduced neoplastic transformation (Bertram and Bortkiewicz, 1995),
induce remission of oral leukoplakia (Garewal, 1995), quench free radicals such as singlet oxygen (Sies and Stahl, 1995), and modulate
immune activity (Meydani et al., 1995). There is recent evidence thatsome of the carotenoid protection against cancer relates to gene regulation (Bertram, 1999).
Several hundred carotenoid research studies have been publishedsince 1996, when two major intervention trials showed a lack of pro77

CHAPTER TWO

tective effect of β-carotene supplements against lung cancer. Recent epidemiologic studies, however, continue to show an association between
high dietary intake of β-carotene and lower risk of lung cancer (Cooper,
1999a).
The predominant carotenoids in human tissues are β−carotene andlycopene. Compared with other carotenoids and other antioxidant compounds, including vitamin E, lycopene has been reported to be a more
efficient quencher of singlet oxygen in vitro (Di Mascio et al., 1991).
Lycopene disappearance precedes the disappearance of β−carotene whenhuman low-density lipoprotein is oxidized in vitro (Esterbauer et al.,1989). When skin is subjected to UV light stress, more skin lycopene isdestroyed compared with β−carotene, suggesting a role of lycopene in
mitigating oxidative damage in tissues (Ribaya-Mercado et al., 1995).
One study suggests that eating tomatoes, which are rich in lycopene, cansignificantly reduce the risk of heart attacks and prostate cancer.

TOXICITY
In general, the possibility of vitamin toxicities for livestock andpoultry is remote. However, of all vitamins, vitamin A is most likely tobe provided in toxic concentrations to both humans and livestock, andexcess vitamin A has been demonstrated to have toxic effects in most
species studied. Presumed upper safe levels are 4 to 10 times the nutritional requirements for nonruminant animals, including birds and fishes, and about 30 times the requirements for ruminants (NRC, 1987). A
high tolerance for ruminants may be due in part to some microbial
degradation of vitamin A in the rumen (Rode et al., 1990). Most of theharmful effects have been obtained by feeding over 100 times the dailyrequirements for a period of time. Thus, small excesses of vitamin A for
short periods should not exert any harmful effects. Recommended upper
safe levels of vitamin A for livestock and poultry are presented in Table
2.10 (NRC, 1987).
Large doses can be toxic, and many cases of overdoses have beenreported in various species. The most characteristic signs of hypervitaminosis A are skeletal malformations, spontaneous fractures, and internal hemorrhage (NRC, 1987). Other signs include loss of appetite, slow
growth, weight loss, skin thickening, suppressed keratinization,
increased blood-clotting time, reduced erythrocyte count, enteritis, congenital abnormalities, and conjunctivitis. Degenerative atrophy, fatty
infiltration, and reduced function of liver and kidney are also typical. A
report on clinical signs of hypervitaminosis A in rainbow trout (Salmo
78

V I TA M I N A■ Table 2.10

Presumed Safe Upper Levels of Vitamin A (IU/kg
Diet) for Livestock and Poultry

Animal
Birds
Chicken, growing
Chicken, laying
Duck
Goose
Quail
Turkey, growing
Turkey, breeding
Cat
Cattle, feedlot
Cattle, pregnant, lactating, or bulls
Dog
Fish
CatfishSalmon
Trout
Goat
Horse
MonkeyRabbitSheep
Swine, growing
Swine, breeding

Presumed
Safe Levela
15,00040,00040,00015,00025,00015,00024,000100,000
66,00066,00033,33033,33025,00025,00045,00016,000100,000
16,00045,00020,00040,000Source: Adapted from NRC (1987).
aFor chronic dietary administration.

gairdneri) listed growth depression; increased mortality; abnormal andnecrotic anal, caudal, pectoral, and pelvic fins; and pale yellow fragile
livers (Hilton, 1983). Bone abnormalities may include extensive bone
resorption and narrowing of the bone shaft, bone fragility, and shortbones because of retarded growth. Abnormalities in bone modeling are
the essential causes of fractures, and the cartilage matrix of bone may bedestroyed. For example, high levels of vitamin A injected into rabbits
may cause ears to curl because of the destruction of cartilage.
Pet foods or vitamin A medication can result in hypervitaminosis A.
The use of vitamin A in dermatology therapy of dogs and the inclusionof livestock by-products high in vitamin A in canned dog foods haveraised concerns about subclinical, long-term vitamin A toxicosis(Hoffmann-La Roche, 1998). Hypervitaminosis A in cats is most frequently associated with diets having a high liver or fish liver oil content(Goldman, 1992; Goldston and Hoskins, 1995). When a cat is found tobe sick, and its diet consists wholly or partly of raw liver, vitamin A toxicity should be suspected. The feeding of raw liver should be stopped
79

CHAPTER TWO

and a balanced feline diet instituted (Goldman, 1992). Cats consuming
raw beef liver containing 20,000 to 40,000 μg per kilogram for 2 to 5
years produced the main clinical findings of cutaneous hyperesthesia ofneck and forelegs, forelimb lameness, pain and skeletal immobility due
to bony exostoses of the cervical vertebrae, and tendinous tuberosities oflong bones (Seawright et al., 1965).
It is believed that release of lysosomal enzymes is responsible fordegradative changes observed in tissues and intact animals sufferingfrom hypervitaminosis A (Fell and Thomas, 1960). In hypervitaminosis
A, retinol penetrates the lipid of the membrane and causes it to expand,
and because the protein of the membrane is relatively inelastic, the membrane is therefore weakened. Thus, many phenomena in hypervitaminosis A can be explained in terms of damage to membranes either ofcells or of organelles within cells.
Excess vitamin A affects metabolism of other fat-soluble vitaminswith competition for absorption and transport demonstrated. Therefore,
in diets containing barely adequate levels of vitamins D, E, and K, amarked increase in dietary vitamin A may cause decreases in growth oregg production due to a deficiency of one or more of the other fat-soluble vitamins rather than a toxic effect of vitamin A. Retinoid-induced
hemorrhaging in rats fed diets deficient in vitamin K have been reported (McCarthy et al., 1989). The mechanisms of absorption and transport apparently are similar for the carotenoid pigments and the fat-soluble vitamins. A marked increase in dietary vitamin A has been shownto interfere with absorption of carotenoids, thereby resulting indecreased pigmentation for poultry.
High dietary levels of vitamin A have depressed vitamin E utilizationin most animals studied (Schelling et al., 1995; Franklin et al., 1998).
However, levels of vitamin A fed to dairy cattle were in excess byapproximately tenfold to highest levels being fed commercially(Schelling et al., 1995). In swine, high levels of injectable vitamin A had
little or no effect on serum and tissue vitamin E concentrations(Anderson et al., 1997).
The efficiency of carotene conversion to vitamin A declines progressively with increasing intakes (see Requirements). This appears to be anatural homeostatic control mechanism that protects grazing livestockfrom any harmful effects due to great abundance of carotene present inhigh-quality, fresh forages when they are the major feed for long periods(Miller, 1979). Likewise, comparatively rapid disposal of very high levels of stored vitamin A is a protective mechanism. On a practical basis,
toxicity is more easily caused by vitamin A than by carotene. Even so,80V I TA M I N A

vitamin A toxicity is not a practical problem for livestock, except when
unreasonably large amounts are given accidentally (Miller, 1979).
The symptomatology of hypervitaminosis A in humans varies considerably depending on the age of the subject and the duration of theexcessive intake. The young child is especially susceptible, with toxicity
occurring in children taking therapeutic doses prescribed for various
skin problems. The onset of clinical vitamin A toxicity is insidious andusually follows consumption of doses of the order of 100,000 IU per day
for periods ranging from weeks to months. The varied clinical signs andsymptoms are well documented and include headache and other symptoms attributable to increased intracranial pressure, dermatologicalchanges, pruritic skin rash, irritability, pain in arms and legs, and hydrocephalus in children. Plasma vitamin A is always elevated, usually well
above 200 μg per deciliter.
Arctic and Antarctic explorers have suffered from acute toxicity
after eating polar bear or seal liver. It has been calculated that consumption of 500 g of polar bear liver (13,000–18,000 IU of vitamin A/g)would result in a toxic dose (Sebrell and Harris, 1967). Symptoms ofthis acute polar bear liver poisoning, which may appear within 2 to 4
hours, are drowsiness, sluggishness, irritability or an irresistible desire to
sleep, severe headache, and vomiting. The medical literature records
death following intake of a single dose of 1,000,000 IU in adults and
500,000 IU in children.
Historically, it has been known for centuries among Eskimos andArctic travelers that the ingestion of polar bear liver by men and dogs
causes severe illness (Rodahl, 1949). There are stories of polar bear liver
being given to dogs, which later became sick and eventually sufferedfrom loss of hair. It is also said that the Eskimos threw the polar bear
livers into the sea to keep the dogs from getting them. On the other
hand, it is known that dogs are reluctant to eat polar bear liver, as is generally experienced by the European trappers in northeastern Greenlandwith their sledge dogs. On one occasion a liver was given to a very hungry dog, which ate a small portion of the liver and subsequently sufferedfrom diarrhea (Rodahl, 1949).
It is interesting that gulls and ravens avoid polar bear liver. On one
occasion ravens pecked a hole in the abdominal wall of a dead bear andate all the entrails but left the liver untouched. From the footsteps in thesnow around the carcass, it could be concluded that a large number ofbirds had been there (Rodahl, 1949).
Hypervitaminosis A in humans is becoming a clinical problem of
increasing frequency as a result of self-medication and overprescription.
81

CHAPTER TWO

According to McLaren (1982), vitamin A accumulation in human liverwith age signifies that hypervitaminosis A is also becoming a publichealth problem in Western countries. He points out that habitual intake
of vitamin A in the United States and Europe is greater than the recommended daily allowance, the body content of vitamin A is in excess ofthat of other vitamins in relationship to requirements, and the rate ofcatabolism of retinol is exponential, regardless of stores. Excess retinol
intake may explain the high incidence of osteoporosis in Northern
Europe (Whiting and Lemke, 1999). Explicit warnings for both children
and adults are provided by the RDA (1989) regarding excessive vitamin
A intakes of more than 25,000 IU on a regular basis.

■ REFERENCES
Adams, C.R., and Zimmerman, C.R. (1984). Feedstuffs 56, 35.
Ahlswede, L, and Konermann, H. (1980). Der praktische Tierarzt 61, 47.
Ahmed, F., Jones, D.B., and Jackson, A.A. (1990). Br. J. Nutr. 63, 363.
Akordor, F.Y., Stone, J.B., Walton, J.S., Leslie, K.E., and Buchanan-Smith, J.B.
(1986). J. Dairy Sci. 69, 2173.
Anderson, L.E., Myer, R.O., Brendemuhl, J.H., and McDowell, L.R. (1997).
Reprod. Nutr. Dev. 37, 213.
Anonymous (1977). Nutr. Rev. 35, 305.
Anonymous (1991). Nutr. Rev. 49, 243.
Anonymous (1993). Nutr. Rev. 51, 81.
Aoe, H.I., Masuda, T., Mimura, T., Saito, T., and Komo, A. (1968). Bull. Jpn.
Soc. Sci. Fish. 34, 959.
Aréchiga, C.F., Staples, C.R., McDowell, L.R., and Hansen, P.J. (1998a). J.
Dairy Sci. 81, 390.
Aréchiga, C.F., Vázquez-Flores, S., Ortíz, O., Hernández-Cerón, J., Porras, A.,
McDowell, L.R., and Hansen, P.J. (1998b). Theriogenology 50, 65.
Arykroyd, W.R. (1958). Fed. Proc. Fed. Am. Soc. Exp. Biol. 17, 103.
Bardos, L. (1991). Magyar-Allatorvosok-Lapja 46, 167.
Bartsch, R.D., Imes Jr., G.D., and Smit, J.P.J. (1975). Onderstepoort J. Vet. Res.
42, 43.
Barua, A.B. (1997). Nutr. Rev. 55, 259.
Bauernfeind, J.C. (1972). J. Agric. Food Chem. 20, 456.
Bauernfeind, J.C., and DeRitter, E. (1972). Feedstuffs 44 (36), 34.
Bebravicius, V., Medzevicius, A., and Medzevicius, A. (1987). Acta
Parasitologica Lituanica 22, 102.
Beeson, W.M. (1965). Fed. Proc. Fed. Am. Soc. Exp. Biol. 24, 924.
Bendich, A. (1989). J. Nutr. 119, 135.
Bertram, J.S. (1999). Nutr. Rev. 57, 182.
Bertram, J.S., and Bortkiewicz, H. (1995). Am J. Clin. Nutr. 62, 1327S.
Berzin, N., and Bauman, V.K. (1987). Br. J. Nutr. 57, 255.
82

V I TA M I N A

Bhandari, N., Bahl, R., Sazawal, S., and Bhan, M.K. (1997). J. Nutr. 127, 59.
Bierer, T.L., Merchen, N.R., and Erdman, J.W. (1995). J. Nutr. 125, 1569.
Blomhoff, R., Green, M.H., Green, J.B., Berg, T., and Norum, K.R. (1991).
Physiol. Rev. 71, 951.
Bonsembiante, M., Bittante, G., and Andrighetto, I. (1980). Zoot. Nutr. Anim.
6, 47.
Booth, A., Reid, M., and Clark, T. (1987). J. Am. Vet. Med. Assoc. 190, 1305.
Braude, R., Foot, A.S., Henry, K.M., Kon, S.K., Thompson, S.Y., and Mead,
T.H. (1941). Biochem J. 35, 693.
Brief, S., and Chew, B.P. (1985). J. Anim. Sci. 60, 998.
Brown, E.D., Chen, W., and Smith, J.C. (1976). J. Nutr. 106, 563.
Brubacher, G.B., and Weiser, H. (1985). Int. J. Vitam. Nutr. Res. 55, 5.
Bruns, N.J., and Webb, K.E. (1990). J. Anim. Sci. 68, 454.
Canfield, L.M., Foprage, J.W., and Valenzuela, J.G. (1992). Proc. Soc. Exp. Biol.
Med. 200, 260.
Chapman, H.L, Shirley, R.L., Palmer, A.Z., Haines, C.E., Carpenter, J.W., andCunha, T.J. (1964). J. Anim. Sci. 23, 669.
Chew, B.P. (1987). J. Dairy Sci. 70, 2732.
Chew, B.P. (1993). J. Dairy Sci. 76, 2804.
Chew, B.P. (1995). J. Nutr. 125, 1804S.
Chew, B.P., and Johnston, L.A. (1985). J. Dairy Sci. 68(Suppl. 1), 191(Abstr.).
Chew, B.P., Luedecke, L.O., and Holpuch, D.M. (1984). J. Dairy Sci. 67, 2566.
Chhabra, A., and Arora, S.P. (1987). Indian J. Dairy Sci. 40, 322.
Chhabra, A., Arora, S.P., and Kishan, J. (1980). Indian J. Anim. Sci. 50, 879.
Claeson, M., and Merson, M.H. (1990). Pediatr. Infect. Dis. J. 9, 345.
Clawitter, J.W., Trout, E., Burke, M.G., Areghi, S., and Roberts, R.M. (1990).
J.Biol. Chem. 265, 3248.
Coelho, M.B. (1991). Vitamin Stability in Premixes and Feeds: A Practical
Approach, p. 56. BASF Technical Symp., Bloomington, Minnesota.
Coffey, M.T., and Britt, J.H. (1993). J. Anim. Sci. 71, 1198.
Cooke, B.C., and Combden, N.A. (1978). Anim. Prod. 26, 356(Abstr.).
Cooper, D.A. (1999a). Nutr. Rev. 57, 133.
Cooper, D.A. (1999b). Nutr. Rev. 57, 201.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Dahlquist, S.P., and Chew, B.P. (1985). J. Dairy Sci. 68(Suppl. 1), 191(Abstr.).
Daniel, L.R., Chew, B.P., Tanaka, T.S., and Tjoelker, L.W. (1991). J. Dairy Sci.
74, 124.
Dash, S.K., and Mitchell, D.J. (1976). Anim. Nutr. Health Oct. 16-17.
Davis, C.Y., and Sell, J.L. (1983). J. Nutr. 113, 1914.
Davis, C.Y., and Sell, J.L. (1989). Poult. Sci. 68, 136.
Di Mascio, P., Murphy, M.E., and Sies, H. (1991). Am. J. Clin. Nutr. 53, 194S.
Donoghue, S., Donawick, W.J., and Kronfeld, D.S. (1983). J. Nutr. 113, 2197.
Eaton, H.D., Lucas, J.J., Neilson, S.W., and Helmboldt, C.F. (1970). J. Dairy Sci.
53, 1775.
Ensminger, M.E., and Olentine, C.G. (1978). Feeds and Nutrition. Ensminger,
Clovis, California.
Erdman, J.W., Thatcher, A.J., Hofmann, N.E., Lederman, J.D., Block, S.S., Lee,
C.M., and Mokady, S. (1998). J. Nutr. 128, 2013.
83

CHAPTER TWO

Esterbauer, H., Striegl, G., Puhl, H., and Rotheneder, M. (1989). Free Radic.
Res. Commun. 6, 67.
Farid, M.F.A., and Ghanem, Y.S. (1982). World Rev. Anim. Prod. 18, 75.
Fawzi, W.W., Herrera, M.G., Willett, W.C., Nestel, P., El Amin, A., Lipsitz, S.,
and Mohamed, K.A. (1994). Am J. Clin. Nutr. 59, 401.
Fawzi, W.W., Herrera, M.G., Willett, W.C., Nestel, P., El Amin, A., andMohamed, K.A. (1995). J. Nutr. 125, 1211.
Fell, H.B., and Thomas, L. (1960). J. Exp. Med. 111, 719.
Fichter, S.A., and Mitchell, G.E. (1997). J. Anim. Sci. 72(Suppl. 1), 266(Abstr.).
Flachowsky, G., Heidemann, B., Schlenzig, M., Wilk, H., and Hennig, A.
(1993). Z. Ernährung Swiss 32, 21.
Folman, Y., Ascarelli, F., Hertz, Z., Rosenberg, M., Davidson, M., and Halevi,
A. (1979). Br. J. Nutr. 41, 353.
Franklin, S.T., Sorenson, C.E., and Hammell, D.C. (1998). J. Dairy Sci. 81,
2623.
Funk, C. (1922). The Vitamins. Williams and Wilkins Co., Baltimore.
Futoryan, T., and Gilchrest, B.A. (1994). Nutr. Rev. 52, 299.
Garewal, H. (1995). Am J. Clin. Nutr. 62, 1401S.
Gaziano, J.M., and Hennekens, C.H. (1993). Ann. N.Y. Acad. Sci. 691, 148.
Gershoff, S.N., Andrus, S.B., Hegsted, D.M., and Lentini, E.A. (1957). Lab.
Invest. 6, 227.
Goldman, A.L. (1992). J. Am. Vet. Med. Assoc. 200(12), 1970.
Goldston, R.T., and Hoskins, J.D. (1995). In Geriatrics and Gerontology of the
Dog and Cat, p. 310. W.B. Saunders Co., Phildelphia.
Goodman, D.S. (1980). Fed. Proc. Fed. Am. Soc. Exp. Biol. 39, 2716.
Grace, M.L., and Bernhard, R.A. (1984). J. Dairy Sci. 67, 1646.
Graves-Hoagland, R.L., Hoaglund, T.A., and Woody, C.O. (1988). J. Dairy Sci.
71, 1058.
Green, M.H., Green, J.B., and Lewis, K.C. (1987). J. Nutr. 117, 694.
Greiwe-Crandell, K.M., Kronfeld, D.S., Gay, L., Sklan, D., Tiegs, W., andHarris, P.A. (1997). J. Anim. Sci. 75, 2684.
Guerin, H.B. (1981). J. Anim. Sci. 53, 758.
Guss, S.B. (1977). Management and Diseases of Dairy Goats. Dairy Goat
Journal Publ. Corp., Scottsdale, Arizona.
Halevy, O., Arazi, Y., Melamed, D., Friedman, A., and Sklan, D. (1994). J. Nutr.
124, 2139.
Hall, W.C., Damjanou, I., Nielsen, S.W., van der Heide, L., and Eaton, H.D.
(1980). Am. J. Vet. Res. 41, 586.
Harmon, B.G., Miller, E.R., Hoefer, J.A., Ullrey, D.E., and Leucke, R.W. (1963).
J. Nutr. 79, 263.
Harris Jr., B. (1975). Feedstuffs 47, 42.
Heanes, D.L. (1990). Aust. Vet. J. 67(8), 291.
Hentges, J.F., Grummer, R.H., Phillips, P.H., Bohstedt, G., and Sorensen, D.K.
(1952). J. Am. Vet. Med. Assoc. 120, 213.
Herrick, J.B. (1972). Vet. Med. 67, 906.
Hidiroglou, N., McDowell, L.R., and Boning, A. (1986). Int. J. Vitam. Nutr.
Res. 56, 339.
Hill, C.H. (1961). Poult. Sci. 40, 762.
84

V I TA M I N A

Hilton, J.W. (1983). J. Nutr. 113, 1737.
Hoffmann-La Roche (1967). Vitamin A, the Key-stone of Animal Nutrition.
Hoffmann-La Roche, Basel, Switzerland.
Hoffmann-La Roche (1994). In Vitamin Nutrition for Ruminants
RCD8775/894. Hoffmann-La Roche Inc., Nutley, New Jersey.
Hoffmann-La Roche (1998). In Vitamin Nutrition for Dogs and Cats.
Hoffmann-La Roche, Nutley, New Jersey (In Press).
Horst, R.L., Reinhardt, T.A., Goff, J.P., Nonnecke, B.J., Gambhir, V.K., Fiorella,
P.D., and Napoli, J.L. (1995). Biodhem. 34, 1203.
Hosokawa, H. (1989). The vitamin requirements of fingerling yellowtail Seriola
quinqueradiata. Ph.D. Dissertation, Kochi University, Japan.
Howell, C.E., Hart, G.H., and Ittner, N.R. (1941). Am. J. Vet. Res. 2, 60.
Howell, J., McC., and Thompson, J.N. (1967). Br. J. Nutr. 21, 741.
Humphrey, J.H., Natadisastra, G., Friedman, D.S., Tielsch, J.M., West, K.P., andSommer, A. (1994). J. Nutr. 124, 1172.
Hussein, L., and El-Tohamy, M. (1990). Int. J. Vitam. Nutr. Res. 60, 229.
Jensen, L.S. (1965). Poult. Sci. 44, 1609.
Johnson, E.J., Qin, J., Krinsky, N.I., and Russell, R.M. (1997). J. Nutr. 127,
1833.
Kasner, P., Chambon, P., and Leid, M. (1994). In Vitamin A in Health andDisease, p. 189 (R. Blomhoff, ed.). Marcel Dekker, New York.
Katz, J. Khatry, S.K., West, K.P., Humphrey, J.H., LeClerq, S.C., Pradhan, E.K.,
Pohkrel, R.P., and Sommer, A. (1995). J. Nutr. 125, 2122.
Kelley, K., and Easter, R. (1987). Feedstuffs 59(22), 14.
Kiatoko, M., McDowell, L.R., Bertrand, J.E., Chapman, H.L., Pate, F.M.,
Martin, F.G., and Conrad. J.H. (1982). J. Anim. Sci. 55, 28.
Kitamura, S., Suwa, T., Ohara, S., and Nakagawa, K. (1967). Bull. Jpn. Soc. Sci.
Fish. 33, 1126.
Kliewer, S.A., Umesono, K., Evans, R.M., and Mangelsdorf, D. (1994). In
Vitamin A in Health and Diseases, p. 239 (R. Blomhoff, ed.). Marcel
Dekker, New York.
Knopf, K., Sturman, J.A., Armstrong, M., and Hayes, K.C. (1978). J, Nutr. 108,
773.
Kohlmeier, R.H., and Burroughs, W. (1970). J. Anim. Sci. 30, 1012.
Krishnan, S., Bhuyan, U.N., Talwar, G.P., and Ramalingas Wami, R. (1974).
Immunology 27, 383.
Larkin, P.J., and Yates, R.J. (1964). E. Afr. Aric. For. J. 30, 11.
Leo, M.A., Lowe, N., and Lieber, C.S. (1987). J. Nutr. 117, 70.
Lindley, C.E., Brugman, H.H., Cunha, T.J., and Warwick, E.J. (1949). J. Anim.
Sci. 8, 590.
Liuzzi, J.P., Cioccia, A.M., and Hevia, P. (1998). J. Nutr. 128, 2467.
Loosli, J.K. (1991). In Animal Science Handbook (P.A. Putnam, ed). Academic
Press, San Diego.
Lopen, G.A., Phillips, R.W., and Nockels, C.F. (1973). Proc. Soc. Exp. Biol.
Med. 144, 54.
Mäenpää, P.H., Pirhonen, A., and Koskinen, E. (1988). J. Anim. Sci. 66, 1424.
Majumdar, B.N., and Gupta, B.N. (1960). Ind. J. Med. Res. 48, 388.
Mares-Perlman, J.A., Brady, W.E., Klein, B.E.K., Palta, M., Bowe, P., and85

CHAPTER TWO

Stacewicz-Sapuntzakis, M. (1995). Invest. Ophthalmol. Vis. Sci. 36, 276.
Martin, F.J., Ullrey, D.E., Miller, E.R., Kemp, K.E., Geasler,M.R., andHenderson, H.E. (1971). J. Anim. Sci. 32, 1233.
Massaro, G.D., and Massaro, D. (1997). Nat. Med. 3, 675.
Maynard, L.A., Loosli, J.K., Hintz, H.F., and Warner, R.G. (1979). Animal
Nutrition, p. 283, 7th Ed. McGraw-Hill, New York.
McCarthy, D.J., Lindamood, C., Gundberg, C.M., and Hill, D.L. (1989).
Toxicology and Applied Pharmacology 97, 300.
McCollum, E.V. (1957). In A History of Nutrition, p. 217. Houghton Mifflin,
Boston.
McDowell, K.J., Adams, M.H., Franklin, K.M., and Baker, C.B. (1995). Biol.
Reprod. 52, 438.
McDowell, L.R. (1985). Nutrition of Grazing Ruminants in Warm Climates.
Academic Press, San Diego.
McDowell, L.R. (1992). Minerals in Animals and Human Nutrition. Academic
Press, San Diego.
McDowell, L.R. (1997). Minerals for Grazing Ruminants in Tropical Regions,
3rd Ed. University of Florida, Gainesville.
McGhee, J.R., Mestecky, J., Dertzbaugh, M.T., Eldridge, J.H., Hirasawa, M.
and Kiyono, H. (1992). Vaccine 10, 75.
McGinnis Jr., C.H. (1988). In Proc. 1988 Georgia Nutrition Conference for the
Feed Industry, Athens, Georgia.
McLaren, D.S. (1982). Nutr. Rev. 40, 303.
McLaren, D.S. (1984). In Nutrition Reviews, Present Knowledge in Nutrition,
p. 192 (R.E. Olson, ed). Nutrition Foundation, Washington, D.C.
McLaren, D.S. (1986). In Vitamin A Deficiency and Its Control, p. 1 (J.C.
Bauernfeind, ed). Academic Press, Orlando, Florida.
Mellanby, E. (1938). J. Physiol. 94, 316.
Meydani, S.N., Wu, D., Santos, M.S., and Hayek, M.G. (1995). Am. J. Clin.
Nutr, 62, 1462S.
Michal, J.J., Heirman, L.R., Wong, T.S., and Chew, B.P. (1994). J. Dairy Sci. 77,
1408.
Miller, R.W., Hemken, R.W., Waldo, D.R., and Moore, L.A. (1969). J. Dairy Sci.
52, 1998.
Miller, W.J. (1979). Dairy Cattle Feeding and Nutrition. Academic Press, NewYork.
Milton, R.C. Reddy, V., and Naidu, A.N. (1987). Am. J. Clin. Nutr. 46, 827.
Mitchell, G.E. (1967). J. Am. Vet. Med. Assoc. 151, 430.
Moore, A.C., Gugger, E.T., and Erdman, J.W. (1996). J. Nutr. 126, 2904.
Morand-Fehr, P. (1981). In Goat Production, p. 193 (C. Gall, ed). Academic
Press, New York.
Nestel, P., Herrera, M.G., El Amin, A., Fawzi, W.W., Mohammed, K.A., andWeld, L. (1993). J. Nutr. 123, 2115.
Nockels, C.F. (1988). In Proc. 1988 Georgia Nutrition Conference for the Feed
Industry, Athens, Georgia.
Nonnecke, B.J., Reinhardt, T.A., and Franklin, S.T. (1993). J. Dairy Sci. 76,
2175.
NRC. Nutrient Requirements of Domestic Animals. National Academy of86

V I TA M I N A

Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of SciencesNational Research Council, Washington, D.C.
Oldham, E.R., Eberhart, R.J., and Muller, L.D. (1986). J. Dairy Sci. 69(Suppl.
1), 103.
Oldham, E.R., Eberhart, R.J., and Muller, L.D. (1991). J. Dairy Sci. 74, 3775.
Olson, J.A. (1989). J. Nutr. 119, 94.
Olson, J.A. (1991). In Handbook of Vitamins (L.J. Machlin, ed). Marcel
Dekker, Inc., New York.
Olson, J.A. (1994). J. Nutr. 124, 1461S.
Ong, D.E. (1993). J. Nutr. 123, 351.
Peltier, M.M., Peltier, M.R., Sharp, D.C., and Ott, E.A. (1997). Theriogenology
48, 893.
Perry, T.W. (1980). Beef Cattle Feeding and Nutrition, Academic Press, NewYork.
Perry, T.W., Beeson, W.M., Smith, W.H., and Mohler, M.T. (1967). J. Anim. Sci.
26, 115.
Pit, G.A.J. (1985). In Fat-Soluble Vitamins, p. 1. (A.T. Diplock, ed.). Technomic
Publ., Lancaster, Pennsylvania.
Poor, C.L., Bierer, T.L., Merchen, N.R., Fahey, G.C., and Erdman, J.W. (1993).
J. Nutr. 123, 1296.
Poston, H.A., Riis, R.C., Rumsey, G.L., and Ketola, H.G. (1977). Cornell Vet.
67, 472.
Power, H.T., and Ihrke, P.J. (1990). Vet. Clin. North Am. Small Anim. Pract.
20(6), 1525.
Power, H.T., Ihrke, P.J., Stannard, A.A., and Backus, K.Q. (1992). J. Am. Vet.
Med. Assoc. 201(3), 419.
Pusateri, A.E., Diekman, M.A., and Singleton, W.L. (1998). J. Anim. Sci. 77, 1532.
Quackenbush, F.W. (1963). Cereal Chem. 40, 266.
Rahmatullah, L., Underwood, B.A., Thulasiraj, R.D., and Milton, R.C. (1991).
Am. J. Clin. Nutr. 54, 568.
87

CHAPTER TWO

Raila, J., Eisenach, C., Buchholz, I., and Schweigert, F.J. (1997). J. Anim. Sci.
75(Suppl. 1), 226(Abstr.).
Rajaraman, V., Nonnecke, B.J., Franklin, S.T., Hammell, D.C., and Horst, R.L.
(1998). J. Dairy Sci. 81, 3278.
Ralston Purina. (1987). Nutrition and Management of Dogs and Cats. Ralston
Purina Co., St. Louis, Missouri.
Ray, S.M. (1963). Proc. World Conf. Anim. Prod. 1st Rome 2, 190(Abstr.).
RDA (1989). Recommended Dietary Allowances, 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
Ribaya-Mercado, J.D., Garmyn, M., Gilchrest, B.A., and Russell, R.M. (1995).
J. Nutr. 125, 1854.
Rodahl, K. (1949). Norsk Polarinstitutt Bull. No. 92. Oslo, Norway.
Rode, L.M., McAllister, T.A., and Cheng, K.J. (1990). Can. J. Anim. Sci. 70,
227.
Roels, O.A., Trout, M., and Dujacquier, R. (1958). J. Nutr. 65, 115.
Ross, A.C. (1992). Proc. Soc. Exp. Biol. Med. 200, 303.
Ross, A.C. (1993). J. Nutr. 123, 346.
Rumsey, T.S. (1975). Feedstuffs 47, 30.
Russell, W.C. (1929). J. Biol. Chem. 85, 289.
Safonova, I., Amri, E., and Ailhaud, G. (1994). Mol. Cell. Endocrinol. 104, 201.
Schelling, G.T., Roeder, R.A., Garber, M.J., and Pumfrey, W.M. (1995). J. Nutr.
125, 1799S.
Schmidt, H. (1941). Am. J. Vet. Res. 2, 373.
Schweigert, F.J., Vehlein-Harrell, S., and Zucker, H. (1990). J. Vet. Med. 37,
605.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Seawright, A.A., English, P.B., and Gartner, R.J.W. (1965). Nature 206, 1171.
Sebrell Jr., W.H., and Harris, R.S. (1967). The Vitamins, p. 3. Academic Press,
New York.
Seijas, H.C. (1995). In Ph.D. Dissertation, Ohio State University, Columbus.
Selke, M.R., Barnhart, C.E., and Chaney, C.H. (1967). J. Anim. Sci. 26, 759.
Semba, R.D. (1998). Nutr. Rev. 56, S38.
Serman, V., and Mazija, H. (1985). Veterinarski Archiv 55(1), 1.
Shields, R.G., Campbell, D.R., Hughes, D.M., and Dillingham, D.A. (1982).
Feedstuffs 22, 27.
Shin, D., and McGrane, M.M. (1997). J. Nutr. 127, 1274.
Sies, H., and Stahl, W. (1995). Am. J. Clin. Nutr. 62, 1322S.
Sijtsma, S.R., West, C.E., Rombout, J.H.W.M., and van der Zijpp, A.J. (1989).
J. Nutr. 119, 932.
Singh, M.M., Sinha, G.K., and Gupta, B.N. (1965). Ind. Vet. J. 42, 879.
Smith, J.E. (1990). Methods Enzymol. 190, 229.
Snodderly, D.M. (1995). Am. J. Clin. Nutr. 62(Suppl.), 1448S.
Sommer, A., Tarwotijo, I., Djunaedi, E., West, K.P., Loeden, A.A., Tilden, R.,
and Mele, L. (1986). Lancet 1 (9481), 1169.
Sommer, A., Tarwotijo, I., Hussani, G., Susanto, D., and Soegiharto, T. (1981).
Lancet 1 (8235), 1407.
Stahl, W., Schwarz, W., Von Laar, J., and Sies, H. (1995). J. Nutr. 125, 2128.
88

V I TA M I N A

Stephensen, C.B., Blount, S.R., Schoeb, T.R., and Park, J.Y. (1993). J. Nutr. 823,
833.
Stephensen, C.B., Moldoveanu, Z., and Gangopadhyay, N.N. (1996). J. Nutr.
126, 94.
Suryawan, A., and Hu, C.Y. (1997). J. Anim. Sci. 75, 112.
Takyi, E.E.K. (1999). J. Nutr. 129, 1549.
Tanumihardjo, S.A., Permaesih, D., Rustan, E., Rusmil, K., Fatah, A.C., Wilbur,
S., Karyadi, D., and Olson, J.A. (1996). J. Nutr. 126, 451.
Thurnham, D.I. (1994). Proc. Nutr. Soc. 53, 77.
Tjoelker, L.W., Chew, B.P., Tanaka, T.S., and Daniel, L.R. (1990). J. Dairy Sci.
73, 1017.
Tokach, M.D., Goodband, R.D., and Nelssen, J.L. (1994). In Kansas State
University Swine Day, Kansas State University, Manhattan, Kansas.
Trout, W.E., Hall, J.A., Stallings-Mann, M.L., Galvin, J.M., Anthony, R.V., andRoberts, R.M. (1992). Endocrinology 130, 2557.
Ullrey, D.E. (1972). J. Anim. Sci. 35, 648.
Underwood, B.A. (1990). J. Nutr. 120, 1459.
Underwood, B.A. (1994). J. Nutr. 124, 1467S.
UNICEF. (1988). Nutr. Rev. 56, 115.
United Nations Administrative Committee on Coordination-Subcommittee on
Nutrition. (1987). First Report on the World Nutrition Situation, p. 36.
United Nations, Geneva.
Vallet, J.L. (1994). J. Anim. Sci. 72, 2449.
Vallet, J.L. (1995). Biol. Reprod. 53, 1436.
Van den Berg, H. (1999). Nutr. Rev. 57, 1.
Van der Pol, F., Purnomo, S.U., and Van Rosmalen, H.A. (1988). Nutr. Rep. Int.
37, 785.
Van Pelt, A.M.M., and De Rooij, D.G. (1990). Biol. Reprod. 43, 363.
Van Poppel, G., and Goldbohn, R.A. (1995). Am. J. Clin. Nutr. 62, 1393S.
Van Vliet, T., Van Vlissingen, M.F., Van Schaik, F., and Van den Berg, H. (1996).
J. Nutr. 126, 499.
VERIS. (1996). In VERIS Research Summary, November. VERIS ResearchInformation Service, La Grange, Illinois.
Wald, G. (1968). Science 162, 230.
Wang, J.Y., Owen, F.G., and Larson, L.L. (1988). J. Dairy Sci. 71, 181.
Wang, X. (1999). Nutr. Rev. 57, 51.
Weiss, W.P. (1998). J. Dairy Sci. 81, 2493.
West, C.E., Sijtsma, S.R., Kouwenhoven, B., Rombout, J.H.W.M., and van derZijpp, A.J. (1992). J. Nutr. 122, 333.
West, K.P., LeClerq, S.C., Shrestha, S.R., Wu, L.S., Pradhan, E.K., Khatry, S.K.,
Katz, J., Adhikari, R., and Sommer, A. (1997). J. Nutr. 127, 1957.
West, K.P., Pokhrel, R.P., Katz, J., LeClerq, S.C., Khatry, S.K., Shrestha, S.R.,
Pradhan, E.K., Tielsch, J.M., Pandey, M.R., and Sommer, A. (1991). Lancet
338 (8759), 67.
Whaley, S.L., Hedgpeth, V.S., and Britt, J.H. (1997). J. Anim. Sci. 75, 1071.
White, W.S., Peck, K.M., Ulman, E.A., and Erdman, J.W. (1993). J. Nutr. 123,
1129.
Whiting, S.J., and Lemke, B. (1999). Nutr. Rev. 57, 192.
89

CHAPTER TWO

Wiedermann, U., Hanson, L.A., Holmgren, J., Kahu, H., and Dahlgren, U.I.
(1993). Infect. Immun. 61, 3952.
Wing, J.M. (1969). J. Dairy Sci. 52, 479.
Wolf, G. (1991). Nutr. Rev. 49(1), 1.
Wolf, G. (1992). Nutr. Rev. 50, 292.
Wolf, G. (1995). Nutr. Rev. 53(5), 134.
Wolf, G. (1998). Nutr. Rev. 56, 156.
Zamora, R., Hidalgo, F.J., and Tappel, A.L. (1991). J. Nutr. 121, 30.
Zhao, Z., and Ross, C. (1995). J. Nutr. 125, 2064.

90

chapter three

V I TA M I N D

INTRODUCTION
Vitamin D is thought of as the “sunshine vitamin” because it is synthesized by various materials when they are exposed to sufficient sunlight. The two major natural sources of vitamin D are cholecalciferol(vitamin D3, which occurs in animals) and ergocalciferol (vitamin D2,
which occurs predominantly in plants). In this chapter, the term vitamin
D in the absence of a subscript will imply either vitamin D2 or vitamin
D3. Under modern farming conditions, many animals, particularly swine
and poultry, are raised in total confinement with little or no exposure tonatural sunlight. Even though with adequate sunlight exposure, vitamin
D is not needed in the diet, it still fits the definition of a vitamin in all
respects for animals and humans who are confined indoors away fromthe sun. In recent years vitamin D receptors have been found in tissues
not associated with the traditional role of calcium (Ca) metabolism. The
additional roles of vitamin D await further elucidation.

HISTORY
Historical aspects of vitamin D have been reviewed by DeLuca(1979), Holick (1987), Loosli (1991), and Collins and Norman (1991).
Vitamin D-deficiency rickets is a disease known since antiquity. There isevidence that rickets occurred in Neanderthal man about 50,000 B.C. In
525 B.C., there was a battle between Persia and Egypt, and 100 years
later, the battlefield was visited by the Greek historian Herodotus.
Herodotus noted that Persian skulls were so fragile they broke whenstruck by a pebble, while Egyptian skulls were strong. The difference
related to sunlight and vitamin D activity; Egyptians went bareheaded91

3CHAPTER THREE

from childhood while Persians covered their heads with turbans.
Hippocrates described conditions that resembled rickets, and Soranus
Ephesius (born A.D. 130) provided a classic description referring to “thebackbone bending” and “legs twisted at the thighs” in a disease noted
to be more common in smoky cities than in the country (Arneil, 1975).
Since the Middle Ages, it was observed that sunlight seemed to have
health-giving effects. During the early stages of the Industrial Revolution
in the late eighteenth and early nineteenth centuries, there was a massmigration of population from the rural countryside into the industrial
centers of Great Britain and Europe. Young children of the working
class who lived in the densely populated cities were affected by a severebone-deforming disease. The combination of industrial pollution andnarrow, shaded alleyways prevented children who were reared in thisenvironment from being exposed to sunlight. When necropsy studies
were performed on children who died of various causes, it was found in
Leyden, The Netherlands, that 85 to 90% of these children sufferedfrom rickets (Holick, 1987).
In 1822 Sniadecki suggested that rickets was caused by lack of
exposure to sunlight. He observed that children who lived in inner cities
of Warsaw, Poland, had a very high incidence of rickets, whereas children living on the farms on the outskirts of the city essentially were freeof this disease. Almost 70 years later Palm concluded from an epidemiological survey that the common denominator in rickets in children waslack of exposure to sunlight. He encouraged systematic sunbathing as ameans of preventing and curing rickets. The majority of scientists andphysicians at the time did not believe that simple exposure to sunlight
could cure or prevent this bone-deforming disease.
In the early twentieth century, Sir Edward Mellanby began his workon rickets, which was in epidemic proportions in the human populationin his native England. Mellanby attempted to produce rickets by nutritional means, and in 1922 he succeeded in producing the disease in dogs
maintained on a diet of oatmeal and, unplanned by him, in absence ofsunlight. However, we now know sunlight would not have been a factor
since vitamin D is not produced in the skin of this species (How et al.,1994a). Although Mellanby incorrectly concluded that the healing of
rickets was a property of the fat-soluble vitamin A, he placed the studyof rickets on an experimental basis.
McCollum, who had discovered the fat-soluble vitamin A, realizedthat the antirachitic activity discovered by Mellanby was distinct fromthe antixerophthalmia activity in cod liver oil. McCollum bubbled oxygen through cod liver oil and heated it to destroy the vitamin A activity,92

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but the properties of cod liver oil in prevention and cure of rickets
remained. Therefore, in 1922 he concluded that this unknown substance
represented a new fat-soluble vitamin, which he called vitamin D.
Although it was known that ultraviolet (UV) light and vitamin D
from cod liver oil were both equally effective in preventing and curing
rickets, the close interdependence of these two factors was not immediately realized. Goldblatt and Soames (1923) conducted experiments inwhich they irradiated rachitic rats with UV light, removed their livers,and demonstrated that they contained the antirachitic substance, whereas if they were not irradiated, no antirachitic substance could be detected. Steenbock and Black (1924) then realized that UV irradiation wascausing the alteration of some substance in animals and proceeded todemonstrate that UV irradiation of not only the animals but of theirfood could heal or prevent rickets.
These important discoveries led to the use of UV light irradiation offoods such as milk and butter to fortify them with vitamin D and thus
eliminate rickets as a major medical problem. In addition, the demonstration that irradiation of food resulted in the production of an antirachitic factor provided the key for the isolation and chemical characterization of vitamin D2 from the provitamin ergosterol, and that irradiation of skin produced vitamin D3 from the provitamin 7-dehydrocholesterol. In 1932, the structure of vitamin D2 was simultaneously determined by Windaus in Germany, who named it vitamin D2, and by
Askew in England, who named it ergocalciferol. In 1936, Windaus succeeded in identifying the structure of vitamin D3. Vitamin D1 had been
isolated earlier by Windaus and his colleagues but was later shown to be
a combination of vitamin D2 and an irradiation side product, lumisterol.
A new phase of vitamin D research began in the late 1960s and has
resulted in a complete change in attitude regarding the problems associated with this substance. It is now recognized that vitamin D is simply
the precursor of a steroid hormone, 1,25-dihydroxyvitamin D [1,25(OH)2D], sometimes referred to as calcitrol. In 1966, DeLuca (1979)
demonstrated the disappearance of vitamin D3 following administrationand the appearance of several metabolites that possessed more potent
antirachitic activity. The first of these metabolites, 25-hydroxyvitaminD(25-OHD), was found to be produced in the liver. The 25-OHD formof vitamin D was chemically synthesized by Blunt and coworkers in
1968. Using the radioactively labeled chemical, 25-OHD was shown to
be metabolically altered to at least three dihydroxy compounds, themost important thought to be 1,25-(OH)2D. In the early 1970s, it wasdetermined that the kidney was the principal site of 1,25-(OH)2D pro93

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duction. From the discoveries that this biologically active vitamin D
metabolite is produced exclusively in kidney and is found in the nuclei
of intestinal cells came the concept that, in terms of its structure andmode of action, vitamin D is similar to steroid hormones. The discovery
that the biological actions of vitamin D can be explained by a hormonelike mechanism of action marked the beginning of the modern era ofvitamin D research.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Vitamin D designates a group of closely related compounds thatpossess antirachitic activity. It may be supplied through the diet or byirradiation of the body. There are about 10 provitamins that, after irradiation, form compounds having variable antirachitic activity. The twomost prominent members of this group are ergocalciferol (vitamin D2)
and cholecalciferol (vitamin D3). Chemical structures of ergocalciferol
and cholecalciferol and their precursors, ergosterol and 7-dehydrocholesterol, are shown in Fig. 3.1. All sterols possessing vitamin D activity
have the same steroid nucleus; they differ only in the nature of the side
chain attached to carbon 17.
Ergocalciferol is derived from a common plant steroid, ergosterol,
and is the usual dietary source of vitamin D. Cholecalciferol is produced
exclusively from animal products. 7-Dehydrocholesterol is derived fromcholesterol or squalene, which is synthesized in the body and present inlarge amounts in skin, intestinal wall, and other tissues. Vitamin D precursors have no antirachitic activity until the B-ring is opened between
the 9 and 10 positions by irradiation and a double bond is formed
between carbons 10 and 19 to form vitamin D.
Vitamin D in the pure form occurs as colorless crystals that are
insoluble in water but readily soluble in alcohol and other organic solvents. It is less soluble in vegetable oils. Cholecalciferol crystallizes asfine white needles from diluted acetone and has a melting point between
84 and 85oC. Vitamin D can be destroyed by overtreatment with UVlight and by peroxidation in the presence of rancidifying polyunsaturated fatty acids. Like vitamins A and E, unless vitamin D3 is stabilized it isdestroyed by oxidation. Its oxidative destruction is increased by heat,moisture, and trace minerals. There is less destruction of vitamin D3 infreeze-dried fish meals during drying, possibly because of decreased
atmospheric oxygen (Scott and Latshaw, 1994). There is negligible lossof crystalline cholecalciferol over 1 year of storage in amber evacuated94

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Fig. 3.1 Vitamin D2 (ergocalciferol), vitamin D3 (cholecalciferol), and their precursors in animal (7-dehydrocholesterol) and plant (ergosterol) tissues.

capsules at refrigerator temperatures or of ergocalciferol for 9 months.
Solutions in corn oil stored in amber bottles have shown no loss of activity during 30 months of storage. Overirradiation of ergocalciferol orcholecalciferol produces numerous irradiation products such as tachysterols, supra-sterol1, supra-sterol2, and others. Some of these compounds have partial vitamin D activity, some are toxic, and some may
be potent antagonists of vitamin D (Scott et al., 1982).

ANALYTICAL PROCEDURES
Vitamin D activity can be expressed in units based on a bioassay
in vitamin D-deficient rats or chicks. When assayed in the rachitic rat,
ergocalciferol and cholecalciferol are equally active with a potency of
40,000 units/mg of pure steroid. However, in chicks and other birds,
ergocalciferol has only about one-tenth the activity of cholecalciferol(Chen and Bosmann, 1964). One international or USP unit of vitamin
D activity is defined as the activity of 0.025 μg of vitamin D3 contained in the USP vitamin D reference standard. For poultry, the terminternational chick unit (ICU) is employed with reference to use of D3
versus D2.
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Methods of analysis for vitamin D are complex and somewhat difficult because there are so many isomers, and not all are biologically
active. The standard method for assay of vitamin D supplements forfeeds is a biological assay. Vitamin D is the only vitamin in which a biological method has not been largely replaced by a chemical, physical, or
microbiological assay. The rat is often used to assay products for humanand animal use, but the chick is the assay animal of choice in assessing
supplements intended for poultry feeding because of unequal activity ofvitamin D2 and D3 for this species compared with most mammals.
Vitamin D3 biological assay using young chicks has been standardized
by the Association of Analytical Official Chemists. It involves feeding astandard rachitogenic ration to young chicks for a 21-day period anddetermining the ash content of dry fat-free tibia, either from chicks thathave received the supplements containing unknown quantities of vitamin D or from those receiving standard quantities of a vitamin D reference standard (Scott et al., 1982).
A curative method involves developing rickets in young rats andchicks; then graded increments of unknown samples and standard vitamin D are added to diets for 7 days, followed by a “line test.” This test
entails staining a section of the metaphysis of the proximal end of thetibia with silver nitrate (AgNO3) to show deposition of Ca salts. The silver precipitates the PO4 and, on exposure to light, Ag3PO4 is reduced toAg, which deposits in a black line. Bones are then graded numericallyaccording to degree of calcification. Also, x-rays may be taken insteadof using the line test. Chemical and physical methods to analyze vitamin
D generally lack sensitivity of biological assays. Thus, they are not adequate for measuring samples that contain low concentrations of vitamin
D. However, these physical and chemical means of vitamin D determination have the advantage of being less time-consuming than biologicalassays and are frequently used on samples known to contain high levelsof vitamin D (Collins and Norman, 1991).
Physical and chemical methods of vitamin D analysis include UV
absorption, colorimetric procedures, fluorescence spectroscopy, gas
chromatography/mass spectroscopy, competitive binding assays, andhigh-pressure liquid chromatography (HPLC). The HPLC procedure isvery promising, with the separation process resulting in an exceedingly
high resolving capability and increased sensitivity (Collins and Norman,
1991; Matilla et al., 1995). A major advantage of HPLC is that compounds are not altered by the heat of gas-liquid chromatography, so they
may be detected as the actual known compounds. Other benefits ofusing HPLC are the reduced labor and time required to separate vitamin96

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D and its metabolites. Compounds may be identified by the retention
time of either an internal or external standard and a new technique ofstop flow in which the UV spectrum of the molecule separated may be
examined.

METABOLISM
Absorption and Conversion from Precursors
Vitamin D obtained from the diet is absorbed from the intestinal
tract, with conflicting reports as to which portion of the small intestine
serves as the primary absorption site. It has also been suggested that thelargest amount of dietary vitamin D is more likely to be absorbed in theileum because of longer retention time of food in the distal portion of
the intestine (Norman and DeLuca, 1963).
Vitamin D is absorbed from the intestinal tract in association withfats, as are all the fat-soluble vitamins. Like the others, it requires thepresence of bile salts for absorption. Because it is fat soluble, vitamin D
is absorbed with other neutral lipids via chylomicra into the lymphatic
system of mammals or the portal circulation of birds and fishes. It has
been reported that only 50% of a dose of vitamin D is absorbed.
However, considering that sufficient vitamin D is usually produced bydaily exposure to sunlight, it is not surprising that the body has not
evolved a more efficient mechanism for dietary vitamin D absorption(Collins and Norman, 1991).
Cholecalciferol is produced by irradiation of 7-dehydrocholesterol
with UV light either from the sun or from an artificial source.
Cholecalciferol is synthesized in the outer skin layers. Presence of theprovitamin 7-dehydrocholesterol in the epidermis of the skin and sebaceous secretions is well recognized. In poultry, Tian et al. (1994) reported that in the chicken, the skin of legs and feet contains about 30 times
as much 7-dehydrocholesterol (provitamin D3) as the body skin.
Vitamin D is synthesized in the skin of many herbivores and omnivores, including humans, rats, pigs, horses, poultry, sheep, and cattle.
However, little 7-dehydrocholesterol is found in the skin of cats anddogs (and likely other carnivores), and therefore little vitamin D is produced in the skin (How et al., 1995).
During exposure to sunlight, the high-energy UV photons (290–315nm) penetrate the epidermis and photolyze 7-dehydrocholesterol (provitamin D3) to previtamin D3. Once formed, previtamin D3 undergoes athermally induced isomerization to vitamin D3 that takes 2 to 3 days to97

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reach completion. In poultry, the time course revealed a fourfold
increase in the circulating concentration of vitamin D3, with a peak
about 30 hours postradiation (Tian et al., 1994). Approximately 15%of provitamin D3 in human skin exposed to 10 minutes of simulated
sunlight is converted in the stratum basale to vitamin D3 (Holick et al.,1981). Longer exposure times do not significantly increase D3 concentrations in the epidermis. Heuser and Norris (1929) showed that 11 to
45 minutes of sunshine daily were sufficient to prevent rickets in growing chicks, and that no further improvements in growth were obtained
under these conditions by adding cod liver oil. During initial exposure
to sunlight, provitamin D3 in the human epidermis is efficiently converted to previtamin D3. However, because previtamin D3 is also labile
to sunlight, once it is formed, it begins to photolyze to additional photoproducts, principally luminsterol and tachysterol. The net result is that
prolonged exposure to sunlight does not significantly increase the previtamin D3 concentration above about 15% of the initial provitamin D3
concentrations (Holick, 1987).
More than 90% of previtamin D3 synthesis in skin occurs in theepidermis. The cholecalciferol formed by irradiation of the 7-dehydrocholesterol in the skin is absorbed through the skin and transported by
the blood to the lipids throughout the body. Clearly, absorption can takeplace, because rickets can be successfully treated by rubbing cod liver oilon the skin. Once vitamin D3 is formed, it is transported in the blood.
Some of the vitamin D3 formed in and on the skin ends up in the digestive tract as many animals consume the vitamin as they lick their skinand hair.

Conversion to Metabolically Active Forms
Prior to 1968, it was almost universally thought that cholecalciferol and ergocalciferol (D3 and D2) were the circulating antirachitic agentsin the living animal system. Starting in 1968, DeLuca and a number ofother researchers demonstrated that vitamin D undergoes a multiple
series of transformations and multi-site interactions in the living system(DeLuca, 1979). Whether the vitamin is ingested orally or produced
photochemically in the skin, these chemical changes occur before any
significant biological response is registered in the intestine or bone.
Production and metabolism of the vitamin D necessary to activate thetarget organs are illustrated in Fig. 3.2.
Once vitamin D (D2, D3, or both) enters the blood, it circulates atrelatively low concentrations. This phenomenon is probably a result of
its rapid accumulation in the liver. Once in the liver, the first transfor98

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Fig. 3.2 The functional metabolism of vitamin D3 necessary to activate the target organs of intestine, bone, and kidney.

mation occurs, in which a microsomal system hydroxylates the 25-carbon in the side chain to produce 25-OH vitamin D. This metabolite is
the major circulating form of vitamin D under normal conditions andduring vitamin D excess (Littledike and Horst, 1982).
Conversion to 25-OHD takes place predominantly in the microsomes but also in the mitochondria. The mitochondrial enzyme is ahigh-capacity, low-affinity enzyme and is thought to hydroxylate vitamin D under conditions of high substrate concentrations, such as vitamin D toxicity. Liver is the major site of 25-hydroxylation of vitamin D;
however, the intestine and kidney can also produce 25-OHD, although
the amount of 25-hydroxylation taking place in these organs is small(Tucker et al., 1973). In the chicken, the D-25-hydroxylase enzyme
exists in the extrahepatic tissues, including the intestine and kidney, but
in mammals, the liver is the predominant site.
The 25-OHD is then transported to the kidney on the vitamin D
transport globulin, where it can be converted to a variety of compounds,of which the most important appears to be 1,25-(OH)2D3 (also called99

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calcitrol) (DeLuca, 1990). This reaction occurs in the mitochondrial
fraction and is catalyzed by a three-component, mixed-function
monoxygenase involving NADPH, molecular oxygen, a flavoprotein, aniron-sulfur protein, and cytochrome P-450 (Ghazarian et al., 1974). The
1,25-(OH)2D that is formed in the kidney is then transported to theintestine, bones, or elsewhere in the kidney, where it is involved in themetabolism of Ca and phosphorus (P). The hormonal form is the metabolically active form of the vitamin that functions in intestine and bone,whereas 25-OHD and vitamin D do not function at these specific sites
under physiological conditions (DeLuca, 1990).
Production of 1,25-(OH)2D is very carefully regulated by parathyroid hormone in response to serum Ca and phosphate concentrations.
Removal of parathyroid glands results in an animal’s inability to adapt
to varying Ca demands by increasing intestinal Ca absorption. It is now
known that the most important point of regulation of the vitamin D
endocrine system occurs through the stringent control of the activity of
the renal 1-hydroxylase. In this way, the production of the hormone
1,25-(OH)2D can be modulated according to the Ca needs of the organism (Collins and Norman, 1991). Factors known to affect the activity of
the 1-hydroxylase include Ca, parathyroid hormone, and vitamin D status. Both intestinal and serum 1,25-(OH)2D concentrations and activityof 1-hydroxylase are inversely related to dietary and serum Ca concentrations.
Later, it was found that the hormone 1,25-(OH)2D is produced by
its endocrine gland, the kidney, and a paracrine system (Norman, 1995).
The 1-hydroxylase activities have been confirmed in placenta and in cultured cell lines (skin, bone, embryonic intestine).
Epidermal cells (keratinocytes) produce vitamin D; metabolize it to
its most biologically active form, 1,25-(OH)2D; and in response to the
1,25-(OH)2D, there is a decrease in proliferation and an increase in differentiation (Bikle, 1995). 1,25-(OH)2D production by keratinocytes istightly controlled and changes as the cells differentiate, increasing during the early stages of differentiation and then decreasing again as terminal differentiation ensues.
Besides 1,25-(OH)2D, the kidney also converts 25-OHD to otherknown compounds, including 24,25-(OH)2D, 25-26-(OH)2D, and
1,24,25-(OH)2D. The role of these compounds in function or inactivation of vitamin D has not been fully evaluated, and significant physiological roles are yet to be discovered. Although controversy exists, it has
been suggested that 24,25-(OH)2D is responsible for the mineralizationof bone and for the suppression of parathyroid hormone secretion.
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Bordier et al. (1978) showed that 24,25-(OH)2D3 is required with 1,25(OH)2D3 for normal healing of osteomalacia in humans. Dogs with
impaired renal function had depressed serum concentration of 24,25(OH)2D3, but no clinical benefit was observed when this form of vitamin D was administered (Dzanis et al., 1992). The hormonal role of
24,25-(OH)2D is very tentative. Although it has been shown that 24hydroxylation appears to have a role in bone mineralization, it is also
believed to be involved in the elimination pathway for 25-hydroxy and1,25-dihydroxy vitamin D.
Henry and Norman (1978) have shown that hatchability of eggs
from hens is severely depressed in vitamin D-deficient hens even when
they are fed 1,25-(OH)2D3. Evidently, 1,25-(OH)2D3 is effective inmaintaining blood Ca levels so that egg production and egg shell thickness remain normal. However, without vitamin D, the upper mandibleof the chicks fails to develop, and consequently, the chicks cannot crack
the shell, resulting in mortality. The suggestion was made that chickmortality resulted from lack of 24,25-(OH)2D. However, the reason forthe chick deaths related to failure of 1,25-(OH)2D3 being passed fromthe hen to the egg, as D3 and 25-OHD3 are readily passed to the egg.

Transport
Vitamin D in plasma, coming from either diet or the skin, is pickedup for transport to the liver by a plasma protein called vitamin D-binding protein (DBP; also called transcalciferin), which is synthesized in theliver. In mammals, vitamin D, 25-OHD, and possibly 24,25-(OH)2D
and 1,25-(OH)2D, are all transported by DBP. This protein has a molecular weight of 50,000 to 60,000 and in humans appears to be a singlechain polypeptide. The DBP binds 25-OHD (the major circulating
metabolite) with a higher affinity than it binds vitamin D or 1,25(OH)2D3 (Haddad and Walgate, 1976). With radioactively labeled
metabolites, Hay and Watson (1976) observed in 65 of the 72 species ofmammals studied that the vitamin D metabolites associated with a protein of α-globulin mobility as determined by disk gel electrophoresis.
This was also observed in bony fish, reptiles, and some species of birds.
However, for several mammalian species and in four species of birds, 25OHD was carried on albumin or on a protein with albumin-like electrophoretic mobility.

Storage and Placental Transfer
In contrast to aquatic species, which store significant amounts ofvitamin D in liver, land animals and humans do not store appreciable
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amounts of the vitamin. The body has some ability to store the vitamin,
although to a much lesser extent than is the case for vitamin A. Principal
stores of vitamin D occur in blood and liver, but it is also found in lungs,
kidneys, and elsewhere. Since it was known that liver serves as a storagesite for vitamin A, another fat-soluble vitamin, it was thought that theliver also functioned as a storage site for vitamin D. However, it has
since been shown that blood has the highest concentration of vitamin Dcompared with other tissues; in pigs, the amount of vitamin D in bloodis several times higher than that in liver (Quaterman et al., 1964).
The persistence of vitamin D in animals during periods of vitamin
D deficiency may be explained by slow turnover rate of vitamin D in
certain tissues, such as skin and adipose tissue. During times of deprivation, vitamin D in these tissues is released slowly, thus meeting vitamin D needs of the animal over a longer period (Collins and Norman,
1991).
Transplacental movement of Ca increases dramatically during thelast trimester of gestation in all species observed. It is well establishedthat in most mammalian species, fetal plasma Ca levels are higher thanmaternal levels at term. In sheep, passive diffusion accounts for aminor component of placental Ca movement, with active transportmechanisms responsible for more than 90% (Braithwaite et al., 1972).
In the pregnant rat (and perhaps humans), 1,25-(OH)2D is a criticalfactor in the maintenance of sufficient maternal Ca for transport to thefetus and may play a role in normal skeletal development of theneonate (Lester, 1986). Although there is no large transfer to the fetus,
liberal intake during gestation provides a sufficient store in newborns
to help prevent early rickets. For example, newborn lambs can be provided enough in this way to meet their needs for 6 weeks. Parenteralcholecalciferol treatment of sows before parturition proved an effective means of supplementing young piglets with cholecalciferol (via thesow’s milk) and its more polar metabolites via placental transport(Goff et al., 1984).

Excretion
Excretion of absorbed vitamin D and its metabolites occurs primarily in feces with the aid of bile salts. Very little vitamin D appears inurine. Ohnuma et al. (1980) suggested that the metabolite 1,25(OH)2D3-26,23-lactone may represent one of the first steps in the catabolism of 1,25-(OH)2D3. Because the half-life of 25,26-(OH)2D in serumis only 10 days, this metabolite might have an excretory role.

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FUNCTIONS
The general function of vitamin D is to elevate plasma Ca and P toa level that will support normal mineralization of bone as well as otherbody functions. The active form of vitamin D, 1,25-(OH)2D, functions
as a steroid hormone. The hormone is produced by an endocrine gland,
circulated in blood bound to a carrier protein (DBP), and transported totarget tissues.
In the target tissue, the hormone enters the cell and binds to a cytosolic receptor or a nuclear receptor. 1,25-(OH)2D regulates gene expression
through its binding to tissue-specific receptors and subsequent interaction
between the bound receptor and the DNA (Collins and Norman, 1991;
DeLuca and Zierold, 1998). The receptor-hormone complex moves tothe nucleus, where it binds to the chromatin and stimulates the transcription of particular genes to produce specific mRNAs, which code forthe synthesis of specific proteins. Evidence for transcription regulation ofa specific gene typically includes 1,25-(OH)2D–induced modulation inmRNA levels. Additionally, evidence may include measurements of transcription and/or the presence of a vitamin D-responsive element within
the promoter region of the gene (Hannah and Norman, 1994). Recent
research provides evidence that a membrane-bound receptor, in addition
to nuclear receptors, exists (Fleet, 1999).
The 1,25-(OH)2D3 receptor has been extensively characterized, andthe cDNA for the human receptor has been cloned (Baker et al., 1988).
The 1,25-(OH)2D3 receptor is a protein with a molecular weight ofabout 67,000 daltons. The nucleotide sequence of the bovine vitamin D3
receptor has been reported (Neibergs et al., 1996).
Common vitamin D receptor gene alleles have been shown to contribute to the genetic variability in bone mass and bone turnover; however, the physiological mechanisms involved are unknown. The vitamin
D receptor alleles are associated with differences in the vitamin D
endocrine system and may have important implications in relation to thepathophysiology of osteoporosis (Howard et al., 1995; Eisman, 1998).
Recent studies have identified a heterodimer of the vitamin D receptor (VDR) and a vitamin A receptor (RXR) within the nucleus of the cellas the active complex for mediating positive transcriptional effects of
1,25-(OH)2D. The two receptors (vitamins D and A) selectively interact
with specific hormone response elements composed of direct repeats ofspecific nucleotides located in the promoter of regulated genes. The
complex that binds to these elements actually consists of three distinct
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elements: the 1,25-(OH)2D3 hormonal ligand, the vitamin D receptor(VDR), and one of the vitamin A (retinoid) X receptors (RXR) (Kliewer
et al., 1992; Whitfield et al., 1995).
Since the late 1980s, it has become apparent that 1,25-(OH)2D3 also
has the potential to generate biological actions through mechanisms notdependent on regulation of gene transcription (Norman, 1995).
Research suggests that 1,25-(OH)2D3 may also generate biologicalresponses via signal transduction mechanisms that are independent of
the nuclear VDRs, which are termed nongenomic pathways.
Nongenomic responses can include stimulation of membrane lipid
turnover, activation of Ca2+ channels, and elevation of intracellular Ca2+
concentrations, all of which have been shown to occur within seconds
after addition of 1,25-(OH)2D3. Progress has been made in identifying
and purifying an integral protein of the basal lateral membrane that may
be a receptor for 1,25-(OH)2D3 (Nemere, 1995). These studies have provided definite correlations between binding to the solubilized membrane
receptor and the ability to initiate transcaltachia (the rapid hormonal
stimulation of Ca transport). The involvement in genomic and nongenomic signal transduction pathways is not unique to the steroid hormone 1,25-(OH)2D3; these same pathways are also utilized by virtually
all steroid hormones (Nemere et al., 1993).

Relationship to Calcium and Phosphorus Homeostasis
Tetany in humans and animals results if plasma Ca levels are appreciably below normal. Two hormones—thyrocalcitonin (calcitonin) andparathyroid hormone (PTH)—function in a delicate relationship with
1,25-(OH)2D to control blood Ca and P levels. Production rate of 1,25(OH)2D is under physiological control as well as dietary control (seeMetabolism, Conversion to Metabolically Active Forms). Calcitonin,
contrary to the other two, regulates high serum Ca levels by (1) depressing gut absorption, (2) halting bone demineralization, and (3) reabsorption in the kidney. Vitamin D brings about an elevation of plasma Ca
and P by stimulating specific pump mechanisms in the intestine, bone,
and kidney. These three sources of Ca and P thus provide reservoirs thatenable vitamin D to elevate the levels of Ca and P in blood to levels thatare necessary for normal bone mineralization and for other functions
ascribed to Ca.

Intestinal Effects
It is well known that vitamin D stimulates active transport of Ca
and P across intestinal epithelium. This stimulation involves the104

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parathyroid hormone directly and the active form of vitamin D.
Parathyroid hormone indirectly stimulates intestinal Ca absorption bystimulating production of 1,25-(OH)2D under conditions of hypocalcemia. As the human body becomes vitamin D insufficient, the efficiency of intestinal Ca absorption decreases from approximately 30 to 50%
to no more than 15%.
The mechanism whereby vitamin D stimulates Ca and P absorption
is still not completely understood. Evidence (Wasserman, 1981) indicates that 1,25-(OH)2D is transferred to the nucleus of the intestinal cell,
where it interacts with the chromatin material. In response to the 1,25(OH)2D, specific RNAs are elaborated by the nucleus, and when these
are translated into specific proteins by ribosomes, the events leading toenhancement of Ca and P absorption occur (Scott et al., 1982).
In the intestine, 1,25-(OH)2D promotes synthesis of Ca-binding
protein (calbindin) and other proteins and stimulates Ca and P absorption. Vitamin D has also been reported to influence magnesium (Mg)absorption as well as Ca and P balance (Miller et al., 1965).
Administration of 1,25-(OH)2D3 to rachitic animals has been shown tostimulate the incorporation of [3H] leucine into several proteins of theintestinal mucosa. This apparent increase in protein synthesis was
accounted for at least in part by the discovery that 1,25-(OH)2D inducessynthesis of a specific intestinal protein that has been identified as calbindin. Calbindin is not present in the intestine of rachitic chicks but
appears after vitamin D treatment.
Intestinal Ca transport relies on the integrated effects of bothgenomic and nongenomic mechanisms of hormone action. Two kinds ofmucosal proteins are dependent on vitamin D: (1) calbindin and (2)intestinal membrane Ca-binding protein (IMCal). IMCal is a membranecomponent of the translocation mechanism rather than a cytosol constituent (Schachter and Kowarski, 1982). It is proposed that the primarynongenomic mechanism by which 1,25-(OH)2D regulates Ca transport
across the luminal membrane of the enterocyte involves inducing a specific alteration in membrane phosphatidylcholine content and structure,
which leads to an increase in membrane fluidity and thereby to anincrease in Ca transport rate. The size of the villus and the microvilli
increases upon 1,25-(OH)2D3 treatment. The brush border undergoes
noticeable alterations in structure and composition of cell surface proteins and lipids, in a time frame corresponding to the increase in Ca2+transport mediated by 1,25-(OH)2D3 (Collins and Norman, 1991). In
addition to inducing calbindin and IMCal, 1,25-(OH)2D3 has beenshown to increase levels of several other proteins in the intestinal
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mucosa. These include alkaline phosphatase, Ca-stimulated ATPase, andphytase enzyme activities (Collins and Norman, 1991). Once Ca istransported to the basolateral membrane, it is extruded from the cellagainst a 1,000-fold concentration gradient by Mg-dependent CaATPase, which is also increased by 1,25-(OH)2D3 (Bronner, 1987).
Originally, it was believed that vitamin D did not regulate P absorption and transport, but in 1963, it was demonstrated, through the use of
an in vitro inverted sac technique, that vitamin D does in fact play sucha role (Harrison and Harrison, 1963). Little is known about the actual
mechanism of phosphate transport, but phosphate is transported against
an electrochemical potential gradient involving sodium in response to
1,25-(OH)2D3.

Bone Effects
Vitamin D plays roles both in the mineralization of bone as well as
demineralization or mobilization of bone mineral. 1,25-(OH)2D is oneof the factors controlling the balance between bone formation andresorption. In young animals during bone formation, minerals aredeposited on the matrix. This is accompanied by an invasion of bloodvessels that gives rise to trabecular bone. This process causes bones toelongate. During vitamin D deficiency, this organic matrix fails to mineralize, causing rickets in the young and osteomalacia in adults. 1,25(OH)2D3 brings about mineralization of the bone matrix, and Weber etal. (1971) provided evidence that 1,25-(OH)2D3 is localized in the nuclei
of bone cells. Also, there is some indication that 24,25-(OH)2D3 andpossibly 25-OHD3 may have unique actions on bone.
Vitamin D also plays a role in the mobilization of Ca from bone tothe extracellular fluid compartment. This function is shared by PTH(Garabedian et al., 1974). However, little is known about the mechanism of bone reabsorption in response to these factors, although it may
be similar or identical to the intestinal transport system. It is an active
process requiring metabolic energy, and presumably it transports Ca andphosphate across the bone membrane by acting on osteocytes and osteoclasts.
Rapid, acute plasma Ca regulation is due to the interaction of plasma Ca with Ca-binding sites in bone mineral since blood is in contact
with bone. Changes in plasma Ca are brought about by a change in the
proportion of high- and low-affinity Ca-binding sites, access to which isregulated by osteoclasts and osteoblasts, respectively (Bronner and Stein,1995). These cells, in turn, respond to hormonal signals by shapechanges. Contraction of osteoclasts and corresponding expansion of106

V I TA M I N D

osteoblasts make more high-affinity sites available, whereas osteoblast
contraction and osteoclast expansion, make more low-affinity sites
available, leading to a decrease or an increase in the blood Ca level,
respectively.
Another role of vitamin D has been proposed in addition to its
involvement in bone; namely, in the biosynthesis of collagen in preparation for mineralization (Gonnerman et al., 1976). A vitamin D deficiency causes inadequate cross-linking of collagen as a result of low lysyl
oxidase activity, which is involved in a condensation reaction for the collagen cross-linking. This may be a direct effect of vitamin D or a resultof mineral changes in blood; it is not considered a major function ofvitamin D.

Kidney Effects
There is evidence that vitamin D functions in the distal renal tubules
to improve Ca reabsorption and is mediated by calbindin (Bronner andStein, 1995). It is known that 99% of the filtered load of Ca is reabsorbed in the absence of vitamin D and the parathyroid hormone. The
remaining 1% is under control of these two hormonal agents, although
it is not known whether they work in concert. It has been shown that
1,25-(OH)2D3 functions in improving renal reabsorption of Ca (Sutton
and Dirks, 1978). With intact parathyroids and without vitamin D,
renal tubular resorption of inorganic phosphate decreases, thereby
increasing phosphate clearance and resulting in hypophosphatemia,
although the parathyroids maintain a normal plasma Ca level. With adequate vitamin D, greater reabsorption of P by the renal tubules occurs.
Without intact parathyroids, vitamin D actually increases renal loss of P.

Calcium and Phosphorus Absorption by Ruminants
Calcium
There is clear evidence that sheep and cattle absorb Ca from their
gut according to need and that they can alter the efficiency of absorption to meet a change in requirement. For example, Braithwaite andRiazuddin (1971) showed that young sheep with a high Ca requirement
absorb Ca at a higher rate and with greater efficiency than mature animals with a low requirement. An increase in both absorption and efficiency of absorption also occurs in mature sheep when their requirement
for Ca is increased through pregnancy or lactation or after a period of
Ca deficiency (Braithwaite, 1974).
Studies in cattle have given similar results. Thus, the efficiency of107

CHAPTER THREE

absorption of Ca in the small intestine of the dairy cow has been shownto rise in response to a reduction in dietary Ca intake and to the onsetof lactation (Scott and McLean, 1981). Calcium absorption has also
been shown to be directly related to milk production, though in earlylactation, when the demand for Ca is greatest, the increase in absorption
falls short of the requirement, with the deficit being met by increasedbone resorption (Braithwaite et al., 1969).
The mechanism by which Ca is adjusted in response to requirement
has received much attention. In this sequence, a fall in plasma Ca concentration resulting from an increase in demand leads in turn to anincrease in parathyroid hormone release. This then stimulates theincreased production by the kidney of 1,25-(OH)2D, which acts on the
gut to increase the production of calbindin and so accelerates Ca absorption. In a reverse manner, an increase in plasma Ca concentration causes suppression of parathyroid hormone release, a reduction in 1,25(OH)2D production, and reduced Ca absorption. Although all aspects ofthis system have not yet been fully examined in ruminants, it appears
that the same mechanism operates, in that an increase in circulatory
1,25-(OH)2D level has been found to precede the increase in Ca absorption that occurs in cattle soon after parturition (Horst et al., 1978).

Phosphorus Absorption
Sheep fed on roughage diets usually excrete little P in their urine(Scott and McLean, 1981), so control of P balance must therefore beachieved within the gut through control of either absorption or secretionor both. Saliva is the main contributor of P in the gut. Little or no net
absorption of P appears to occur from either the forestomach or thelarge intestine, and it is generally accepted that the upper small intestineis the major absorptive site. Using sheep fitted with reentrant cannulas,
several workers have shown that the secretion of P before the pylorus(salivary) is closely matched by net absorption in the small intestine.
Ability to balance absorption against secretion has been shown to beunaffected by wide variations in dietary Ca:P values (Scott and McLean,
1981).
Until recently, secretion of P in saliva has usually been viewed in thecontext of its role as a buffer against the volatile fatty acids produced in
the rumen. However, studies by Australian workers (Tomas, 1974) havesuggested that salivary glands, apart from their role as a source of buffer,
may also play an important role in P homeostasis by controlling theamount of P secreted into the gut. Evidence for this function was provided by sheep studies in which both parotid salivary ducts were ligat108

V I TA M I N D

ed, a procedure that led to a small increase in urinary P excretion and a
proportional reduction in fecal P excretion (Tomas and Somers, 1974).
Similar changes in the pathway of P excretion were also seen in sheep inwhich part of the parotid salivary flow was collected and returned
directly to the circulation.
Ruminants have a higher renal threshold for P excretion than domonogastric species, and it is important to consider what advantage thisconfers. Poor-quality roughage diets, apart from their low digestibility,
also tend to contain little P (McDowell, 1997). Therefore, the ratio
between the amount of P required for saliva production and dietaryintake is wide. If the renal threshold for P excretion in ruminants wereas low as in monogastric species, then at times when the concentrationof inorganic P in the plasma rises in response to reabsorption, P would
be excreted in the urine. This P would not, however, in any real sense be
surplus to requirement, and its loss would have to be met from a diet
low in available P. Under such conditions, there is clearly an advantage
to the ruminant in maintaining the high renal threshold for P excretion.
Concentrate diets, especially those that include fish meal, containmuch larger quantities of P than do roughage diets, to the point where
intake may equal or exceed the amount secreted in the saliva. Under
these conditions, need to control P absorption is clearly less critical and,as a result, a different level of control may operate. Increasing dietary P
intake leads to increased absorption and increased urinary P excretion.
Administration of large amounts of parathyroid hormone over several days has been shown to reduce fecal P excretion in cattle (Mayer etal., 1968), though whether this was due to reduced secretion or
increased absorption is not clear. In sheep, parathyroidectomy has beenshown to result in a negative balance for both Ca and P, and it has beensuggested that the effect on P balance was the result of a reduction in theamount of salivary P reabsorbed (McIntosh and Tomas, 1978). 1,25(OH)2D has also been suggested as a possible regulator of intestinal P
absorption in ruminants (Scott and McLean, 1981), though whether thiswas a primary effect or secondary to its effects on Ca absorption anddeposition in bone is not clear.

Other Vitamin D Functions
The well-known effects of vitamin D relate to biochemical changes
occurring in the intestine, bone, and kidney. Vitamin D has also beenshown to be required for chick embryonic development. Vitamin D
treatment stimulated yolk Ca mobilization, and the vitamin D-dependent Ca-binding protein, calbindin, is present in the yolk sac (Tuan and109

CHAPTER THREE

Suyama, 1996). These findings strongly suggest that the hormonal
action of 1,25-(OH)2D on yolk sac Ca transport is mediated by the regulated expression and activity of calbindin, analogous to the response of
the adult intestine. 1,25-(OH)2D is also essential for the transport ofeggshell Ca to the embryo across the chorioallantoic membrane(Elaroussi et al., 1994).
Vitamin D is important for more than just its traditional role in Ca
metabolism. A receptor for the active metabolite 1,25-(OH)2D has
been isolated in the pancreas, parathyroid glands, bone marrow, certain cells of the ovary and brain, endocrine cells of the stomach, breastepithelial cells, skin fibroblasts, and keratinocytes, suggesting that
1,25-(OH)2D has additional functions in a wide variety of cells,
glands, and organs (Machlin and Sauberlich, 1994). In the pancreas,
1,25-(OH)2D is essential for normal insulin secretion. Experimentswith rats have shown that vitamin D increases insulin release from isolated perfused pancreas, both in the presence or absence of normal
serum Ca levels (Cade and Norman, 1986). More than 50 genes havebeen reported to be transcriptionally regulated by 1,25-(OH)2D(Hannah and Norman, 1994).
The actions of 1,25-(OH)2D3 are recognized as being involved inregulation of the growth and differentiation of a variety of cell types,
including those of hematopoietic and immune systems (Lemire, 1992).
Studies have suggested 1,25-(OH)2D3 as an immunoregulatory hormone. Elevated 1,25-(OH)2D also has been associated with a significant
70% enhancement of lymphocyte proliferation in cells treated withpokeweed mitogen (Hustmeyer et al., 1994). 1,25-(OH)2D3 also inhibits
growth of certain malignant cell types and promotes their differentiation(Colston et al., 1981; DeLuca, 1992). 1,25-(OH)2D has been reported toinhibit proliferation of leukemic cells (Pakkala et al., 1995), breast cancer cells (Vink van Wijngaarden et al., 1995), and colorectal cancer cells(Cross et al., 1995). A deficiency of vitamin D may promote prostatecancer (Skowronski et al., 1995). Also, 1,25-(OH)2D and its analogs
may be effective in treating some forms of psoriasis (Kragballe et al.,1991). The therapeutic value of vitamin D and its analogs has been
under rigorous evaluation in numerous laboratories around the world.
Thus, the concept has emerged that 1,25-(OH)2D, and possibly
other vitamin D metabolites, have functions that extend beyond those ofregulating bone mineralization and intestinal Ca transport. The skin is
one such tissue in which this broader role is being intensively explored.
Besides producing vitamin D, epidermal cells (keratinocytes) make 1,25(OH)2D and respond to the 1,25-(OH)2D they produce with a decrease110

V I TA M I N D

in proliferation and an increase in differentiation (Bikle et al., 1995).
1,25-(OH)2D production by keratinocytes is tightly controlled andchanges as the cells differentiate, increasing during the early stages ofdifferentiation and then decreasing again as terminal differentiation
ensues. The 1,25-(OH)2D produced endogenously or supplied exogenously acts in concert with Ca and products of phosphoinositide metabolism to stimulate the transition from a proliferating basal cell to a terminally differentiated corneocyte. The mechanisms involved include
changes in gene transcription and messenger RNA stability.

REQUIREMENTS
Most animals and humans do not have a nutritional requirement forvitamin D when sufficient sunlight is available, since vitamin D3 is produced in skin through action of UV light on 7-dehydrocholesterol.
Unlike humans, rats, common poultry, and livestock, dogs and cats
(and perhaps other carnivores) have a nutritional requirement for vitamin D even when sufficient sunlight is available, since vitamin D3 is notproduced in skin through action of UV irradiation on 7-dehydrocholesterol in sufficient quantities to prevent rickets (How et al., 1994a,b,
1995). Hazewinkel et al. (1987) found that rickets in dogs could not be
prevented or treated by UV radiation; treated dogs developed clinical,
biochemical, and histological signs of rickets. In the skin of the dog andcat, the concentrations of the precursor 7-dehydrocholesterol are low,
and the precursor is inadequately converted to vitamin D. It is suggested that carnivores do not need to provide their own vitamin D, since fat,liver, and blood of their prey will fulfill this need (How et al., 1995). In
addition to sunlight, other factors influencing dietary vitamin D requirements include (1) amount and ratio of dietary Ca and P, (2) availabilityof P and Ca, (3) species, and (4) physiological factors.
Vitamin D becomes a nutritionally important factor in the absenceof sufficient sunlight. Sunlight that comes through ordinary window
glass is ineffective in producing vitamin D in skin, since glass does notallow penetration of UV rays, and its effectiveness is dependent onlength and intensity of UV rays that reach the body. The radiation thatreaches the earth contains only a small part of the UV range that has anantirachitic effect. Sunlight is more potent in the tropics than in the temperate or arctic zones, more potent in summer than in winter, morepotent at noon than in the morning or evening, and more potent at highaltitudes. Sunlight provides most of its antirachitic powers during the 4hours around noon.
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Season and latitude greatly influence skin synthesis of vitamin D3.
In Boston (42.2oN), previtamin D3 was produced in the irradiated skinsamples exposed to sunlight from March through October, but not at allfrom November through February (Webb et al., 1988). In Edmonton,
Alberta, Canada (52o N), the effective period for the synthesis of previtamin D3 commenced at the beginning of April and ceased after October.
In Los Angeles (34o N) and Puerto Rico (18oN), sunlight effectively photoconverted 7-dehydrocholesterol to previtamin D3 even in the middleof winter. These data confirm that the conversion of 7-dehydrocholesterol to previtamin D3 is dependent on the intensity of UV radiation asaffected by the zenith angle of the sun at various latitudes and by seasons of the year.
Besides geographical and seasonal considerations, sunlight may beblocked by many means. Potency of UV light varies greatly with differences in atmospheric conditions and because of clouds, mist, or smoke.
Air pollution screens out many UV rays as does the use of sunscreens.
Air pollution became prevalent during the Industrial Revolution and, atthe same time, the incidence of rickets became widespread in industrial
cities. It is now known that this epidemic of rickets was partly due to thelack of sunlight because of the presence of air pollution. Thus, rickets
has been called the first air-pollution disease. Animals housed for muchof the year must depend on their feed for the vitamin D they need; in amodern agricultural economy, this applies particularly to intensive swine
and poultry production.
The colors of the coat and skin are important in determining
response to irradiation. UV irradiation is more effective on exposed skinthan through a heavy coat of hair. Irradiation is less effective on darkpigmented skin. This has been shown to be true for white and black
breeds of hogs as well as for people. White pigs have taken about twice
as long as colored pigs to show signs of vitamin D deficiency (Cunha,1977).
In white humans, 20 to 30% of the UV radiation is transmitted
through the epidermis, while in black people, less than 5% of the UV
radiation penetrates the epidermis (Holick, 1987). When specimens ofsurgically obtained white and black skin were exposed to simulated sunlight under the same conditions, longer exposure times were needed tomaximize vitamin D3 formation in the samples of black skin. As skinpigmentation increased, the time required to maximize vitamin D3 formation increased from 30 minutes to 3 hours.
Aging has an effect on the cutaneous production of vitamin D3. In
humans older than about 20 years, skin thickness decreases linearly with112

V I TA M I N D

time. A comparison of amounts of previtamin D3 generated in the skinof young and elderly subjects showed that aging can decrease more than
twofold the capacity of the skin to produce previtamin D3 (Holick,1986).
Amounts of dietary Ca and P, and the physical and chemical forms
in which they are presented, must be considered when determining vitamin D requirements. High dietary Ca concentrations can precipitate
phosphates as insoluble Ca phosphate. If Ca is given in the form of a relatively insoluble compound, or even a Ca compound that is normallyeasily soluble but is too coarsely ground, it may be comparatively
unavailable, for example, coarse limestone (Ca carbonate) (Franklin andJohnstone, 1948). Soluble Ca salts are more readily absorbed, andoxalates tend to interfere with absorption, but some of this interference
can be overcome by dietary vitamin D or irradiation.
Correspondingly, while the P of inorganic orthophosphate tends tobe well absorbed, other factors being favorable, that of phytic acid (seeChapter 17), which is the predominant P compound of unprocessedcereal grains and oilseeds, seems to be poorly available except to species(such as ruminants) with massive populations of microorganisms in the
gut that synthesize phytase enzymes. Phosphorus absorption is mostly
independent of vitamin D intake, with the inefficient absorption and theimprovement upon vitamin administration being a result of improving
Ca absorption.
The need for vitamin D depends to a large extent on the Ca-P ratio.
As this ratio becomes either wider or narrower than the optimum, thevitamin D requirement increases, but no amount will compensate forsevere deficiencies of either Ca or P. The dietary dry matter of rapidly
growing young stock should contain approximately 0.6 to 1.2% Ca,with a Ca-P ratio in the range of about 1.2:1 to 1.5:1. For adult maintenance, lower Ca levels and wider Ca-P ratios are possible. In these situations, vitamin D requirements seem to be at a minimum, and risks ofvitamin D deficiency are less probable. The young rat needs very little
vitamin D. Indeed, to make the species suitably responsive in the biological assay of the vitamin, the dietary P content has to be kept low and
the Ca-P ratio high (Abrams, 1978).
Intestinal pH as well as other dietary nutrients influence Ca and P
requirements, and thus vitamin D requirements. Fermentation of excesscarbohydrates makes intestinal contents more acid, which favors both
Ca and P absorption, probably by converting less soluble alkaline salts
to the more soluble acid salts. High intakes of fats containing higher
fatty acids increase highly insoluble Ca soaps. Potassium may increase P
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CHAPTER THREE

absorption, but cations that form insoluble phosphates such as iron andaluminum interfere.
Vitamin D requirements of various species (Table 3.1) are sufficiently high to produce normal growth, calcification, production, andreproduction in the absence of sunlight, provided that diets contain recommended levels of Ca and available P. Species differences can be illustrated by the fact that adequate intakes of Ca and P in a diet that contains only enough vitamin D to produce normal bone in rats or pigs willquickly cause the development of rickets in chicks. Turkeys and pheasants have higher requirements than chicks. Surprisingly, human babies■ Table 3.1

Vitamin D Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Milk replacer
Growing bulls
Lactating
All classes

275 IU/kg
300 IU/kg
300 IU/kg
1,000 IU/kg
1,400 IU/kg

Leghorn, 0–18 weeks
Leghorn, laying (100-g
intake)Broilers, 0–8 weeks
0–7 weeks
All classes
Growing
All classes
Adult

200 ICU/kg
300 ICU/kg

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1989a)
Morand-Fehr(1981)NRC (1994)
NRC (1994)

Goat
ChickenDuck (Peking)
Geese
Japanese quail
Turkey
Sheep
Swine
Horse
Cat
Dog
Fish
Rat
Mouse
Guinea pig
Human

Growing-finishing, 3–10 kg
Growing-finishing, 20–120 kg
Adult
Maintenance and working
Growing
Pregnancy and lactation
Growing
GrowingCatfishRainbow trout
Growing-reproductive
Growing
GrowingChildren
AdultsLactating

200 ICU/kg
400 ICU/kg
200 IU/kg
750 ICU/kg
1,100 ICU/kg
555 IU/100 kg
body weight
220 IU/kg
150 IU/kg
200 IU/kg
300 IU/kg
800 IU/kg
600 IU/kg
500 IU/kg
22 IU/kg bodyweight250–1,000 mg/kg
1,600–2,400 IU/kg
1,000 IU/kg
1,000 IU/kg
1,000 IU/kg
400 IU/day
200–400 IU/day
400 IU/day

NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1989b)
NRC (1989b)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as IU/day (1 IU= 0.025 μg vitamin D).

114

V I TA M I N D

are more like birds in this respect than like the other mammals mentioned. Greater absorption of Ca and greater growth rates occurred infull-term infants given 400 IU per day compared to those given 100 IU
per day, although 100 IU is enough to prevent rickets (Collins andNorman, 1991). In addition to the RDA (1989) suggested requirements
for humans, it is recognized that without casual exposure to sunlight,
the dietary intake for vitamin D should be at least two to three times
more than the recommended intake (Holick, 1995).

NATURAL SOURCES
Ergocalciferol and cholecalciferol contents of natural materials areshown in Table 3.2. Sources of vitamin D are natural foods, irradiatedsebaceous material licked from skin or hair, and directly absorbed products or irradiation formed on or in the skin.
In dogs and cats (and presumably other carnivores), vitamin D must
be obtained from dietary sources due to the inability of these species tosynthesize and utilize vitamin D from precursors in the skin (How et al.,1995). The distribution of vitamin D is very limited in nature, althoughprovitamins D occur widely. Of feeds for livestock, grains, roots, andoilseeds, as well as their numerous by-products, contain insignificant
amounts of vitamin D. With a few notable exceptions, vitamin D3 is notfound in plants. Those exceptions include the species Solanum malacoxylon, Cestrum diurnum, and Trisetum flavescens (see Toxicity), in
which vitamin D occurs as water-soluble β−glycosides of vitamin D3, 25OHD3 and 1,25-(OH)2D3.
Typical green forages are extremely poor in vitamin D. However,when plants—especially pasture species—begin to die and the fading
leaves are favorably exposed to UV light, some vitamin D2 is formed. So
arises the vitamin D of hay, the potency of which depends on local climatic conditions, for if it is made very quickly in the absence of direct
sunlight and is baled when still quite green, its potency will be low. The
principal source of the antirachitic factor in the diets of farm animals isthus provided by the action of radiant energy upon ergosterol in forages.
Legume hay that is cured to preserve most of its leaves and green color
contains considerable amounts. Alfalfa, for example, will range from
650 to 2,200 IU/kg (Maynard et al., 1979); timothy and other grass hays
contain less. Stemmy hay that is lacking in leaves and color and has been
exposed to a minimum of sunlight may contain none, whether legume
or nonlegume. The antirachitic value of the alfalfa crop increases withstate of maturity because of the increase in dead leaves.
115

CHAPTER THREE■ Table 3.2

Vitamin D Concentrations in Various Foods and Feedstuffs

Food or Feedstuff
Alfalfa pasture
Alfalfa hay, sun cured
Alfalfa silage
Alfalfa wilted silage
Birdsfoot trefoil hay, sun cured
Barley straw
Cocoa shell meal, sun driedCorn grain
Corn silage
Molasses, sugar beet
Red clover, fresh
Red clover, sun cured
Sorghum grain
Sorghum silage

IU/100 ga
Ergocalciferol (D2)
4.6
142
12
60
142
60
3,500
0
13
58
4.7
192
2.6
66

Food or Feedstuff
Blue fin tuna liver oil
Cod liver oil meal
Cod liver oil
Eggs
Halibut liver oil
Herring, entire body oil
Menhaden, entire body oil
Milk, cow’s whole (summer)
Milk, cow’s whole (winter)
Sardine, entire body oil
Sturgeon liver oil
Swordfish liver oil

IU/100 g
Cholecalciferol (D3)4,000,000
4,000
10,000100
120,000
10,000
5,000
4
1
8,000
0
1,000,000

Sources: Adapted from NRC (1982b) and Scott et al. (1982).

aConcentrations are on as fed-basis.

Artificially dried and barn-cured hay contains less vitamin D than
hay that is properly sun cured. Even hay dried in the dark immediately
after cutting contains some of the vitamin, because the dead or injured
leaves on the growing plant respond to irradiation even though the living tissues do not. This fact is also largely responsible for the vitamin D
found in corn silage (Maynard et al., 1979). Under normal conditions,even wilted legume silage furnishes ample vitamin D for dairy calves.
For non–forage-consuming species, the vitamin D that occurs naturally in unfortified food is generally derived from animal products.
Saltwater fish, such as herring, salmon, and sardines, contains substantial amounts of vitamin D, and fish liver oils are extremely rich sources.
The probable origin of vitamin D in fish liver is a result of food chainsfrom plankton (Takeuchi et al., 1991). Veal, beef, unfortified milk, and116

V I TA M I N D

butter supply only small quantities. Among animal products, eggs, especially the yolks, are a good source if the hen’s diet is rich in the vitamin.
Milk contains a variable amount in its fat fraction (5 to 40 IU in cow’s
milk per quart), but neither cow’s milk nor human milk contains enoughto protect the baby against rickets. Cow’s milk is reportedly higher invitamin D when produced during the summer rather than winter.
It has generally been assumed that for all but a few species, vitamin
D2 and vitamin D3 are equally potent. In poultry, other birds, and a few
of the rarer mammals, including some New World monkeys, vitamin D3
has been found to be many times more potent than D2 on a weight basis.
Vitamin D3 may be 30 times more effective than D2 for poultry; therefore, plant sources (vitamin D2) of the vitamin should not be provided
to these species. The dogma that mammals (other than the New World
monkey) do not discriminate between the two sources has been provenincorrect. Data for pigs (Horst and Napoli, 1981) and ruminants(Sommerfeldt et al., 1981) have suggested that these species discriminate
in the metabolism of vitamin D2 and D3, with vitamin D3 being the preferred substrate. Pigs given oral doses of a mixture of vitamin D2 and D3(1:w/w) had significantly higher concentrations of plasma vitamin D3,
25-OHD3, 24,25-(OH)2D3, and 1,25-(OH)2D3 than those of corresponding counterparts given vitamin D2. Sommerfeldt et al. (1983) reported
that the amount of 1,25-(OH)2D in the plasma of ergocalciferol-treateddairy calves was one-half to one-fourth the amount in the plasma ofcholecalciferol-treated calves. Discrimination against vitamin D2 byruminants may be in part a result of its preferred degradation by rumen
microbes or its less efficient absorption by the intestine. Similarly,
Harrington and Page (1983) found vitamin D3 to be more hypercalcemicand overtly toxic to horses than vitamin D2, likewise suggesting preference for D3. Vitamin D3 has been reported to be at least three times as
effective as vitamin D2 in satisfying the vitamin D requirement in trout(NRC, 1993). Although the more recent data suggest a preference for D2
by a number of animals, in practice D2 is still relatively comparable toD3 in antirachitic function except for poultry and certain monkeys.

DEFICIENCY
Effects of Deficiency
The outstanding disease of vitamin D deficiency is rickets, generally
characterized by a decreased concentration of Ca and P in the organic
matrices of cartilage and bone. The signs and symptoms are similar to117

CHAPTER THREE

those of a lack of Ca or P or both, as all three are concerned with proper bone formation. In the adult, osteomalacia is the counterpart of rickets and, since cartilage growth has ceased, is characterized by adecreased concentration of Ca and P in the bone matrix. Osteoporosis
is defined as a decrease in the amount of bone, leading to fractures after
minimal trauma. In osteoporosis, bone mineral and protein matrix are
lost, resulting in less overall bone but normal composition.
Osteomalacia is also characterized by inadequate bone mineralization;
however, in contrast to osteoporosis, persons with osteomalacia havenormal protein matrix that is not fully mineralized. When bone mass
becomes too low, mechanical support and skeletal integrity cannot be
maintained, and fractures can occur with minimal trauma.
Clinical signs of vitamin D deficiency are seen mainly in the young.
General consequences of deficiency can appear as inhibited growth,weight loss, and reduced or lost appetite before characteristic signs that
relate primarily to the bone system become apparent. The role of vitamin D in the adult appears to be much less important except during
reproduction and lactation. Congenital malformations in newborns
result from extreme deficiencies in the diet of the mother during gestation, and the mother’s skeleton is injured as well.
The same disruption of the orderly processes of bone formation
with vitamin D deficiency occurs in animals as it does in humans andincludes the following characteristics (Kramer and Gribetz, 1971):
1. Failure of Ca salt deposition in the cartilage matrix.
2. Failure of cartilage cells to mature, leading to their accumulation
rather than destruction.
3. Compression of the proliferating cartilage cells.
4. Elongation, swelling, and degeneration of proliferative cartilage.
5. Abnormal pattern of invasion of cartilage by capillaries.

Outward signs of rickets include the following skeletal changes,
varying somewhat with species depending on anatomy and severity:
1. Weak bones, causing curving and bending of bones.
2. Enlarged hock and knee joints.
3. Tendency to drag hind legs.
4. Beaded ribs and deformed thorax.

Although there appear to be differences between species in thesusceptibility of different bones to such degenerative changes, differ118

V I TA M I N D

ences that probably reflect bodily conformation (e.g., pigs comparedwith sheep) and stance (e.g., humans compared with the common
quadrupeds), there is nevertheless an apparent common pattern(Abrams, 1978). Spongy parts of individual bones, and bones relatively rich in spongy tissue, are first and worst affected. As in simple
Ca deficiency, the vertebrae and the bones of the head suffer thegreatest degree of resorption. Next come the scapula, sternum, andribs. The most resistant bones are the metatarsals and the shafts oflong bones.

Ruminants
Clinical signs of vitamin D deficiency in ruminants are decreased
appetite and growth rate, digestive disturbances, rickets (Figs. 3.3 and3.4), stiffness in gait, labored breathing, irritability, weakness, and occasionally tetany and convulsions. There is enlargement of joints, slightarching of the back, and bowing of legs, with erosion of joint surfaces

A

B

Fig. 3.3 Calves developed severe rickets
while receiving ration
deficient in vitamin D,
and kept away fromsunlight. (A: Courtesyof W. Krauss, Ohio
Agriculture Experiment
Station. B: Courtesy ofMichigan Agriculture
Experiment Station,
NRC, 1958.)
119

CHAPTER THREE

Fig. 3.4 Calves and young bulls with rickets. (Courtesy of the late Francisco
Megale, Universidade Federal de Minas Gerais, Escola de Veterinaria, Belo
Horizante, MG, Brazil.)

causing difficulty in locomotion (NRC, 1996). Young ruminants may beborn dead, weak, or deformed.
Clinical signs involving bones begin with thickening and swelling of
the metacarpal or metatarsal bones. As the disease progresses, theforelegs bend forward or sideways. In severe or prolonged vitamin D
deficiency, tension of the muscles will cause bending and twisting of longbones to give the characteristic deformity of bone. There is enlargement
at ends of bones from deposition of excess cartilage, giving the characteristic “beading” effect along the sternum where ribs attach (NRC,
1989a, 1996). The lower jaw bone (mandible) becomes thick and soft;in severe cases, eating is then difficult. Calves may experience slobbering, inability to close the mouth, and protrusion of the tongue (Craig
and Davis, 1943). Joints (particularly the knee and hock) become
swollen and stiff, the pastern straight, and the back humped. In more
severe cases, synovial fluid accumulates in the joints (NRC, 1989a).
Posterior paralysis may also occur as the result of fractured vertebrae.
The structural weakness of the bones appears to be related to poor mineralization. The advanced stages of the disease are marked by stiffness120

V I TA M I N D

of gait, dragging of the hind feet, irritability, tetany, labored and fastbreathing, weakness, anorexia, and retardation of growth. Calves may
be born dead, weak, or deformed (Rupel et al., 1933).
Clinical signs of vitamin D deficiency in sheep and goats are similar
to those in cattle, including rickets in young animals and osteomalacia
in adults (NRC, 1981, 1985b). An early report of rickets in Scotlandreferred to the condition as “bent leg,” which occurred in ram lambs 7
to 12 months of age (Elliot and Crichton, 1926). The condition was prevented by administration of small doses of vitamin D in the form of cod
liver oil. Newborn lambs can receive enough vitamin D from their dams
to prevent early rickets if the dams have adequate storage (Church andPond, 1974). Newborn kids had rickets if the dam was deficient in vitamin D during pregnancy (NRC, 1981).
In older animals with vitamin D deficiency (osteomalacia), bones
become weak and fracture easily, and posterior paralysis may accompany vertebral fractures. In dairy cattle, milk production may be decreased
and estrus inhibited by inadequate vitamin D (NRC, 1989a). Cows fed
a diet deficient in vitamin D and kept out of direct sunlight showed definite signs of vitamin D deficiency within 6 to 10 months (Wallis, 1944).
Functions that deplete vitamin D are high milk production and advancing pregnancy, especially during the last few months before calving. The
visible signs of vitamin D deficiency in dairy cows are similar to those of
rickets in calves. The animal begins to show stiffness in limbs and joints,
which makes it difficult to walk, lie down, and get up. The knees, hocks,
and other joints become swollen, tender, and stiff. The knees often
spring forward, the posterior joints straighten, and the animal is tilted
on its toes. The hair becomes coarse and rough, and the animal has an
overall appearance of unthriftiness (Wallis, 1944). As the deficiency
advances, the back often becomes stiff, humped, bent, and flexed. In
vitamin D-deficient herds, calving rates are lower and calves have beenborn dead or weak.
Milk fever (parturient paresis) is a paralyzing metabolic diseasecaused by hypocalcemia near parturition and initiation of lactation inhigh milk-producing dairy cows. Milk fever is an impaired metabolic
condition that is related to Ca status, previous Ca intake, and malfunction of the hormone form of vitamin D [1,25-(OH)2D] and PTH.
Animals that develop milk fever are unable to meet the sudden demand
for Ca that is brought about by the initiation of lactation. Milk fever
usually occurs within 72 hours after parturition and is manifested bycirculatory collapse, generalized paresis, and depression of consciousness. The most obvious and consistent abnormality is acute hypocal121

CHAPTER THREE

cemia, in which serum Ca decreases from a normal 8 to 10 mg/dL to 3
to 7 mg/dL (average, 5 mg/dL). Early in the onset, the cow may exhibit
some unsteadiness as she walks. More frequently, the cow is found lying
on her sternum with her head displaced to one side, causing a kink in
the neck, or turned into the flank. The eyes are dull and staring and thepupils dilated. If treatment is delayed many hours, the dullness gives way
to coma, which becomes progressively deeper, leading to death.
Aged cows are at the greatest risk of developing milk fever. Heifers
almost never develop milk fever. Older animals have a decreased
response to dietary Ca stress due to both decreased production of 1,25(OH)2D and decreased response to the 1,25-(OH)2D. Target tissues ofcows with milk fever may have defective hormone receptors, and thenumber of receptors declines with age. In older animals, fewer osteoclasts exist to respond to hormone stimulation, which delays the abilityof bone to contribute Ca to the plasma Ca pool (Goff et al., 1989). The
aging process is also associated with reduced renal 1α-hydroxylase
response to Ca stress, therefore reducing the amount of 1,25-(OH)2D
produced from 25-OHD (Goff et al., 1991b).
Parturient paresis also occurs in ewes. It is a disturbance of metabolism in pregnant and lactating ewes characterized by acute hypocalcemia and the rapid development of hyperexcitability, ataxia, paresis,coma, and death. The disease occurs any time from 5 weeks before to
10 weeks after lambing, principally in highly conditioned older ewes atpasture. The onset can be associated with an abrupt changing of feed, asudden change in weather, or short periods of fasting imposed by circumstances such as shearing or transportation. The degree of involvement of vitamin D with Ca metabolism in parturient paresis with sheep
is unclear.
Vitamin D should be supplied to growing animals that are denied
sunlight over extended periods because of cloud cover or confinement
housing. In northern latitudes during winter, photochemical conversion
of provitamin D to its active compound in the skin of ruminants is limited because of insufficient UV radiation. Hidiroglou et al. (1979)
reported that 25-OHD was higher in plasma of cattle in summer than in
winter. Richter et al. (1990) found that 25-OHD concentrations ofblood plasma were higher if bulls were kept outdoors versus indoors(21.1 versus 14.3 ng per milliliter, respectively). Vitamin D deficiency
may be observed in young ruminants that are closely confined and do
not consume sun-cured roughage. Hidiroglou et al. (1978) reported theclinical history of a flock of sheep kept under total confinement thatshowed a high incidence (8%) of an osteodystrophic condition (a vita122

V I TA M I N D

min D-responsive disease). A form of osteodystrophia has also been produced experimentally in goats (NRC, 1981).
When grazing ruminants have normal exposure to direct sunlight or
are fed normal amounts of sun-cured forage, little chance for vitamin D
deficiency exists. However, seasons of minimum sunlight, artificially
cured forages, sheep with full fleece, feedlot animals without access tosunlight or sun-cured forages, and high-producing dairy cows with limited access to sunlight or sun-cured forage may lead to the need for
dietary supplementation.

Swine
In swine, vitamin D deficiency causes poor growth, stiffness, lameness (Fig. 3.5), stilted gait, a general tendency to “go down,” or lose theuse of the limbs (posterior paralysis), frequent cases of fractures, softness of bones, bone deformities, beading of the ribs, enlargement anderosion of joints, and unthriftiness (Cunha, 1977). Bones may also be
deformed by the weight of the animal and the pull of body muscles.
Pigs with severe vitamin D deficiency may exhibit signs of Ca andFig. 3.5 Pig with advanced rickets caused by lack of vitamin D. The pig was
not exposed to sunlight. It shows leg abnormalities and was unable to walk; itlater responded to supplementary vitamin D. (Courtesy of J.M. Bell, Universityof Saskatchewan, Canada.)123

CHAPTER THREE

magnesium deficiency, including tetany (NRC, 1998). It takes 4 to 6
months for pigs fed a vitamin D-deficient diet to develop signs of a deficiency (Johnson and Palmer, 1939; Quarterman et al., 1964).
The trend toward confinement of swine in completely enclosed
houses through the life cycle increases the importance of adequate
dietary vitamin D fortification. Goff et al. (1984) concluded that subclinical rickets may become more of a problem as swine producers convert to swine-confinement operations, which deprive sows and piglets of
the UV irradiation needed for endogenous production of cholecalciferol.
Research has shown that sunshine cannot always be depended on tomeet vitamin D requirements of growing or finishing pigs during wintermonths in northern latitudes.
Of all the vitamins provided in swine feeds, vitamin D is one of two(the other is vitamin B12) that is most likely to be deficient. Typicalgrain- and soybean-based diets contain virtually none of the vitamin.
Also, the trend toward complete confinement will eliminate UV light asa source of the vitamin; therefore, supplemental vitamin D must be provided for all swine operations in which growing and breeding animals
remain in confinement.

Poultry
Little difference exists among poultry species in relation to clinicalsigns of deficiency. Clinical signs in all poultry species include rickets
and lowered growth rate, egg production, and hatchability.
In addition to retarded growth, the first sign of vitamin D deficiency in chicks is rickets, which is characterized by severe weakness of thelegs. During vitamin D deficiency, growing birds develop hypocalcemia,
which in turn stunts skeletal development through widened cartilage at
epiphyses of long bones and weakened shafts (NRC, 1994). Young,growing chickens and turkeys tend to rest frequently in a squatting position, are disinclined to walk, and have a lame, stiff-legged gait. These are
distinguished from the clinical signs of vitamin A deficiency in that vitamin D-deficient birds are alert rather than droopy, and walk with a lamerather than a staggering gait (ataxia). The beaks and claws become soft
and pliable (Fig. 3.6), usually between 2 and 3 weeks of age. The mostcharacteristic internal sign of vitamin D deficiency in chicks is a beadingof the ribs at their juncture with the spinal column (Scott et al., 1982).
In chronic vitamin D deficiency, marked skeletal distortions become
apparent (Scott et al., 1982), in which the spinal column may bend
downward in the sacral and coccygeal region. The sternum usually

124

A

B
Fig. 3.6 Vitamin D-deficient and normal poult: (A) Note rubbery beak fromvitamin D deficiency; (B) (Courtesy of L.S. Jensen and Washington State
University.)125

CHAPTER THREE

shows both a lateral bend and an acute dent near the middle of thebreast. These changes reduce the size of the thorax, with consequent
crowding of the vital organs.
A disease condition known as endochondral ossification defects(EOD) produces bone deformations, fractures, and lameness in broiler
chickens throughout the world within the first few weeks after hatching.
Flocks with a high incidence of EOD have significantly lower bone ash
and 1,25-(OH)2D3 than do mildly affected flocks, and it seems probablethat higher systemic concentrations of 1,25-(OH)2D3 between 7 and 14
days of age will enhance the ability of broiler chickens to effectively mineralize the cartilaginous growth plates in the appendicular skeleton during early bone maturation (Vaiano et al., 1994).
As in many other nutritional diseases of poultry, the feathers soon
become ruffled. In red or buff-colored breeds of chickens, vitamin D
deficiency causes an abnormal black pigmentation of some of the feathers, especially those of the wings. If the deficiency is very marked, theblackening becomes pronounced, and nearly all the feathers may beaffected (NRC, 1994). When vitamin D is supplied in adequate quantity, the new feathers and newer parts of older feathers are normal incolor, although the discolored portion remains black.
Signs of vitamin D deficiency begin to occur in laying hens in confinement about 1 to 2 months after they are deprived of vitamin D.
When laying chickens are fed a diet deficient in vitamin D, the first signof the deficiency is a thinning of the shells of their eggs. Commercial layers will continue to lay eggs with reduced shell quantity for weeks. If thediet is also completely devoid of vitamin D3, egg production may
decrease rapidly, and eggs with a very thin shell or no shell will be produced. In laying hens, eggshell strength tends to decrease as the hens age.
The decline in shell strength may be due to a decrease in the hen’s ability to synthesize 1,25-(OH)2D. A study of the effect of dietary 1,25(OH)2D3 on eggshell strength in older hens found that within 3 weeks,
the percentage of cracked or broken eggs was lower for hens supplemented with 1,25-(OH)2D (Tsang, 1992).
Vitamin D nutriture of the hen also influences its content in egg yolk
and the subsequent need for this vitamin by the chick (Stevens and Blair,
1985). Hatchability is markedly reduced, with embryos frequently dying
at 18 to 19 days of age. These embryos show a short upper mandible orincomplete formation at the base of the beak. Eventually, breast bones
become noticeably less rigid, and there may be beading at the ends of theribs. Individual hens may show temporary loss of use of the legs, with126

V I TA M I N D

recovery after laying an egg (usually shell-less) (Scott et al., 1982).
During periods of extreme leg weakness, hens show a characteristic posture that has been described as a penguin-like squat.
Of all vitamins provided in poultry feeds, vitamin D is one of two(the other is vitamin B12) that is most likely to be deficient. Typicalgrain- and soybean-based diets contain virtually none of these vitamins.
Also (as mentioned for swine above), the trend to complete confinement
will eliminate UV light as a source of the vitamin; therefore, supplemental vitamin D must be provided for all poultry operations in whichbirds remain in confinement.

HorsesDeficiency of Ca, P, or vitamin D can cause bone deformities in thehorse caused by the weight of the animal and the pull of the muscles on
weak, porous bones. Vitamin D deficiency is characterized by reducedbone calcification, stiff and swollen joints, stiffness of gait, bone deformities, frequent fractures, and reduction in serum Ca and P (NRC,
1989b). El Shorafa et al. (1979) observed that early stages of rickets inponies included decreased bone ash, decreased cortical area and bonedensity, and delayed epiphyseal closure.
Young Shetland ponies kept outdoors in Florida clearly do not needdietary vitamin D (El Shorafa et al., 1979). However, when they weredeprived of sunlight and no vitamin D was included in the diet, ponies
lost their appetite and had difficulty standing.
Grazing horses or horses that exercise regularly in sunlight and consume sun-cured hay will get the amount of vitamin D they require.
However, if exposure to sunlight is restricted by confinement, hay may
not always supply the requirement. Sunlight provides most of its antirachitic powers during the 4 hours around noon. This should be considered by those who exercise race horses briefly in the early morning
before housing them for the rest of the day (Abrams, 1978).

Other Animal Species
DOGS AND CATS
The dog was one of the first animals in which rickets was produced
experimentally. In 1922, Mellanby produced rickets in dogs by feedingthem oatmeal (NRC, 1985a). Rickets in dogs is radiographically,
histopathologically, and biochemically similar to the disorder as manifested in other animals or in human beings. Rickets with typical bone127

CHAPTER THREE

lesions is readily produced in dogs, but clinical signs are frequently confounded by simultaneous deficiency or imbalance of Ca and P (NRC,
1985a), both of which result in initial hypocalcemia.
Campbell and Douglas (1965) fed a diet of 0.5% Ca and 0.3% P,with no supplemental vitamin D, to puppies for 15 weeks without signsof rickets or osteoporosis. However, when the diet contained 0.08 to
0.10% Ca and 0.13 to 0.15% P and no supplemental vitamin D, rickets
complicated by osteoporosis was observed.
A diagnosis of rickets in a 12-week-old female St. Bernard was
attributed to an inborn error in vitamin D metabolism (Johnson et al.,1988). Physical examination revealed enlargement of the costochondraljunctions and the distal metaphyses of the radius, ulna, femur, and tibia.
When the dog was standing, its elbows were slightly abducted, and there
was mild valgus deviation of the front paws. The dog showed no lameness but was lethargic and inactive. Radiographically, the physes wereenlarged radially and axially, and metaphyseal bone adjacent to the physes was widened and cup-shaped. Serum biochemical abnormalities werehypocalcemia, hypomagnesemia, and hyperparathyroidism.
Severe rickets in kittens resulted in enlarged costochondral junctions(“rachitic rosary”), with disorganization in new bone formation andexcessive osteoid (NRC, 1986). Classic signs of rickets are rare in kittens
but can arise from queens fed vitamin D-deficient diets.
Severe rickets in kittens was produced using vitamin D-deficient
diets containing either 1% Ca and 1% P or 2% Ca and 0.65% P(Gershoff et al., 1957). Weight gain was less with the latter diet, andrickets was less severe. Rickets that developed in about 4 to 5 months
was characterized by radiographic and morphological changes that weresimilar to bone lesions observed in other species with the disease. The
cats that died during acute rickets had a lower percent of femur ash thancats supplemented with vitamin D.

FISH
Channel catfish raised in aquaria and fed a vitamin D-deficient dietfor 16 weeks showed reduced weight gain, lower body ash, lower bodyP, and lower body Ca than controls (NRC, 1993). Reduced growth withvitamin D deficiency has also been reported with juvenile lobsters, juvenile grass shrimp, blue tilapia, and trout (NRC, 1993; O’Connell and
Gatlin, 1994; Shiau and Hwang, 1994). Although effects of vitamin D3
on mineralization of hard tissues were negligible in blue tilapia, weightgains were reduced with deficiency (O’Connell and Gatlin, 1994).
Signs of vitamin D deficiency in trout include slow growth, impaired
128

V I TA M I N D

Ca homeostasis manifested by clinical signs of tetany of the white skeletal muscles, and ultrastructural changes in the white muscle fibers of themusculature (Barnett et al., 1982; NRC, 1993). No changes in bone ash
have been detected in this species of fish. A lordosis-like droopy tail syndrome observed in vitamin D-deficient trout (Barnett et al., 1982) was
suggested to be related to muscle weakness.

FOXES AND MINK
Rickets can be produced in both foxes and mink by feeding diets
having low vitamin D content and abnormal Ca-P ratios (NRC, 1982a).
Signs of rickets in growing kits are generally seen between 2 and 4
months of age.

LABORATORY ANIMALS
Rickets is classically brought about in rats by a diet lacking vitaminD, adequate in Ca, and low in P. Bones of rachitic rats show decreased
or absent calcification, with wide areas of uncalcified cartilage at thejunction of diaphysis and epiphysis. Bone ash may be less than half normal (NRC, 1995).
Guinea pigs housed without access to UV light and fed a purified
diet low in vitamin D, with 0.028% Ca and 0.2% P, did not grow normally and developed rickets. Typical lesions occurred in the zone of cartilage proliferation at the epiphyseal plate of long bones and ribs. Also,
incisors exhibited a high degree of enamel hypoplasia, while enamel anddentin were disorganized and irregular, with poor calcification (NRC,1995). Hamsters do not require dietary vitamin D for prevention of rickets when the dietary Ca-P ratio is about 2:1 and Ca is at 0.6% (NRC,1995). However, Sergeev et al. (1990) reported changes in vitamin D status of guinea pigs fed a vitamin D-deficient diet containing 6 g Ca/kg
diet and 6 g P/kg diet. Serum Ca and P concentrations were reduced,
serum alkaline phosphatase was increased, serum 25-OHD concentrations were extremely low, kidney 1-α-hydroxylase activity was morethan twice the normal concentration, and bone Ca content was aboutfour-fifths of the control concentration. Rickets may be induced in hamsters in the absence of vitamin D and when dietary Ca is 4 g/kg and P is
0.2 g/kg (NRC, 1995).

NONHUMAN PRIMATES
Softening, demineralization, and fibrous dysplasia of bone, which
are compatible with a diagnosis of rickets and responsive to vitamin D
treatment, have been observed in many species of nonhuman primates
129

CHAPTER THREE(NRC, 1978). Rickets also has been induced unintentionally in New
World primates kept under laboratory conditions when it was incorrectly assumed that they could utilize vitamin D2 as efficiently as D3 (seeNatural Sources).

HumansVitamin D deficiency in children leads to the pathological bone condition called rickets, which is characterized by disordered cartilage cell
growth and enlargement of the epiphyseal growth plates in the longbones (Fraser, 1984). There is also a prominent accumulation of unmineralized bone matrix (osteoid) on trabecular bone surfaces. Classic bone
symptoms associated with rickets, such as bowlegs (Fig. 3.7), knockknees, curvature of the spine, and pelvic and thoracic deformities, result
from the application of normal mechanical stress to demineralized bone
(Collins and Norman, 1991). Beading of the ribs, referred to as the“ricketic rosary,” is almost a constant sign after the age of 6 months,
and is caused by the swollen cartilaginous ends of the ribs. The chest
may be narrow and rather funnel-shaped, and described as a “pigeon”
chest. In severe cases, this may interfere with breathing. Enlargement ofbones, especially in the knees, wrists, and ankles, and changes in the costochondral junctions also occur. With these defects, the bones become
structurally weak, bend under the weight of the child, and are liable tofracture. Rickets also results in inadequate mineralization of tooth
enamel and dentin. If the disease occurs during the first 6 months of life,
convulsions and tetany can develop.
When humans changed from a hunting and gathering culture to an
agricultural one, they also moved from tropical and subtropical climates
to temperate zones, where houses and clothes were necessary for protection from the cold for a considerable part of the year (Harrison,1978). This limited the production of vitamin D in skin, particularly of
infants during those months when exposure of large areas of skin to sunshine was not possible. When a further change to urban living occurred,children in crowded city slums had little opportunity for exposure tosunshine. As noted earlier, this was accentuated by the Industrial
Revolution, during which the use of fossil fuels contaminated the atmosphere and thus blocked out UV radiation.
Rickets resulting from vitamin D deficiency became widespread innorthern cities, particularly in the British Isles, Europe, the UnitedStates, and Canada. The disease usually appeared in the first year of life
and was characterized by muscle weakness; deformities of long bones,
including bowed legs; knuckle-like projections along the rib cage,130

V I TA M I N D

Fig. 3.7 Child with rickets exhibits marked bowlegs. Note the angle of the feet.
(Courtesy of Alan T. Forrester, Scope Manual on Nutrition, The Upjohn
Company, Kalamazoo, Michigan.)

known as the rachitic rosary; and deformities of the pelvis, which were
often permanent. Consequences of this disease were quite profound,especially for young women, in whom a deformed pelvis would cause
difficulty with childbirth and result in a high incidence of infant andmaternal morbidity and mortality (Holick, 1987).
Osteomalacia occurs after skeletal development is complete. As inrickets, even though bone mineralization has ceased, collagen formation
continues, resulting in formation of uncalcified bone matrix. In adults,
the bones stop growing in length but are constantly remodeled. The
main symptoms of osteomalacia are muscular weakness and bone pain.
As the disease progresses, bone fractures occur.
Season and latitude greatly affect the ability of sunlight to promote
synthesis of vitamin D3 in skin; therefore, individuals from more temperate regions in the winter are more susceptible to vitamin D deficiency. Studies have indicated less vitamin D activity in northern climates131

CHAPTER THREE(see Natural Sources). In Manitoba, Canada, 43% of children and 76%
of mothers had serum 25-OHD levels below the normal range (Lebrun
et al., 1993). In a study of infant growth in two northern and two southern cities in China, infants from the more southern latitudes had thegreatest body length (Feliciano et al., 1994). Rickets is not limited tonorthern climates; the disease was established in 41 Sudanese children(El-Hag and Karrar, 1995). Possible causes in the Sudan were poorsocioeconomic background, inadequate dietary intake in both mothers
and children, prolonged breast feeding, limited sun exposure, and typeof residence.
Although it is accepted that vitamin D is absolutely essential for
growing children, it is not well appreciated that it is also essential for
maintenance of a healthy mineralized skeleton in adults. One reason forvitamin D deficiency in older populations is that individuals do not getenough exposure to sunlight. Older persons require more sunlight to get
the same vitamin effect as do young individuals (see Requirements). A
high incidence (73%) of vitamin D deficiency as evidenced by low blood
25-OHD was reported for elderly women in a nursing home in Holland(Lowik et al., 1992). The low 25-OHD concentrations were associatedwith limited exposure to UV radiation and nonusage of vitamin D supplements.
Holick (1987) observed that one of the primary causes of poor vitamin D nutrition in the elderly in the United States is a decrease in orcomplete abstinence from consumption of milk, which is the principal
food source that is fortified with vitamin D. Also, milk and milk products are the principal sources of Ca and P in many diets. In GreatBritain, where dairy products were not routinely fortified with vitaminD, it was found that 30% of women and 40% of men who incurred a
hip fracture were deficient in vitamin D (Doppelt et al., 1983).
In relation to diet, alcoholics have a high risk of vitamin D deficiency. Low plasma 25-OHD was highly related to alcohol consumption(Guilland et al., 1994). Many studies have shown the indigenous elderly population and Asian immigrants to be groups at particular risk ofvitamin D deficiency and osteomalacia. In one study, blood 25-OHD
was significantly lower in elderly Asians (21/37) and young Asians(7/17) compared with white controls (Solanki et al., 1995).
Vitamin D is involved not only in the regulation of Ca homeostasis
and bone metabolism, but in the regulation of cell proliferation, differentiation, and immune response. The extent of non–bone-related vitamin D deficiency is unknown. Vitamin D treatment to prevent various132

V I TA M I N D

types of cancer and other conditions, such as psoriasis, may be based on
the pharmacological effects of the vitamin versus a function of vitamin
D. Vitamin D has a role in heart function as evidenced by abnormal electrocardiograms (Sood et al., 1995), with congestive heart failure reported as a result of the deficiency (Brunvand et al., 1995).

Assessment of Status
Several methods have been used to assess vitamin D nutritional status of individuals. Poor production rates in livestock as well as boneabnormalities in both animals and humans are the chief indications thatvitamin D deficiency is substantially advanced. The incomplete calcification of the skeleton is easily detectable with x-rays, but, like other production-related signs, would not be specific for vitamin D deficiency versus other nutrient inadequacies.
Low serum Ca levels in the range of 5 to 7 mg/100 ml and highserum alkaline phosphatase activity can be used to diagnose rickets andosteomalacia. Also, a marked reduction in circulating 1,25-(OH)2D levels in individuals with osteomalacia has been reported. The plasma concentration of 25-OHD is related to and therefore is an index of the vitamin D supply for both animals and humans. The concentration of 25OHD in plasma, which is the principal form of vitamin D in plasma, isabout 10 times and 500 to 1,000 times higher than those of 24,25(OH)2D and 1,25-(OH)2D, respectively (Fraser, 1984).
The circulating concentration of 25-OHD is a good index for determining the cumulative effect of sunlight on synthesis of vitamin D3 inskin, and dietary sources of the vitamin. In plasma of the vitamin D-deficient child, a low concentration of 25-OHD (less than 5 ng/ml) has beenfound (Fraser, 1984). Oldham et al. (1980) fed a vitamin D-deficient dietto dogs. This diet required at least 3 months to depress the serum Ca
approximately 2 mg/dL while lowering the serum 25-(OH)2D3 level toless than 0.4 ng/ml (30 to 60 ng/ml is the normal value for humans).
Toxicity caused by excess vitamin D administration is also associated with plasma 25-OHD; concentrations of more than 400 ng/ml havebeen reported (Hughes et al., 1976). Patients suffering from hypervitaminosis D have been shown to exhibit a 15-fold increase in plasma 25OHD concentration compared with normal individuals.
Osteocalcin can also be used to evaluate bone mineralization. Carter
et al. (1996) have suggested that serum osteocalcin and 1,25-(OH)2D
are better predictors of bone mineralization and/or turnover in pigs thanis serum alkaline phosphatase.

133

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SUPPLEMENTATION
For animals that are kept indoors or live in climates where the sunlight is not adequate for vitamin D production, the vitamin D content of
the diet becomes important. In modern poultry, swine, and dairy operations, animals are often entirely without sunlight. Under these intensive
conditions, supplemental vitamin D must be provided. It seems that supplementation is important for dogs and cats since studies have shownconclusively that neither dogs nor cats receive a significant benefit fromsynthesis of vitamin D in the skin through exposure to UV irradiation(How et al., 1994a,b, 1995).
Animals on pasture during the summer never suffer from the lack of
the antirachitic factor even though their diet may be practically devoid of
it. In winter, however, animals are outside only a part of the time, there
are generally fewer sunny days, and the sunlight that reaches the animal
is much less effective than in summer. In winter at far southern and northern latitudes, it is unsafe to rely on exposure to sunlight to provide theantirachitic factor as no significant production of vitamin D3 in skinoccurs (Webb et al., 1988). Milk is not especially rich in vitamin D, but
young animals can obtain adequate amounts by skin irradiation if
exposed to the sunlight for 1 to 2 hours per day. Sun-cured forage is thebest natural source of vitamin D. Even silages generally have dead (thussun-cured) leaves that will provide some vitamin D to growing animals
housed indoors. However, it is probably prudent to provide a vitamin D
supplement to young calves in their milk replacer and starter diets until
they are turned out or are consuming adequate forage (e.g., 6 to 8 weeks).
Special Ca and P supplementation is required for high-producing
dairy cows to prevent parturient paresis (milk fever). Parturient paresis
can be prevented effectively by feeding a prepartum diet low in Ca andadequate in P. Prepartal low-Ca diets are associated with increased plasma PTH and 1,25-(OH)2D2 and 1,25-(OH)2D3 concentrations during
the prepartal period. Green et al. (1981) suggested that these increased
PTH and 1,25-(OH)2D concentrations resulted in “prepared” and effective intestinal and bone Ca homeostatic mechanisms at parturition thatprevented parturient paresis.
Supplemental vitamin D has been used to prevent parturient paresis
in dairy cows for a number of years. Treatment with high levels of vitamin D has been successful, but toxicity problems have sometimes resulted, and for some animals, the disease has been induced by the treatment.
Because of the extreme toxicity of vitamin D3 in pregnant cows, and the134

V I TA M I N D

low margin of safety between doses of vitamin D3 that prevent milk
fever and doses that induce milk fever, Littledike and Horst (1982) concluded that vitamin D3 cannot be used practically to prevent milk feverwhen injected several weeks prepartum. However, a later report fromthe same laboratory provided data suggesting that injection of 24-F1,25-(OH)2D3 (fluoridation at the 24R position) delivered at 7-day
intervals prior to parturition can effectively reduce incidence of parturient paresis (Goff et al., 1988). This compound, however, never advanced
beyond the experimental stage. An analogue that has been marketed in
Israel and used with varying degrees of success was 1α-hydroxycholecalciferol [1α(OH)D3] (Sachs et al., 1977).
This analogue was developed as a precursor in the chemical synthesis of 1,25-(OH)2D3 and, fortunately, can be activated by the body following 25-hydroxylation in the liver to form 1,25-(OH)2D3. Use of thisanalogue, however, shared the disadvantages of earlier compounds:
hypercalcemia was induced, and the endogenous synthesis of 1,25(OH)2D3 was inhibited. Hodnett et al. (1992) used a combination of 25OHD3 plus 1α−hydroxycholecalciferol to reduce parturient paresis indairy cows fed high dietary Ca. The incidence of the disease was reduced
from 33 to 8%. Supplementation of 1α-hydroxycholecalciferol, which isless costly to produce than 1,25-(OH)2D3, has improved phytate P utilization in poultry but not swine (Biehl and Baker, 1997a,b).
Anion-cation balance of prepartum diets (sometimes referred to asacidity or alkalinity of a diet) also can influence the incidence of milk
fever (Oetzel et al., 1988; Gaynor et al., 1989). Diets high in cations,especially sodium and potassium, tend to induce milk fever, but thosehigh in anions, primarily chlorine and sulfur, can prevent milk fever. The
incidence of milk fever depends on the abundance of the cations Na+
and K+ relative to the anions Cl- and SO42-. This concept is now generally referred to as the cation-anion difference (CAD). Because mostlegumes and grasses are high in potassium, many of the commonly usedprepartum diets are alkaline. Addition of anions to a prepartal diet isthought to induce metabolic acidosis in the cow, which facilitates bone
Ca resorption and intestinal Ca absorption (Lomba et al., 1978; Fredeen
et al., 1988; Horst et al., 1997). Diets higher in anions increase osteoclastic bone resorption and synthesis of 1,25-(OH)2D3 in cows (Goff etal., 1991a). Both of these physiological processes are controlled by PTH.
Workers at the Rowett Research Institute (Abu Damir et al., 1994) havealso reported that 1,25-(OH)2D3 production is enhanced in cows fed
acidifying diets. Commercial preparations of HCL mixed into common
feed ingredients as a premix could offer an inexpensive and palatable
135

CHAPTER THREE

alternative to anionic salts as a means of controlling the incidence of
milk fever in dairy cows (Goff and Horst, 1998).
Collectively, these data suggest that a major underlying cause of
milk fever is metabolic alkalosis, which causes an inability of cow tissues
to respond adequately to PTH (Horst et al., 1997). This lack of response
in turn reduces the ability of the cow to draw on bone Ca stores, andreduces production of the second Ca-regulating hormone, 1,25-(OH)2D,
which is needed for active transport of Ca within the intestine. The presumption is that metabolic alkalosis somehow disrupts the integrity of
PTH receptors on target tissues. Low CAD diets prevent metabolic alkalosis, increasing target tissue responsiveness to PTH, which controls
renal 1α-hydroxylase and resorption of bone Ca.
Several options exist regarding methods for the control of milk fever(Horst et al., 1997). The current understanding of the CAD concept suggests that milk fever could be managed more effectively if dietary K werereduced (Goff and Horst, 1997). Calcium chloride has been used toreduce blood pH. This reduction is beneficial, but excessive oral Ca
chloride can induce metabolic acidosis (Goff and Horst, 1994), which
can cause inappetence at a time when feed intake is already compromised. Calcium propionate treatment has been beneficial in reducing
subclinical hypocalcemia in all trials and reduced the incidence of milk
fever in a herd having a problem with milk fever (Goff et al., 1996).
Due to lack of vitamin D in feeds and management systems without
direct sunlight, modern livestock operations must provide a supplemental source of the vitamin. Vitamin D is available in two forms: vitamin
D3 of animal origin and vitamin D2 of plant origin. Vitamin D3 has been
the primary source of supplemental vitamin D for domestic animals,
whereas vitamin D2 has been the chief source of supplemental vitamin
D in foods and pharmaceuticals. Vitamin D3 is commercially available
as a resin, usually containing 24 to 26 million IU per gram, for use as
the vitamin D source in various vitamin products. Vitamin D3 products
for feed include gelatin beadlets (with vitamin A), oil dilutions, oil
absorbates, emulsions, and spray- and drum-dried powders. Test results
have shown that the gelatin beadlet offers optimum vitamin D3 stability. A commercially available vitamin product containing stabilized,
high-purity vitamin D for feed or drinking water use should be used toensure the vitamin D levels needed to prevent deficiency and allow optimum performance. Continued irradiation eventually destroys the vitamin D produced, but the chief cause of loss from foods is doubtless oxidation, as was recognized long ago (Fritz et al., 1942). Higher vitaminD levels in freeze-dried fish meals suggest less destruction during drying,136

V I TA M I N D

possibly because of decreased atmospheric oxygen (Scott and Latshaw,
1994).
Stabilization of the vitamin can be achieved by (1) rapid compression of the mixed feed, for example, into cubes, so that air is excluded,(2) storing feed under cool, dry, dark conditions, (3) preventing close
contact between the vitamin and potent metallic oxidation catalysts, forexample, manganese, and (4) including natural or synthetic antioxidants
in the mix. The vitamin can also be protected by enclosing it in tough,gelatin beadlets. But all stability is relative; warmth and moisture willsoften gelatin, while temperatures below about 10oC cause it to become
hard, brittle, and friable (Abrams, 1978).
Pure vitamin D3 crystals or vitamin D3 resin is very susceptible todegradation upon exposure to heat or contact with mineral elements. In
fact, the resin is stored under refrigeration with nitrogen gas. Dry, stabilized supplements retain potency much longer and can be used in highmineral supplements. It has been shown that vitamin D3 is much morestable than D2 in feeds containing minerals. When the unprotected vitamin is thinly spread over the surface of free-flowing powders, as in someof the mineral components of compound animal feeds, its life in storage
may be no more than 1 month. In complete feeds and mineral-vitaminpremixes, Schneider (1986) reported activity losses of 10 to 30% after
either 4 or 6 months of storage at 22oC.
Stability of dry vitamin D supplements is affected most by high temperature, high moisture content, and contact with trace minerals such as
ferrous sulfate, manganese oxide, and others. Hirsh (1982) reports theresults of a “conventional” or nonstabilized vitamin D3 product being
mixed into a trace mineral premix or into animal feed and stored atambient room temperature (20 to 25oC) for up to 12 weeks. The mash
feed had lost 31% of its vitamin D activity after 12 weeks, and the tracemineral premix had lost 66% of its activity after only 6 weeks in storage.
In addition to providing supplemental vitamin D in feed and water,
injectable sources are available. Parenteral vitamin D3 treatment of sows
before parturition provided an effective means of supplementing piglets
with vitamin D3 (via the sow’s milk) and its dehydroxymetabolites byplacental transport (Goff et al., 1984).
Naturally occurring sources of vitamin D in feeds must likewise beprotected from loss. Poor handling of hay, which can otherwise be an
important source of vitamin D and various other nutrients for forageconsuming animals, can lead to extensive fragmentation and loss of theleaf, which is much richer in vitamin D than the stem. Animals fed on137CHAPTER THREE

grain, silage, or hay that is poorly produced or stored are extremely
liable to have dietary deficiency of the vitamin (Abrams, 1978).
Cost of vitamin D supplementation to livestock diets is nominal(Rowland, 1982). In contrast, the cost of not adding enough vitamin D
to prevent deficiencies is very high. Supplemental levels of vitamin D
administered to livestock through the feed should be adjusted to provide
the margin of safety needed to offset the factors influencing vitamin D
needs. This is important to prevent deficiency and allow optimum performance.
Rowland (1982) notes that diets for young, rapidly growing chickens must contain liberal amounts of vitamin D to prevent field problems.
He further observed that the NRC level (200 IU/kg of feed) for young
chickens is extremely unrealistic for broilers. Even under low-stressresearch conditions, three to five times the NRC level is required to support maximum weight gain of broilers, and under commercial conditions, 10 times the NRC level is prudent for broilers fed under field conditions. The NRC vitamin D levels for laying hens and turkeys are somewhat more realistic, with a factor of five times the NRC level generally
supporting optimum performance and providing some margin of safety(Rowland, 1982). The most logical approach is to adjust supplemental
vitamin D levels to expected production conditions.
Besides inadequate quantities of dietary vitamin D, deficiencies may
result from (1) errors in vitamin addition to diets, (2) inadequate mixing
and distribution in feed, (3) separation of vitamin D particles after mixing, (4) instability of the vitamin content of the supplement, or (5) excessive length of storage of diets under environmental conditions causing
vitamin D loss (Hirsch, 1982).
Supplementation considerations are dependent on other dietary
ingredients. The requirements for vitamin D3 are increased several-foldby inadequate levels of Ca and/or P or by improper ratios of these twoelements in the diet (see Requirements). Vitamin D treatment considerably enhances Ca and P utilization in pigs fed on a phytate-P diet but
does not completely overcome the negative effects of phytate feeding on
mineral metabolism (Pointillart et al., 1986). A number of reports haveindicated that molds in feeds interfere with vitamin D (Cunha, 1977);
for example, when corn contains the mold Fusarium roseum, a metabolite of this mold prevents vitamin D3 in the intestinal tract from being
absorbed by the chick. Other molds may also be involved, and they
result in a large percentage of birds with bone disorders. A number offlocks have been successfully treated by adding water-dispersible forms

138

V I TA M I N D

of vitamin D to drinking water at three to five times the normally recommended vitamin D levels.
Other factors that influence vitamin D status are diseases of theendocrine system, intestinal disorders, liver malfunction, kidney disorders, and drugs. Liver malfunction limits production of the active formsof the vitamin, while intestinal disorders reduce absorption. Persons
with kidney failure are unable to synthesize 1,25-(OH)2D, and forpatients in renal dialysis who are waiting for a compatible kidney transplant donor, 1,25-(OH)2D has been found to alleviate the painful andserious bone disease associated with chronic renal failure. The prolonged use of anticonvulsant drugs can result in an impaired response tovitamin D (Collins and Norman, 1991), and Cunha (personal communication, 1987) suggested the possibility that livestock with certain diseases or heavy infestation of internal parasites might be unable to synthesize the metabolically active forms of vitamin D as a result of liver orkidney damage. Unknown factors in feeds may increase vitamin D
requirements. For example, there is evidence of a factor in rye and insoybean fractions that can produce malabsorption of this vitamin in theintestine (Mac-Auliff and McGinnis, 1976).
Widespread fortification of human diets with vitamin D and provision of oral supplements over the past 60 years have greatly reduced theincidence of rickets in children. However, most of the world’s population still depends on exposure to sunlight for their vitamin D nutritional needs because very few natural foods contain sufficient quantities tomeet the daily requirement. It has been assumed that in countries where
food is fortified with vitamin D, such as the United States, exposure to
sunlight is no longer necessary for vitamin D nutrition. Although thismay be true for young children who drink milk fortified with vitamin D,
many elderly persons who do not drink milk or take a vitamin D supplement are prone to vitamin D deficiency when they are not exposed to
sufficient sunlight. Among the fortified foods are milk (both fresh andevaporated), margarine and butter, cereals, and chocolate mixes. Milk is
fortified to supply 400 IU of vitamin D per quart, and margarine usually contains 4,400 or more IU per kilogram (Collins and Norman, 1991).
The major source of vitamin D for most humans is casual exposure
to sunlight. It is estimated that more than 80 to 90% of the body’s
requirement for vitamin D comes from this source (Holick, 1994).
Several factors can affect the cutaneous synthesis of vitamin D3 (seeRequirements). Anything that limits the amount of solar ultraviolet B(UVB) photons to reach the skin’s surface and penetrate the viable epi-

139

CHAPTER THREE

dermis can significantly affect this vital photosynthetic process. Skin pigment, clothing, and topically applied sunscreens absorb UVB photons
and, therefore, can significantly diminish the synthesis of vitamin D3(Holick, 1994). The topical application of a sunscreen with a sun protection factor of 8 can almost completely eliminate the cutaneous production of vitamin D3 and cause vitamin D deficiency (Matsuoka et al.,1988). This is probably not a logical concern as sufficient summer sunlight is received, probably through both the sunscreen itself and the lackof total skin cover at all times, to allow adequate vitamin D productionin people who are advised to use sunscreens regularly (Marks et al.,1995). Season, latitude, and time of day can significantly affect the cutaneous production of vitamin D3.
Many bone problems in humans are the result of Ca and/or vitamin
D deficiencies. These problems are most severe in young children, adolescents, and elderly persons (manifested as hip fracture and fractures ofthe vertebrae). Bone problems are generally preventable by adequate
dietary or supplementary Ca and vitamin D and exercise at a young age,with the goal of developing peak bone mass prior to advancing age.
Optimal intake of both Ca and vitamin D may be an easily implemented strategy to maintain existing bone mass and reduce the risk of fracture in older men and women (O’Brien, 1998).
Epidemiological studies have suggested that sunlight deprivation,and the associated reduction in the circulating levels of vitamin D3 derivatives, may lead to an increased incidence of carcinomas of the breast,colon, and prostate (Studzinski and Moore, 1995). Likewise, sunlight(vitamin D) may be a protective factor in ovarian cancer mortality(Lefkowitz and Garland, 1994).
The Food and Drug Administration does not permit 1,25-(OH)2D
to be added to feeds. However, this hormone or its analogs have proveneffective in human medicine in treating psoriasis (Bikle, 1995), colorectal carcinoma (Cross et al., 1995), leukemia (Jung et al., 1994), breast
cancer (James et al., 1994), and prostate cancer (Skowronski et al.,1995). This function relates to 1,25-(OH)2D modulating cell growthand differentiation, including that of malignant cells.

TOXICITY
Excess consumption of vitamin A results in toxicity. After vitaminA, vitamin D is the vitamin most likely to be consumed in concentrations
toxic to both humans and livestock. Excessive intake of vitamin D produces a variety of effects, all associated with abnormal elevation of140

V I TA M I N D

blood Ca. Elevated blood Ca is caused by greatly stimulated bone
resorption, as well as increased intestinal Ca absorption.
The main pathological effect of ingestion of massive doses of vitamin D is widespread calcification of soft tissues. Pathological changes inthese tissues are observed to be inflammation, cellular degeneration, andcalcification. Diffuse calcification affects the joints, synovial membranes, kidney, myocardium, pulmonary alveoli, parathyroids, pancreas,
lymph glands, arteries, conjunctivae, and cornea. More advanced casesinterfere with cartilage growth. The abnormal calcification is grossly
observed as a whitish chalky material, and kidney insufficiency is themost critical development of these processes. Initial kidney damage isdue to Ca deposition in distal tubules, causing inflammation and later
obstruction, which in turn causes hypertension and related pathology.
As would be expected, the skeletal system undergoes a simultaneous
demineralization that results in the thinning of bones.
Other common observations of vitamin D toxicity are anorexia (lossof appetite), extensive weight loss, elevated blood Ca, and lowered
blood phosphate. Many investigators have described the clinical signs ofhypervitaminosis D in mammals (NRC, 1987). Cows receiving 30 million IU of vitamin D2 orally for 11 days developed anorexia, reduced
rumination, depression, premature ventricular systoles, and bradycardia. Toxicosis in monkeys resulted in weight loss, anorexia, elevatedblood urea nitrogen, diarrhea, anemia, and upper respiratory infections.
In pigs, signs of toxicity were anorexia, stiffness, lameness, arching of
the back, polyuria, and aphonia. Dogs receiving toxic concentrations ofvitamin D exhibited anorexia, polyuria, bloody diarrhea, polydipsia,
prostration, and excessive calcification of the lungs (Morgan, 1947),
while cats receiving excess vitamin D exhibited heavy calcification ofarteries and the adrenals (Suter, 1957). Symptoms of vitamin D intoxication for humans include hypercalcemia, hypercalciuria, anorexia, nausea, vomiting, thirst, polyuria, muscular weakness, joint pains, diffusedemineralization of bones, and disorientation (Collins and Norman,
1991).
In poultry, excess vitamin D elevates 1,25-(OH)2D with hypercalcemia and soft tissue mineralization (NRC, 1994). Leg problems may
arise with growing birds because of bone Ca loss (Cruickshank and Sim,1987), but few obvious changes occur with hens other than a general
depression in performance (Ameenuddin et al., 1986). Toxic levels ofvitamin D may be transferred into the egg to create similar problems forthe embryo; however, the hypercalcemia occurs from shell resorption,and bone mineralization is enhanced (Narbaitz and Fragiskos, 1984).
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Harrington and Page (1983) compared toxicity of D2 to D3 in horses. Signs of toxicity included weight loss, hypercalcemia, hyperphosphatemia, and cardiovascular calcinosis. However, the signs of illness,deviations of blood chemistry from normal, and severity of tissue
pathology, were much more pronounced for vitamin D3.
Severity of the effects and pathogenic lesions in vitamin D intoxication depend on such factors as the type of vitamin D (D2 versus D3), thedose, the functional state of the kidneys, and the composition of the diet
(NRC, 1987). Studies indicate that vitamin D3 is 10 to 20 times more
toxic than vitamin D2. When equal amounts of vitamin D3 and vitamin
D2 are provided together in diets of mammals, the predominant circulating form of the vitamin is usually 25-OHD3 rather than 25-OHD2.
Similarly, in toxicity experiments in which vitamin D2 was less toxic
than vitamin D3, the metabolite 25-OHD2 was found to be present at
lower plasma concentrations than 25-OHD3 (Harrington and Page,
1983).
Vitamin D toxicity is enhanced by elevated supplies of dietary Ca
and P and is reduced when the diet is low in Ca. Toxicity is also reduced
when the vitamin is accompanied by intakes of vitamin A or by thyroxin injections (Payne and Manston, 1967). Route of administration also
influences toxicity. Parenteral administration of 15 million IU of vitamin
D3 in a single dose caused toxicity and death in many pregnant dairy
cows (Littledike and Horst, 1982). On the other hand, oral administration of 20 to 30 million IU of vitamin D2 daily for 7 days resulted in little or no toxicity in pregnant dairy cows (Hibbs and Pounden, 1955).
Rumen microbes are capable of metabolizing vitamin D to an inactive
compound, which may partially explain the difference in toxicity
between oral and parenteral vitamin D. The toxic dose of vitamin D isquite variable, with an important factor being duration of intake, as thisis a cumulative toxicity. However, there is marked variation among individuals in tolerance to excessive doses of vitamin D, the mechanism ofwhich is unknown. Some species are less affected by toxicity due to vitamin D metabolism differences. For example, in the hooded seal, there isincreased conversion of 25-OHD to 24,25-(OH)2D and a high capacityfor vitamin D storage in its large blubber mass, which provides resistance for this species to toxicity (Keiver et al., 1988).
Although it is usually assumed that living plants do not contain vitamin D2, some plants contain compounds that have vitamin D activity(Mello and Habermehl, 1995). Grazing animals in several parts of theworld develop calcinosis, a disease characterized by the deposition of Ca
salts in soft tissues (Carrillo, 1973; Morris, 1982). The ingestion of the142

V I TA M I N D

leaves of the shrub Solanum malacoxylon by grazing animals causes
enzootic calcinosis in Argentina and Brazil, where the disease is referred
to as enteque seco and espichamento, respectively. As few as 50 fresh
leaves per day (200 g of fresh leaves per week) over a period of 8 to 20
weeks are enough for the disease to develop in cows (Fig. 3.8) (Okada
et al., 1977).
Another solanaceous plant, Cestrium diurnum (a large ornamental
plant), causes calcinosis in grazing animals in Florida, while the grass
Trisetum flavescens is the causative agent in the Alpine regions in
Europe. Solanum torvum is suspected of causing calcinosis in cattle in
Papua, New Guinea. A condition known as “humpy-back,” in whichclinical symptoms reminiscent of calcinosis occur, may be caused bysheep grazing the fruits of S. esuriale in Australia. In Jamaica,“Manchester wasting disease” and in Hawaii, “Naalehu disease” areconditions seen in cattle that are virtually identical to enteque seco in
relation to clinical and pathological signs (Wasserman, 1975; Arnold
and Fincham, 1997).
The calcinogenic factor in S. malacoxylon and C. diurnum is awater-soluble glycoside of 1,25-(OH)2D3 (Wasserman, 1975). The digestive system of the animal releases the sterol, which promotes a massive
increase in the absorption of dietary Ca and P such that accommodationof these elements by the normal physiological processes is ineffective,
and soft tissue calcification results. Vitamin D3 has been identified in T.
flavescens but at concentrations that would not be calcinogenic.
Evidence is emerging that the grass may also contain 1,25-(OH)2D3 or asubstance able to mimic its actions as well as an aqueous-soluble factor
that promotes phosphate absorption.
During development of plant-induced calcinosis diseases, destruction of connective tissues occurs, and this precedes mineralization inwhich magnesium is involved as well as Ca and phosphate. Clinical signsof the disease are stiffened and painful gait, with progressive weight loss.
If the animals are removed in the early stages from the affected areas,
they recover quickly, but they may die if they are not removed. In
advanced cases, joints cannot be extended completely, and animals tend
to walk with an arched back, carrying the weight on the forepart of thehooves.
Animals with calcinosis may also show signs of acute cardiac andpulmonary insufficiency. Postmortem examination shows widespreadmetastatic calcification of the vascular system and soft tissues. The heart
and aorta exhibit the most marked effects, and calcification in the lungdevelops later than in the vascular system. Cartilage of the appendicular
143

A

B
Fig. 3.8 Vitamin D toxicity (enteque seco) in Argentina: (A) Cow that had consumed the shrub Solanum malacoxylon; (B) Calcium deposits in soft tissue.
(Courtesy of Bernardo Jorge Carrillo, CICV, INTA, Castelar, Argentina.)

144

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skeleton is eroded in advanced cases, with the joints almost denuded of
cartilage. The kidney may also show some calcification. Animals grazing in areas where the disease occurs show high serum levels of Ca andinorganic P. All clinical signs are similar to those found in vitamin D
intoxication (Wolker and Carrillo, 1967).
It is clear that the calcinogenic plants are economically important,particularly because of their toxic effects. which result in enormous losses in meat and milk production (Morris, 1982). In some fields inArgentina, between 10 and 30% of cattle show signs of enteque seco,and S. malacoxylon is now regarded as one of the most important poisonous plants of that country. A survey conducted in Bavarian slaughterhouses revealed mineral deposits in soft tissues in 22 to 52% of cattle from the south of Bavaria that had been grazing at an altitude above
500 m. Trisetum flavescens is also an important toxic plant in these
alpine pastures. In 1977, it was estimated that the annual loss in revenue
resulting from S. malacoxylon-induced calcinosis in Argentina was ofthe order of $10 million. No assessment of the economic losses arisingfrom plant-induced calcinosis is available for any other country.
With the exception of grazing animals consuming certain plants containing 1,25-(OH)2D3 in several parts of the world, excessive amounts ofvitamin D are not available from natural sources. Although certain fish
liver and body oils are good sources of vitamin D, concentrations are notso high that toxic amounts of vitamin D would likely be ingested.
Therefore, danger of vitamin D toxicosis is from dietary supplementation. Extensive whole-body irradiation with UV light will not result invitamin D toxicity. Vitamin D toxicosis from dietary sources can result in
400 ng/ml of plasma 25-OHD, while extensive UV light exposure canresult in only 40 to 80 ng of this transport form of vitamin D (Davie andLawson, 1980). Therefore, supplying vitamin D by mouth bypasses protective mechanisms that prevent excessive 25-OHD formation.
For most species the presumed maximum safe level of vitamin D3
for long-term feeding conditions (more than 60 days) is 4 to 10 times therecognized dietary requirement (NRC, 1987). Catfish and rainbow
trout, on the other hand, can tolerate as much as 50 and 500 times theirrequirements, respectively. Under short-term feeding conditions (lessthan 60 days), most species can tolerate as much as 100 times theirapparent dietary requirements. Table 3.3 provides recommendations forsafe upper dietary levels of vitamin D3 for animals.
Rodents are sensitive to vitamin D toxicity and have an LD50 of
43.6 mg/kg body weight for cholecalciferol compared to 88 and 200

145

CHAPTER THREE■ Table 3.3

Estimation of Safe Upper Dietary Levels of Vitamin D3 (IU/kg in Diet) for
Animals
Exposure Time

Animal
Birds
ChickenJapanese quail
TurkeyCow
Fish
CatfishRainbow trout

Dietary
Requirement< 60 days> 60 days

200
1,200
900

40,000120,000
90,000

2,800
4,700
3,500

300

25,000

2,200

1,000
1,800

20,0001,000,000

Horse

400

Sheep

275

25,000

2,200
2,200

Swine

220

33,0002,200

Source: Modified from NRC (1987).

mg/kg in dogs and humans, respectively (Tindall, 1985). Cholecalciferol
is marketed as an effective rodenticide, with low oral toxicity to nonrodent species. However, cholecalciferol rodenticides have caused significant toxicity in dogs at a fraction of the manufacturer’s reported LD50
for dogs, 88 mg pure cholecalciferol per kilogram of body weight(Garlock et al., 1991). These researchers described a dog that consumed
pure cholecalciferol in an estimated amount of 1.5 to 3.0 mg/kg bodyweight. Clinical signs most commonly associated with the resultant
hypercalcemia were polyuria, polydipsia, depression, anorexia, weakness, and vomiting.
In the early stages of vitamin D intoxication, the effects are usually
reversible. Treatment consists merely of withdrawing vitamin D and perhaps reducing dietary Ca intake until serum Ca levels fall. However, it isusually not immediately successful because of the long plasma half-lifeof vitamin D (5 to 7 days) and 25-OHD (20 to 30 days). This is in contrast to the short plasma half-life of 1α-OHD3 (1 to 2 days) and 1,25(OH)2D3 (4 to 8 hours). Sodium phytate, an agent that reduces intestinal Ca absorption, has also been used successfully in vitamin D toxicity
management in monogastrics. This treatment would be of little benefit
to ruminants because of the presence of rumen microbial phytases.
There have also been reports that calcitonin, glucagon, and glucocorticoid therapy reduces serum Ca levels resulting from vitamin D intoxication (NRC, 1987).
146

V I TA M I N D■ REFERENCES
Abrams, J.T. (1978). In Handbook Series in Nutrition and Food, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 2, p. 179. CRC Press, Boca
Raton, Florida.
Abu Damir, H., Phillippo, M., Thorp, B.H., Milne, J.S., Dick, L., and Nevison,
I.M. (1994). Res. Vet. Sci. 56, 310.
Ameenuddin, S., Sunde, M.L., DeLuca, H.F., and Cool, M.E. (1986). Brit. Poult.
Sci. 27, 671.
Arneil, G.C. (1975). Proc. Nutr. Sci. 34, 101.
Arnold, R.M., and Fincham, I.H. (1997). Trop. Anim. Health Prod. 29, 174.
Baker, A.R., McDonnell, D.P., Hughes, M., Crisp, R.M., Manglesdorf, D.J.,
Haussler, M.R., Pike, J.W., Shine, J., and O’Malley, B.W. (1988). Proc.
Natl. Acad. Sci. 85, 3294.
Barnett, B.J., Young, Cho, C., and Slinger, S.J. (1982). J. Nutr. 112, 2011.
Biehl, R.R., and Baker, D.H. (1997a). J. Nutr. 127, 2054.
Biehl, R.R., and Baker, D.H. (1997b). J. Anim. Sci. 75, 2986.
Bikle, D.D. (1995). J. Nutr. 125, 1709S.
Blunt, J.W., DeLuca, H.F., and Schnoes, H.K. (1968). Biochemistry 7, 3317.
Bordier, P., Rasmussen, H., Marie, P., Miravet, L., Gueris, J., and Ryckwaert, A.
(1978). J. Clin. Endocrinol. Metab. 46, 284.
Braithwaite, G.D. (1974). Br. J. Nutr. 31, 319.
Braithwaite, G.D., and Riazuddin, S.H. (1971). Br. J. Nutr. 26, 215.
Braithwaite, G.D., Glascock, R.F., and Riazuddin, S.H. (1969). Br. J. Nutr. 23,
827.
Braithwaite, G.D., Glascock, R.F., and Riazuddin, S.H. (1972). Br. J. Nutr. 27,
417.
Bronner, F. (1987). J. Nutr. 117, 1347.
Bronner, F., and Stein, W.D. (1995). J. Nutr. 125, 1987S.
Brunvand, L., Haga, P., Tangsrud, S.E., and Haug, E. (1995). Acta Paediatrica
84, 106.
Cade, C., and Norman, A.W. (1986). Endocrinology 119, 84.
Campbell, J.R., and Douglas, T.A. (1965). Br. J. Nutr. 19, 339.
Carrillo, B.J. (1973). Rev. Invest. Agropecu. Serv. 4, Patol. Anim. 10, 65.
Carter, S.D., Cromwell, G.L., Combs, T.R., Colombo, G., and Fanti, P. (1996).
J. Anim. Sci. 74, 2719.
Chen, Jr., P.S., and Bosmann, H.B. (1964). Science 150, 19.
Church, D.C., and Pond, W.G. (1974). Basic Animal Nutrition and Feeding.
Albany Printing, Albany, New York.
Collins, E.D., and Norman, A.W. (1991). In Handbook of Vitamins (L.
Machlin, ed.) p. 59. Marcel Dekker, New York.
Colston, K., Colston, M.J., and Feldman, D. (1981). Endocrinology 108, 1083.
Craig, J.F., and Davis, G.O. (1943). J. Comp. Pathol. Ther. 53, 196.
Cross, H.S., Hulla, W., Tong, W., and Peterlik, M. (1995). J. Nutr. 125, 2004S.
Cruickshank, J.J., and Sim, J.S. (1987). Avian Dis. 31, 332.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Davie, M., and Lawson, D.E.M. (1980). Clin. Sci. 58, 235.
147

CHAPTER THREE

DeLuca, H.F. (1979). Nutr. Rev. 37, 161.
DeLuca, H.F. (1990). In Proc. 1990 National Feed Ingredients Association
Nutrition Institute: Developments in Vitamin Nutrition and HealthApplications, Kansas City, Missouri. National Feed Ingredients Association(NFIA), Des Moines, Iowa.
DeLuca, H.F. (1992). Ann. N.Y. Acad. Sci. 669, 59.
DeLuca, H.F., and Zierold, C. (1998). Nutr. Rev. 56, S 4.
Doppelt, S.H., Neer, R.M., Daly, M., Bourvet, L., Schiller, A., and Holick, M.F.
(1983). Orthop. Trans. 7, 512.
Duncan, C.W. (1944). J. Dairy Sci. 27, 636.
Dzanis, D.A., Corbellini, C.N., Krook, L., and Kallfelz, F.A. (1992). J. Nutr.
121, 11S.
Eisman, J.A. (1998). Nutr. Rev. 56, S 22.
Elaroussi, M.A., Uhland-Smith, A., Hellwig, W., and DeLuca, H.F. (1994).
Biochem. Biophys. Acta 1192, 1.
El-Hag, A.I., and Karrar, Z.A. (1995). Annals Trop. Pediat. 15, 69.
Elliot, W., and Crichton, A. (1926). J. Agar. Sci. 16, 65.
El Shorafa, W.M., Feaster, J.P., Ott, E.A., and Asquith, R.L. (1979). J. Anim. Sci.
48, 882.
Feliciano, E.S., Ho, M., Specker, B.L., Falciglia, G., Shui, Q.M., Yin, T.A., andChen, X.C. (1994). J. Trop. Pediat. y40, 162.
Fleet, J.C. (1999). Nutr. Rev. 57, 60.
Franklin, M.C., and Johnstone, I.L. (1948). Maintenance of Serum Calcium
Level by Calcium Supplements to the Diet, p. 63. C.S.I.R.O., Melbourne,
Florida.
Fraser, D.R. (1984). In Nutrition Reviews, Present Knowledge in Nutrition (R.E.
Olson, ed.), p. 209. Nutrition Foundation, Washington, D.C.
Fredeen, A.H., DePeters, E.J., and Baldwin, R.L. (1988). J. Anim. Sci. 66, 159.
Fritz, J.C., Archer, W.F., and Barker, D.K. (1942). Poult. Sci. 21, 361.
Garabedian, M., Tanaka, Y., Holick, M.F., and DeLuca, H.F. (1974).
Endocrinology 94, 1022.
Garlock, S.M., Matz, M.E., and Shell, L.G. (1991). J. An. Anim. Hosp. Assoc.
27, 356.
Gaynor, P.J., Mueller, F.J., Miller, J.K., Ramsey, N., Goff, J.P., and Horst, R.L.
(1989). J. Dairy Sci. 72, 2525.
Gershoff, S.N., Legg, M.A., O’Connor, F.J., and Hegsted, D.M. (1957). J. Nutr.
63, 79.
Ghazarian, J.G., Jefcoate, C.R., Knutson, J.C., Orme-Hohnson, W.H., andDeLuca, H.F. (1974). J. Biol.Chem. 249, 3026.
Goff, J.P., and Horst, R.L. (1994). J. Dairy Sci. 77, 1451.
Goff, J.P., and Horst, R.L. (1997). J. Dairy Sci. 80, 176.
Goff, J.P., and Horst, R.L. (1998). J. Dairy Sci. 81, 2874.
Goff, J.P., Horst, R.L., and Littledike, E.T. (1984). J. Nutr. 114, 163.
Goff, J.P., Horst, R.L., Beitz, D.E., and Littledike, E.T. (1988). J. Dairy Sci. 71,
1211.
Goff, J.P., Horst, R.L., Jardon, P.W., Borelli, C., and Wedam, J. (1996). J. Dairy
Sci. 79, 378.
Goff, J.P., Horst, R.L., Mueller, F.J., Miller, J.K, Kiess, G.A., and Dowlen, H.H.
148

V I TA M I N D(1991a). J. Dairy Sci. 74, 3863.
Goff, J.P., Reinhardt, T.A., and Horst, R.L. (1989). Endocrinology 125, 49.
Goff, J.P., Reinhardt, T.A., and Horst, R.L. (1991b). J. Dairy Sci. 74, 4022.
Goldblatt, H., and Soames, K.M. (1923). BIochem J. 17, 446.
Gonnerman, W.A., Toverud, S.V., Ramp, W.K., and Mechanic, G.L. (1976).
Proc. Soc. Exp. Biol. Med. 151, 453.
Green, H.B., Horst, R.L., Beitz, D.C., and Littledike, E.T. (1981). J. Dairy Sci.
64, 217.
Guilland, J.C., de Carvalho, M.J.C., Moreau, D., Boggio, V., Lhuissier, M., andFuchs, F. (1994). Nutr. Res. 14, 1317.
Haddad, J.G., and Walgate, J. (1976). J.Biol. Chem. 251, 4803.
Hannah, S.S., and Norman, A.W. (1994). Nutr. Rev. 52, 376.
Harrington, D.D., and Page, E.H. (1983). J. Am. Vet. Med. Assoc. 182, 1358.
Harrison, H.C., and Harrison, H.E. (1963). Am. J. Physiol. 205, 107.
Harrison, H.E. (1978). In Handbook Series in Nutrition and Food, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 3, p. 117. CRC Press, Boca
Raton, Florida.
Hay, A.W.M., and Watson, G. (1976). Comp. Biochem. Physiol. 56, 167.
Hazewinkel, H.A.W., How, K.L., Bosch, R., Goedegebuure, S.A., and Voorhout,G. (1987). In Ernahrung, Fehlernäbrung und Diätetik Bei Hund und Katze(H. Meyer and E. KIenzle, eds.) p. 125. Hanover, Germany.
Henry, H.L., and Norman, A.W. (1978). Science 201, 853.
Heuser, G.F., and Norris, L.C. (1929). Poult. Sci. 8, 89.
Hibbs, J.W., and Pounden, W.D. (1955). J. Dairy Sci. 38, 65.
Hidiroglou, M., Prouix, J.G., and Roubos, D. (1979). J. Dairy Sci. 62, 1076.
Hidiroglou, M., Williams, C.J., and Ho, S.K. (1978). Can. J. Anim. Sci. 58, 621.
Hirsch, A. (1982). Vitamins—The Life Essentials, p. 1. National Feed
Ingredients Association, Des Moines, Iowa.
Hodnett, D.W., Jorgensen, N.A., and DeLuca, H.F. (1992). J. Dairy Sci. 75, 485.
Holick, M.F. (1986). Curr. Top. Nutr. Dis. 5, 241.
Holick, M.F. (1987). Fed. Proc. Fed. Am. Soc. Exp. Biol, 46, 1876.
Holick, M.F. (1994). Am. J. Clin. Nutr. 60, 619.
Holick, M.F. (1995). Am. J. Clin. Nutr. 61, 638.
Holick, M.F., MacLaughlin, J.A., and Doppelt, S.H. (1981). Science 211, 590.
Horst, R.L., and Napoli, J.L. (1981). Fed. Prod. Fed. Am. Soc. Exp. Biol. 40,
898(Abstr.).
Horst, R.L., DeLuca, H.F., and Jorgensen, N.A. (1978). Metab. Bone Dis. Relat.
Res. 1, 29.
Horst, R.L., Goff, J.P., Reinhardt, T.A., and Buxton, D.R. (1997). J. Dairy Sci.
80, 1269.
How, K.L., Hazewinkel, H.A.W., and Mol, J.A. (1994a). Endocrinology 96, 12.
How, K.L., Hazewinkel, H.A.W., and Mol, J.A. (1994b). Vet. Rec. 134, 384.
How, K.L., Hazewinkel, H.A.W., and Mol, J.A. (1995). Proc. Voorjaarsdagen
Congress (H.P. Meyer and H.A.W. Hazewinkel, eds.) p. 1, Amsterdam.
Howard, G., Nguyen, T., Morrison, N., Watanabe, T., Sambrook, P, Eisman, J.,
and Kelly, P.J. (1995). J. Clin. Endocrinol. Metab. 80, 2800.
Hughes, M.R., Baylink, D.J., Jones, P.G., and Haussler, M.R. (1976). J. Clin.
Invest. 58, 61.
149

CHAPTER THREE

Hustmeyer, F.G., Beitz, D.C., Goff, J.P., Nonnecke, B.J., Horst, R.L., andReinhardt, T.A. (1994). J. Dairy Sci. 77, 3324.
James, S.Y., Mackay, A.G., Binderup, L., and Colston, K.W. (1994). J. Endocrin.
14, 555.
Johnson, D.W., and Palmer, L.S. (1939). J. Agr. Res. 58, 929.
Johnson, K.A., Church, D.B., Barton, R.J., and Wood, A.K.W. (1988). J. Small
Anim. Pract. 29, 657.
Jung, S.J., Lee, Y.Y., Pakkala, S., DeVos, S., Elstner, E., Norman, A.W.,
Uskokovic, J.G.M., and Koeffler, H.P. (1994). Leukemia Res. 18, 453.
Keiver, K.M., Draper, H.H., and Ronald, K. (1988). J. Nutr. 118, 332.
Kliewer, S.A., Umesono, K., Mangelsdorf, D.J., and Evans, R.M. (1992).
Nature, 355, 446.
Kragballe, K., Gjertsen, B.T., and DeHoop, D. (1991). Lancet 337, 193.
Kramer, B., and Gribetz, D. (1971). In The Vitamins (W.H. Sebrell Jr. and R.S.
Harris, eds.), p. 259. Academic Press, New York.
Lebrun, J.B., Moffatt, M.E.K., Mundy, R.J.T., Sangster, R.K., Postl, B.D.,
Dooley, J.P., Dilling, L.A., Godel, J.C., and Haworth, J.C. (1993). Can. J.
Public Health 84, 394.
Lefkowitz, E.S. and Garland, C.F. (1994). Int. J. Epidemiol. 23, 1133.
Lemire, J.M. (1992). Cell Biochem. 49, 26.
Lester, G.E. (1986). Fed. Proc. Fed. Am. Soc. Exp. Biol. 45, 2524.
Littledike, E.T., and Horst, R.L. (1982). J. Dairy Sci. 65, 749.
Lomba, F.G., Chauvaux, G., Teller, E., Lengele, L., and Beinfet, V. (1978). Br. J.
Nutr. 39, 425.
Loosli, J.K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.)Academic Press, San Diego.
Lowik, M.R.H., Van Den Berg, H., Schrijver J., Odink, J., Wedel, M., and Van
Houten, P. (1992). J. Am. Coll. Nutr. 11, 673.
Mac-Auliff, T., and McGinnis, J. (1976). Poult. Sci. 55, 2059(Abstr.).
Machlin, L.J., and Sauberlich, H.E. (1994). Nutr. Today Jan/Feb., 25.
Marks, R., Foley, P.A., Jolley, D., Knight, K.R., Harrison, J., and Thompson,
S.C. (1995). Arch. Dermatol. 13, 415.
Matsuoka, L.Y., Wortsman, J., Hanifan, N., and Holick, M.F. (1988). Arch.
Dermatol. 124, 1802.
Mattila, P.H., Piironen, V.I., Uusi-Rauva, E.J., and Koivistoinen, P.E. (1995). J.
Agri. Food Chem. 43, 2394.
Mayer, G.P., Ramberg, C.F., and Kronfeld, D.S. (1968). J. Nutr. 95, 202.
Maynard, L.A., Loosli, J.K., Hintz, H.F., and Warner, R.G. (1979). Animal
Nutrition, 7th Ed., p. 283. McGraw-Hill, New York.
McDowell, L.R. (1997). Minerals for Grazing Ruminants in Tropical Regions.
University of Florida, Gainesville.
McIntosh, G.H., and Tomas, F.M. (1978). Q. J. Exp. Physiol. 63, 119.
Mello, J.R., and Habermehl, G. (1995). Pesquisa Vet. Brasileria 15, 73.
Miller, E.R., Ullrey, D.E., Zutaut, C.L., Hoeffer, J.A., and Luecke, R.W. (1965).
J. Nutr. 85, 255.
Morand-Fehr, P. (1981). In Goat Production (G. Gall, ed.), p. 193. Academic
Press, New York.

150

V I TA M I N DMorgan, A.F., Axelrod, H.E., and Groody, M. (1947). Am. J. Physiol. 149, 333.
Morris, K.M.L. (1982). Vet. Hum. Toxicol. 24, 34.
Narbaitz, R., and Fragiskos, B. (1984). Calcif. Tissue Res. 36, 392.
Neibergs, H.L., Bosworth, B.T., and Reinhardt, T.A. (1996). J. Dairy Sci. 79,
1313.
Nemere, I. (1995). J. Nutr. 125, 1695S.
Nemere, I., Zhou, I.X., and Norman, A.W. (1993). Receptor 3, 277.
Norman, A.W. (1995). J. Nutr. 125, 1687S.
Norman, A.W., and DeLuca, H.F. (1963). Biochemistry 2, 1160.
NRC. Nutrient Requirements of Domestic Animals. National Academy of
Sciences-National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of SciencesNational Research Council, Washington, D.C.
O’Brien, K.O. (1998). Nutr. Rev. 56, 148
O’Connell, J.P., and Gatlin, D.M. (1994). Aquaculture 125, 107.
Oetzel, G.R., Olson, J.D., Curtis, C.R., and Fettman, M.J. (1988). J. Dairy Sci.
71, 3302.
Ohnuma, N., Bannai, K., Yamagauchi, H., Hashimoto, Y., and Norman, A.W.
(1980). Arch. Biochem. Biophys. 204, 387.
Okada, K.A., Carrillo, B.J., and Tilley, M. (1977). Econ. Bot. 31, 225.
Oldham, S.B., Mitnick, S.A., and Coburn, J.W. (1980). J. Biol. Chem. 255.
5789.
Pakkala, S., DeVos, S., Elstner, E., Rude, R.K., Uskokovic, M., Binderup, L., andKoeffler, H.P. (1995). Leukemia Res. 18, 65.
Payne, J.M., and Manston, R. (1967). Vet. Rec. 81. 214.
Pointillart, A., Fontaine, N., and Thomasset, M. (1986). Brit. J. Nutr. 56, 661.
Quaterman, J., Dalgarno, A.C., Adam, A., Fell, B.F., and Boyne, R. (1964). Br.
J. Nutr. 18, 65.
RDA. (1989). Recommended Dietary Allowances, 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
Richter, V.G.H., Flachowsky, F., Matthey, M., Ochrimenko, W.I., Wolfram, D.,
and Schade, T. (1990). Vet. Med. 45, 227.

151

CHAPTER THREE

Rowland, R. (1982). Vitamins—The Life Essentials, p. 1. National Feed
Ingredients Association, Des Moines, Iowa.
Rupel, I.W., Bohstedt, G., and Hart, E.B. (1933). Wisc. Agric. Exp. Stn. Res.
Bull. 115.
Sachs, M., Bar, A., Cohen, R., Mazur, Y., Mayer, E., and Horwitz, S. (1977).
Am. J. Vet. Res. 38, 2039.
Schachter, D., and Kowarski, S. (1982). Fed. Proc. Fed. Am. Soc. Exp. Biol. 41,
84.
Schneider, J. (1986). In Proc. of the NFIA Nutrition Institute, p. 1, Ames, Iowa.
Scott, D., and McLean, A.F. (1981). Proc. Nutr. Soc. 40, 257.
Scott, K.C., and Latshaw, J.D. (1994). Anim. Feed Sci. Tech. 47. 99.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Sergeev, I.N., Arkhapchev, Y.P., and Spirichev, V.B. (1990). J. Nutr. 120, 1185.
Shiau, S., and Hwang, J. (1994). J. Nutr. 124, 2445.
Skowronski, R.J., Peehl, D.M., and Feldman, D. (1995). Endocrinology 136, 20.
Solanki, T., Hyatt, R.H., Kemm, J.R., Hughes, E.A., and Cowan, R.A. (1995).
Age and Aging 24, 103.
Sommerfeldt, J.L., Horst, R.L., Littledike, E.T., Beitz, D.C., and Napoli, J.L.
(1981). J. Dairy Sci. 64, 157.
Sommerfeldt, J.L, Napoli, J.L., Littledike, E.T., Beitz, D.C., and Horst, R.L.
(1983). J. Nutr. 114, 2595.
Sood, S., Reghunandanan, R., and Reghunandanan, V. (1995). Indian J. Exp.
Biol. 33, 61.
Steenbock, H., and Black, A. (1924). J. Biol. Chem. 61, 405.
Stevens, V.I., and Blair, R. (1985). Poult. Sci. 64, 510.
Studzinski, G.P., and Moore, D.C. (1995). Cancer Res. 55, 4014.
Suter, P. (1957). Schweiz. Arch. Tierheilkd. 99, 421.
Sutton, R.A.L., and Dirks, J.H. (1978). Fed. Proc. Fed. Am. Soc. Exp. Biol. 37,
2112.
Takeuchi, A., Okano, T., Tanda, M., and Kobayashi, T. (1991). Comp Biochem.
Physiol. 100A, 483.
Tian, X.Q., Chen, T.C., Liu, Z., Shao, Q., and Holick, M.F. (1994).
Endocrinology 135, 655.
Tindall, B. (1985). Anim. Health Nutr. 40, 26.
Tomas, F.M. (1974). Aust. J. Agric. 25, 495.
Tomas, F.M., and Somers, M. (1974). Aust. J. Agric. Res. 25, 475.
Tsang, C.P.W. (1992) Poult. Sci. 71, 215.
Tuan, R.S., and Suyama, E. (1996). J. Nutr. 126, 1309S.
Tucker, G., Gagnon, R.E., and Hassler, M.R. (1973). Arch. Biochem. Biophys.
155, 47.
Vaiano, S.A., Azuolas, J.K., and Parkinson, G.B. (1994). Poult. Sci. 73, 1296.
Vink van Wijngaarden, T., Birkenhager, J.C., Kleinekoort, W.M., Van Den Bend,
G.J., Pols, H.A., and Van Leeuwen, J.P. (1995). Endocrinology 136, 812.
Wallis, G.C. (1944). S.D. Agric. Exp. Stn. Bull. No. 372. South Dakota State
University, Brookings.
Wasserman, R.H. (1975). Nutr. Rev. 33, 1.

152

V I TA M I N D

Wasserman, R.H. (1981). Fed. Proc Fed. Am. Soc. Exp. Biol. 40, 68.
Webb, A.R., Kline, L., and Holick, M.F. (1988). J. Clin. Endocrinol. Metab. 67,
373.
Weber, J.C., Pons, V., and Kodicek, E. (1971). Biochem J.. 125, 147.
Whitfield, G.K., Hsieh, J., Jurutha, P.W., Selznick, S.H., Haussler, C.A.,
MacDonald, P.N., and Haussler, M.R. (1995). J. Nutr. 125, 1690S.
Wolker, N.A., and Carrillo, B.J. (1967). Nature (London) 215, 72.

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chapter fourV I TA M I N E

INTRODUCTION
Vitamin E is recognized as an essential nutrient for all species of animals, including humans. However, opinions differ among research
workers as well as practical livestock producers regarding conditions
under which vitamin E supplementation is required and at what levels it
should be fed. For years, vitamin E in human nutrition was described as“a vitamin looking for a disease.” Some vitamin E-deficiency conditions
that occurred in animals were not seen in humans; however, a numberof medical claims for physiological benefits from the vitamin have beenmade. In more recent years, vitamin E has been shown to be important
against free-radical injury; enhancing the immune response; and playing
a role in prevention of cancer, heart disease, cataracts, Parkinson’s disease, and a number of other disease conditions.

HISTORY
Excellent reviews of the history of vitamin E have been provided by
Scott (1980) and Ullrey (1982). By the early 1920s, existence of vitamins
A, B (thiamin), and C was established, and that of vitamin D was virtually assured.
As a result of the stimulus to experiment with purified diets that followed the discovery of the first vitamins, it was frequently observed thaton certain diets, which were satisfactory for growth and health, rats
failed to reproduce. Vitamin E was discovered in 1922 by Evans andBishop (University of California, Berkeley) as an unidentified factor invegetable oils required for reproduction in female rats. In their experiments, estrus, mating, and all detectable phases of the beginning of preg155

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nancy were normal, but fetuses soon died and were resorbed unless thediet was supplemented with small amounts of wheat germ, dried alfalfa
leaves, or fresh lettuce.
At first this substance was known as factor X, but Sure (1924) andEvans (1925) soon proposed the name vitamin E, since this was the next
serial alphabetical designation. Vitamin E was isolated as α-tocopherol.
The name tocopherol is derived from the Greek tokos meaning childbirth or offspring, the Greek pherein meaning to bring forth, and ol todesignate an alcohol. The structure of α-tocopherol was determined by
E. Fernholz in 1938, and the substance was first synthesized by P. Karrer,
also in 1938. δ-Tocopherol was reported in 1947, and tocotrienols were
described in about 1959.
It also became apparent that male rats were affected by a deficiencyof this nutrient that resulted in testicular degeneration. Throughout the
1920s, vitamin E was recognized only as a factor required for reproduction in rats. Many clinical studies were undertaken to determine theeffects of vitamin E on various reproductive problems in humans,though in most cases it was found to have little or no effect.
In 1931, Pappenheimer and Goettsch conducted a series of whatbecame classic experiments showing that vitamin E is also required forprevention of encephalomalacia in chicks and of nutritional muscular
dystrophy in rabbits and guinea pigs. By 1944, it was found that a multiplicity of clinical signs occur in animals suffering from vitamin E deficiency. In a single species, the chick, three distinct vitamin E deficiency
diseases were documented: exudative diathesis, encephalomalacia, andmuscular dystrophy. Also, embryonic failure in poultry was found to becaused by damage to the circulatory system.
The first controlled study of vitamin E deficiency in swine was thatof Adamstone et al. (1949). These workers reported a decline in reproductive efficiency and signs of locomotor incoordination and muscularnecrosis. Obel (1953) described a naturally occurring dietary disease invitamin E-deficient swine that was characterized by hepatic necrosis, fibrinoid degeneration of blood vessel walls, and muscular dystrophy. In
1957, Klaus Schwarz and associates, studying dietary liver necrosis ofrats receiving a diet low in vitamin E, showed that dried brewer’s yeast,
which contains no vitamin E, was as effective as vitamin E in preventing
liver necrosis. Shortly after this discovery, selenium (Se) was found to be
the active ingredient in brewer’s yeast and able to replace vitamin E forprevention of exudative diathesis in poultry, tissue degeneration inswine, and muscular degeneration in young ruminants. Much confusion
existed because earlier discoveries had shown that synthetic antioxidants156

V I TA M I N E

as well as sulfur amino acids were as effective as vitamin E for prevention of some vitamin E deficiency diseases. Considerable effort has been
made to clarify the mechanisms of vitamin E, Se, sulfur amino acids, andantioxidants in relation to diseases that are vitamin E responsive.
Likewise, considerable research in the last 10 years has confirmed thebeneficial aspects of vitamin E supplementation for both livestock andhumans.

CHEMICAL STRUCTURE AND PROPERTIES
Vitamin E activity in food derives from a series of compounds ofplant origin, the tocopherols and tocotrienols. Eight forms of vitamin E
are found in nature: four tocopherols (α, β, γ, and δ) and four
tocotrienols (α, β, γ, and δ). All have a 6-chromanol ring structure and aside chain. The structures of α-tocopherol and the commercially available α-tocopheryl acetate are presented in Fig. 4.1, while different active
forms of vitamin E are shown in Fig. 4.2. Differences among α, β, γ, andδ are due to the placement of methyl groups on the ring. The difference
between tocopherols and tocotrienols is due to unsaturation of the side
chain in the latter.
The dl-α-tocopheryl acetate (also called all-rac-α-tocopheryl
acetate) is accepted as the International Standard (1 mg = 1 international unit). Synthetic-free tocopherol, dl-α-tocopherol, has a potency of 1.1
IU/mg. Activity of naturally occurring α-tocopherol, d-α-tocopherol

Fig. 4.1 Structure of α-tocopherol and α-tocopheryl acetate.
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Fig. 4.2 Structural differences among vitamin E forms.

(also called RRR-tocopherol, see below), is 1.49 IU/mg, and of its
acetate, 1.36 IU/mg.
The dl-α-tocopheryl acetate is made by the extraction of natural
tocopherols from vegetable oil. Extracted tocopherols undergo distillation to obtain the alpha form, and are then acetylated to produce theacetate ester. α-Tocopherol, the most active compound, is fully methylated, with methyl groups at positions 5, 7, and 8 (2 R, 4´R, 8´R-α-tocopherol, abbreviated RRR). The loss of one or both of the methyl groupsat position 5 or 7 on the ring sharply reduces vitamin E activity of thestructures. The position of the methyl groups influences the vitamin E
activity of the molecule.
dl-α-Tocopheryl acetate is the most widely available source of vitamin E for supplementation. The acetate ester is very stable to in vitro
oxidation and has no activity as an in vitro antioxidant. However, it isreadily hydrolyzed in the intestine to nonesterified or free tocopherol,
which is the potent in vivo antioxidant.
A number of vitamin E products are available that are highly effective in preventing or curing conditions associated with inadequate
dietary vitamin E. Ochoa et al. (1992) compared six vitamin E sourcesthat differed in relation to carrier used, emulsification, liquid or dry
158

V I TA M I N E

product, and a micellized form. All tissues tested resulted in relatively
similar tissue vitamin E concentrations. The naturally derived (d) andsynthetic (dl) chemical forms of vitamin E are not used equally andbecause of this have different biopotencies. Technically, the d formshould not be referred to as “natural” as it is derived from natural
sources but then undergoes chemical processing (e.g., methylation andhydrogenation). Free d- and dl-α-tocopherol as alcohol forms and theirrespective ester forms have the highest biopotency. Since the free form iseasily oxidized, more stable forms such as acetate and succinate esters
have been synthesized with reduced biopotencies.
Serum and certain tissue vitamin E concentrations are influenced bymethod of supplementation, dosage levels, chemical formulation, andcarrier of vitamin E supplements (Charmley et al., 1992; Hidiroglou etal., 1990; Hidiroglou and Charmley, 1991; Njeru et al., 1992, 1994b).
Some studies with sheep and cattle have indicated that the d form resulted in higher serum and selected tissue α-tocopherol concentrations thanthe dl when administered on an equal IU basis (Hidiroglou andMcDowell, 1987; Hidiroglou et al., 1988a,b). On the contrary, althoughhigher serum α-tocopherol was found in preruminant calves (Roquet etal., 1992) and sheep (Hidiroglou et al., 1992a), and in selected tissues
for sheep (Hidiroglou et al., 1994) for d versus dl forms of vitamin E,
the relative IU biopotencies were similar. The ratios of d to dl-α-tocopherol were in close agreement with the established U.S. Pharmacopeia
biopotency of 1.36 for d- to dl-α-tocopheryl acetate. It is not just natural versus synthetic that is important for vitamin E biopotency but also
the ester and carrier used. In sheep, Hidiroglou and Singh (1991) reported that with equivalent IU dosage, the natural form of d-α-tocopheryl
succinate had only one-third the biopotency of the synthetic dl-α-tocopheryl acetate, indicating that the ester succinate has less value than theacetate. Jensen et al. (1999) also found the acetate to be a better form ofvitamin E than succinate.
Although less stable, alcohol forms of vitamin E resulted in higher
concentrations of α-tocopherol in serum and tissues compared to ester
forms (e.g., acetate) (Hidiroglou et al., 1988a, 1989; Ochoa et al.,1992). From a practical viewpoint, this may be important as there is asuggestion that a young ruminant may not utilize ester forms (e.g.,
acetate) as well as the alcohol forms of vitamin E for the first few weeks
of life.
More research is needed to evaluate the different forms of vitamin E
and the carriers of these products. Until more research is conducted,
opinion will remain divided on biopotencies of various vitamin E forms,
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CHAPTER FOUR

particularly d versus dl. Studies are needed to compare forms withdietary levels closer to ruminant vitamin E requirements since, for anumber of nutrients, homeostasis mechanisms discriminate less between
chemical forms when administered at the physiologically required concentration.
α-Tocopherol is a yellow oil that is insoluble in water but soluble inorganic solvents. Tocopherols are extremely resistant to heat but arereadily oxidized. Natural vitamin E is subject to destruction by oxidation, which is accelerated by heat, moisture, rancid fat, light, alkali, andcertain trace minerals (e.g., copper and iron). α-Tocopherol is an excellent natural antioxidant that protects carotene and other oxidizable
materials in feed and in the body. However, in the process of acting as
an antioxidant, it is destroyed. Since esterification of the vitamin
improves its stability, commercial supplements usually contain d-α-tocopheryl acetate or dl-α-tocopheryl acetate.

ANALYTICAL PROCEDURES
Many methods for the determination of tocopherols in feedstuffs
and animal tissues have been introduced, most of them based on separation of the tocopherols by column, paper, or thin-layer chromatography, followed by a colorimetric reaction. Separation steps, however, areusually laborious and time consuming, and colorimetric reactions suchas that of Emmerie and Engel are often subject to interference fromother compounds (McMurray and Blanchflower, 1979). High-pressureliquid chromatography (HPLC) offers the possibility of combining rapid
analysis with separation of tocopherols from interfering substances. The
method consists of three main steps via extraction, saponification, andchromatography (McMurray and Blanchflower, 1979; Cohen andLapointe, 1980). Either reverse-phase or normal-phase procedures areused, coupled with either fluorometric or ultraviolet detectors. The procedures are specific, rapid, simple, and very sensitive. With fluorometricdetection, nanogram quantities can be detected; HPLC procedures permit separation and quantitation of the α, β, and γ isomers of tocopherol.
Biological assay methods for vitamin E determine the ability of an
unknown to prevent or reverse specific vitamin E deficiency signs of animals in vivo (e.g., fetal resorption, encephalomalacia, muscular degeneration, erythrocyte hemolysis) or indications of vitamin E status (e.g.,liver and serum concentrations). The biological assay developed by
Herbert Evans was the rat fetal resorption test. International units havebeen established on the basis of the resorption test. This is the ultimate
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V I TA M I N E

measure of vitamin E activity and is based on development of live fetuses in rats. Litter efficiency is calculated as the number of females withliving young divided by total number of pregnancies.
Biological variations are inherent in bioassays, considerably more sothan in chemical assays as a rule. They involve variations in response
between individuals, families, strains, and species of animals, as well as
management skill, stability of the test material, diet standardization andpurity, and control of other environmental factors. Dietary history (particularly previous vitamin E intakes) of parent stock can have an influence. Table 4.1 illustrates a comparison among vitamin E forms related
to biological assays.

METABOLISM
Absorption and Transport
Vitamin E absorption is related to fat digestion and is facilitated bybile and pancreatic lipase (Sitrin et al., 1987). The primary site ofabsorption appears to be the medial small intestine. Whether presented
as free alcohol or as esters, most vitamin E is absorbed as the alcohol.
Esters are largely hydrolyzed in the gut wall, and the free alcohol enters
the intestinal lacteals and is transported via lymph to the general circulation. Medium-chain triglycerides particularly enhance absorption,
whereas polyunsaturated fatty acids (PUFAs) are inhibitory. Balance■ Table 4.1

Relative Biopotency of Vitamin E Forms

Rat
Antisterility

RabbitRat
Cure of Muscular
Weight Gain
Dystrophy

Hemolysis of
Erythrocytes
in vivoin vitro

Tocopherols
d-α
dl-α
d-β
dl-β
d-γ
dl-γ
d-δ

135
100
54
27
1
1–11
1

—
100
—
25
—
19
—

100
90
30
—
20
6
—

130
100
30
25
4–22
18
3

100
100
40
54
30
67
20

Tocotrienols
d-β
d-α
d-γ

5
29
3

—
—
—

—
—
—

1–5
23
—

133
106
88

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CHAPTER FOUR

studies indicate that much less vitamin E is absorbed, or at least
retained, in the body than vitamin A. Vitamin E recovered in feces froma test dose was found to range from 65 to 80% in the human, rabbit,
and hen, although in chicks, it was reported at about 25%. It is notknown how much fecal vitamin E represents unabsorbed tocopherol andhow much may come via secretion in the bile. As the intake increases,
the percentage of tocopherol absorbed decreases, suggesting a saturation
process.
The tocopherol form, which is the naturally occurring one, is subject to destruction in the digestive tract to some extent, whereas theacetate ester is not. Much of the acetate is readily split off in the intestinal wall, and the alcohol is reformed and absorbed, thereby permitting
the vitamin to function as a biological antioxidant. Any acetate form
absorbed or injected into the body evidently is converted there to thealcohol form.
Vitamin E in plasma is attached mainly to lipoproteins in the globulin fraction within cells and occurs mainly in mitochondria and microsomes. The vitamin is taken up by the liver and is released in combination with low-density lipoprotein (LDL) cholesterol. Rates and amounts
of absorption of the various tocopherols and tocotrienols are in the samegeneral order of magnitude as their biological potencies. α-Tocopherol
is absorbed best, with γ-tocopherol absorption 85% that of α-forms butwith a more rapid excretion. One can generally assume that most of thevitamin E activity within plasma and other animal tissues is α-tocopherol (Ullrey, 1981). In humans, whose natural diet contains a highpercentage of non-alpha forms, blood serum tocopherols identified consisted of about 87% α-, 11% γ-, and 2% β-tocopherol (Hoffmann-La
Roche, 1972).

Placental and Mammary Transfer
Vitamin E does not cross the placenta in any appreciable amounts;
however, it is concentrated in colostrum (Van Saun et al., 1989). With
respect to neonatal ruminants (Hidiroglou et al., 1969; Van Saun et al.,1989) and baby pigs (Mahan, 1991), several investigators have reported limited placental transport of α-tocopherol, making neonates highly
susceptible to vitamin E deficiency. This may be related to a decrease inefficiency of placental vitamin E transfer as gestation proceeds, a dilution effect as a result of rapid fetal growth, or a decrease in available
maternal vitamin E. With limited placental transfer of vitamin E, newborns must rely heavily on ingestion of colostrum as a source of vitamin
E. Van Saun et al. (1989) reported decreased fetal serum vitamin E con162

V I TA M I N E

centrations with increasing fetal age. Additionally, these authors reported less decline in fetal serum vitamin E concentration during gestation infetuses from vitamin E adequate dams.
There is inefficient placental transfer of vitamin E, but high levels of
the vitamin have been shown in calves (Nockels, 1991) and lambs(Njeru et al., 1994a) after consumption of colostrum. Nockels (1991)
reported α-tocopherol levels in plasma from beef calves prior tocolostrum consumption and for several days thereafter. Precolostral
plasma vitamin E levels averaged 0.2 μg/ml and increased to 3.3 μg/ml
at 5 to 8 days of age. Njeru et al. (1994a) fed ewes dl-α-tocopheryl
acetate at graded levels (0, 15, 30, and 60 IU per head daily) to study
placental and mammary gland transfer. Supplemental vitamin E had noeffect on serum α-tocopherol of lambs prior to nursing, averaging 0.35μg/ml. By day 3, lamb serum tocopherol increased to 1.41, 1.84, 2.43,
and 4.46 μg/ ml, respectively, for the four supplemental dietary levels ofvitamin E (Table 4.2). Vitamin E at the given levels of supplementation
increased colostral α-tocopherol at a linear rate of 3.3, 6.8, 8.0, and 9.6
μg/ml, respectively. The importance of providing colostrum rich in vitamin E is quite apparent, as both calves and lambs are born with low levels of the vitamin (Nockels, 1991; Njeru et al., 1994a). Low blood vitamin E may lead to diminished disease resistance and immune responsein the neonate (Nockels, 1991).

Storage and Excretion
Vitamin E is stored throughout all body tissues; major deposits arein adipose tissue, liver, and muscle, with highest storage in the liver.
However, liver contains only a small fraction of total body stores, in■ Table 4.2

Effect of Supplemental Vitamin E to Prepartum Ewes on α-Tocopherol
Concentration in Serum and Colostrum (μg/ml)
Ewes

Lambs

Supplemental Vitamin E(IU per day)

Serum atParturition

Colostrum
Day 1

Serum Prior
to Nursing

Serum
Day 3

0
15
30
60

0.94
1.94
2.53
4.07

3.3
6.8
8.0
9.6

0.40
0.40
0.38
0.23

1.41
1.84
2.43
4.46

Source: Njeru et al. (1994b).
Note: Treatments administered as dl-α-tocopheryl acetate 28 days prepartum through28 days postpartum.
Linear (P < 0.05) treatment effects for α-tocopherol in ewe serum and colostrum
and lamb serum at day 3.

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CHAPTER FOUR

contrast to vitamin A, for which about 95% of the body reserves are in
the liver. The extent of storage is shown by the fact that females born of
mothers whose diets contained a liberal supply frequently have enoughin their bodies at birth to carry them through a first pregnancy. Rats
reared on natural foods rich in the vitamin and then placed on a deficient diet may produce three or four litters before exhausting theirreserves (Maynard et al., 1979). However, Gallo-Torres (1980) reportsthat unlike vitamin A, lower body stores of vitamin E are available for
periods of low dietary intake.
Tocopherol entering the circulatory system becomes distributed
throughout the body, with most of it localizing in the fatty tissues.
Subcellular fractions from different tissues vary considerably in their
tocopherol content; the highest levels are found in membranousorganelles, such as microsomes and mitochondria, which contain highly
active redox systems (McCay et al., 1981; Taylor et al., 1976).
Small amounts of vitamin E will persist tenaciously in the body fora long time. However, stores are exhausted rapidly by PUFAs in the tissues; the rate of disappearance is proportional to the intake of PUFAs.
A major excretion route of absorbed vitamin E is bile, in which tocopherol appears mostly in the free form. Usually less than 1% of orally
ingested vitamin E is excreted in the urine.

FUNCTIONS
Vitamin E has been shown to be essential for integrity and optimumfunction of the reproductive, muscular, circulatory, nervous, andimmune systems (Hoekstra, 1975; Sheffy and Schultz, 1979; Bendich,
1987; McDowell et al., 1996). It is well established that some functionsof vitamin E, however, can be fulfilled in part or entirely by traces of Se
or by certain synthetic antioxidants. Even the sulfur-bearing aminoacids, cystine and methionine, affect certain vitamin E functions. Much
evidence points to undiscovered metabolic roles for vitamin E that may
be paralleled biologically by roles of Se and possible other substances.
The most widely accepted functions of vitamin E are discussed in thissection.

Vitamin E as a Biological Antioxidant
Vitamin E has a number of different but related functions. One of
the most important functions is its role as an intercellular and intracellular antioxidant. Vitamin E is part of the body’s intracellular defense
against the adverse effects of reactive oxygen and free radicals that ini164

V I TA M I N E

tiate oxidation of unsaturated phospholipids (Chow, 1979) and criticalsulfhydryl groups (Brownlee et al., 1977). Vitamin E functions as aquenching agent for free radical molecules with single, highly reactive
electrons in their outer shells. Free radicals attract a hydrogen atom,along with its electron, away from the chain structure of a PUFA, satisfying the electron needs of the original free radical but leaving the PUFA
short one electron. Thus, a fatty acid free radical is formed that joinswith molecular oxygen to form a peroxyl radical that steals a hydrogenelectron unit from yet another PUFA. This reaction can continue in achain, resulting in the destruction of thousands of PUFA molecules(Gardner, 1989; Herdt and Stowe, 1991). Free radicals can be extremely damaging to biological systems (Padh, 1991). Free radicals, includinghydroxy, hypochlorite, peroxy, alkoxy, superoxide, hydrogen peroxide,and singlet oxygen, are generated by autoxidation or radiation, or from
activities of some oxidases, dehydrogenases, and peroxidases.
Highly reactive oxygen species, such as superoxide anion radical(O2), hydroxyl radical (HO), hydrogen peroxide (H2O2), and singlet
oxygen (O2), are continuously produced in the course of normal aerobic
cellular metabolism. Also, phagocytic granulocytes undergo respiratory
burst to produce oxygen radicals to destroy the intracellular pathogens.
However, these oxidative products can, in turn, damage healthy cells ifthey are not eliminated. Antioxidants serve to stabilize these highly reactive free radicals, thereby maintaining the structural and functional
integrity of cells (Chew, 1995). Therefore, antioxidants are very important to the immune defense and health of humans and animals.
The antioxidant function of vitamin E is closely related to and synergistic with the role of Se. Selenium has been shown to act in aqueous
cell media (cytosol and mitochondrial matrix) by destroying hydrogen
peroxide and hydroperoxides via the enzyme glutathione peroxidase(GSHpx) of which it is a co-factor. In this capacity, it prevents oxidationof unsaturated lipid materials within cells, thus protecting fats within
the cell membrane from breaking down. It is the oxidation of vitamin E
that prevents oxidation of other lipid materials to free radicals and peroxides within cells, thus protecting the cell membrane from damage(Drouchner, 1976). If lipid hydroperoxides are allowed to form in theabsence of adequate tocopherols, direct cellular tissue damage canresult, in which peroxidation of lipids destroys structural integrity of thecell and causes metabolic derangements.
Vitamin E reacts or functions as a chain-breaking antioxidant,
thereby neutralizing free radicals and preventing oxidation of lipids
within membranes. Free radicals may not only damage their cell of ori165

CHAPTER FOUR

gin but migrate and damage adjacent cells in which more free radicals
are produced in a chain reaction leading to tissue destruction (Nockels,
1991). At least one important function of vitamin E is to interrupt production of free radicals at the initial stage. Myodystrophic tissue is common in cases of vitamin E-Se deficiency with leakage of cellular compounds such as creatinine and various transaminases through affected
membranes into plasma. The more active the cell (e.g., the cells of skeletal and involuntary muscles), the greater the inflow of lipids for energy
supply and the greater the risk of tissue damage if vitamin E is limiting.
This antioxidant property also ensures erythrocyte stability and maintenance of capillary blood vessel integrity.
Interruption of fat peroxidation by tocopherol explains the wellestablished observation that dietary tocopherols protect or spare body
supplies of such oxidizable materials as vitamin A, vitamin C, and the
carotenes. Certain deficiency signs of vitamin E (e.g., muscular dystrophy) can be prevented by diet supplementation with other antioxidants,
thus lending support to the antioxidant role of tocopherols. Semen quality of boars was improved with Se and vitamin E supplementation, withvitamin E playing a role in maintaining sperm integrity in combination
with Se (Marin-Guzman et al., 1989). Chemical antioxidants are stored
only at very low levels, thus they are not as effective as tocopherol. It isclear that highly unsaturated fatty acids in the diet increase vitamin E
requirements. When acting as an antioxidant, vitamin E supplies
become depleted, thus furnishing an explanation for the often observed
fact that the presence of dietary unsaturated fats (susceptible to peroxidation) augments or precipitates vitamin E deficiency.

Membrane Structure and Prostaglandin Synthesis
α-Tocopherol may be involved in the formation of structural components of biological membranes, thus exerting a unique influence onarchitecture of membrane phospholipids (Ullrey, 1981). It is reported
that α-tocopherol stimulated the incorporation of 14C from linoleic acidinto arachidonic acid in fibroblast phospholipids. Also, it was foundthat α-tocopherol exerted a pronounced stimulatory influence on formation of prostaglandin E from arachidonic acid, while a chemical
antioxidant had no effect.

Blood Clotting
Vitamin E is an inhibitor of platelet aggregation in pigs (McIntosh
et al., 1985), and may play a role by inhibiting peroxidation of arachidonic acid, which is required for formation of prostaglandins involved
166

V I TA M I N E

in platelet aggregation (Panganamala and Cornwell, 1982; Machlin,1991).

Disease ResistanceIn addition to the relationship of vitamin E and Se, vitamin E, vitamin C, and β-carotene as antioxidant vitamins together have importanttissue defense mechanisms against free-radical damage. The dietary andtissue balance of all these nutrients are important in protecting tissues
against free-radical damage. Both in vitro and in vivo studies showedthat the antioxidant vitamins generally enhance different aspects of cellular and noncellular (humoral) immunity. The antioxidant function ofthese vitamins could, at least in part, enhance immunity by maintaining
the functional and structural integrity of important immune cells. A
compromised immune system will affect animal health and result in
reduced animal production efficiency through increased susceptibility todiseases, thereby leading to increased animal morbidity and mortality.
One function of vitamin C is that this vitamin can regenerate thereduced form of α-tocopherol, perhaps accounting for observed sparing
effects of these vitamins (Jacob, 1995; Tanaka et al., 1997). In theprocess of sparing fatty acid oxidation, tocopherol is oxidized to the
tocopheryl free radical. Ascorbic acid can donate an electron to the
tocopheryl free radical, regenerating the reduced antioxidant form of
tocopherol.
Considerable attention is being directed to the role vitamin E and Se
play in protecting leukocytes and macrophages during phagocytosis, themechanism whereby animals immunologically kill invading bacteria.
Both vitamin E and Se may help these cells to survive the toxic productsthat are produced in order to effectively kill ingested bacteria (Badwey
and Karnovsky, 1980). Macrophages and neutrophils from vitamin Edeficient animals have decreased phagocytic activity (Burkholder andSwecker, 1990).
Since vitamin E acts as a tissue antioxidant and aids in quenching
free-radicals produced in the body, any infection or other stress factors
may exacerbate depletion of the limited vitamin E stores from various
tissues. With respect to immunocompetency, dietary requirements may
be adequate for normal growth and production; however, higher levels
have been shown to positively influence both cellular and humoral
immune status of ruminant species. The former two responses are generally used as criteria for determining the requirement of a nutrient.
During stress and disease, there is an increase in production of glucocorticoids, epinephrine, eicosanoids, and phagocytic activity. Eicosanoid
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CHAPTER FOUR

and corticoid synthesis and phagocytic respiratory bursts are prominent
producers of free radicals that challenge the animal’s antioxidant systems. Vitamin E has been implicated in stimulation of serum antibody
synthesis, particularly IgG antibodies (Tengerdy, 1980). The protective
effects of vitamin E on animal health may be involved with its role in
reduction of glucocorticoids, which are known to be immunosuppressive (Golub and Gershwin, 1985). In rats, an in vivo inflammatory challenge decreased vitamin E blood and liver concentrations (Fritsche andMcGuire, 1996). Vitamin E also most likely has an immuno-enhancing
effect by virtue of altering arachidonic acid metabolism and subsequent
synthesis of prostaglandin, thromboxanes, and leukotrienes. Understress conditions, increased levels of these compounds by endogenous
synthesis or exogenous entry may adversely affect immune cell function(Hadden, 1987).
The effects of vitamin E and Se supplementation on protection
against infection by several types of pathogenic organisms, as well as
antibody titers and phagocytosis of the pathogens, have been reported
for calves (Cipriano et al., 1982; Reddy et al., 1987a; Rajaraman et al.,1998) and lambs (Reffett et al., 1988; Finch and Turner, 1989; Turnerand Finch, 1990). As an example, calves receiving 125 IU of vitamin E
daily were able to maximize their immune responses compared withcalves receiving low dietary vitamin E (Reddy et al., 1987b). In sows,
vitamin E restriction depressed lymphocytes and polymorphonuclear
cells for immune function (Wuryastuti et al., 1993). Dogs with vitamin
E deficiency had a depressed proliferative lymphocyte responsiveness(Langweiler et al., 1983).
Antioxidants, including vitamin E, play a role in resistance to viral
infection. Vitamin E deficiency allows a normally benign virus to cause
disease (Beck et al., 1994). In mice, enhanced virulence of a virus resulted in myocardial injury that was prevented with vitamin E adequacy.
Selenium or vitamin E deficiency leads to a change in viral phenotype,such that an avirulent strain of a virus becomes virulent and a virulent
strain becomes more virulent (Beck, 1997).

Electron Transport and Deoxyribonucleic Acid (DNA)There is limited evidence that vitamin E is involved in biologicaloxidation-reduction reactions (Hoffmann-La Roche, 1972). Vitamin E
also appears to regulate the biosynthesis of DNA within cells.

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V I TA M I N E

Relationship to Toxic Elements or Substances
Both vitamin E and Se provide protection against toxicity with threeclasses of heavy metals (Whanger, 1981). In one class, which includes
metals like cadmium and mercury, Se is highly effective in altering toxicities, but vitamin E has little influence. In the second class, which
includes silver and arsenic, vitamin E is highly effective; Se is also effective but only at relatively high levels. The third class, which includes
lead, is counteracted by vitamin E, but Se has little effect.
Vitamin E can be effective against other toxic substances. For
example, treatment with vitamin E gave protection to weanling pigs
against monensin-induced skeletal muscle damage (Van Vleet et al.,1987).

Relationship with Selenium in Tissue Protection
There is a close working relationship between vitamin E and Se
within tissues. Selenium has a sparing effect on vitamin E and delays
onset of deficiency signs. Likewise, vitamin E and sulfur amino acids
partially protect against or delay onset of several forms of Se deficiency syndromes. Tissue breakdown occurs in most species receiving diets
deficient in both vitamin E and Se, mainly through peroxidation.
Peroxides and hydroperoxides are highly destructive to tissue integrity and lead to disease development. It appears that vitamin E in cellular and subcellular membranes is the first line of defense against peroxidation of vital phospholipids, but even with adequate vitamin E,
some peroxides are formed. Selenium, as part of the enzyme glutathione peroxidase, is a second line of defense that destroys these peroxides before they have an opportunity to cause damage to membranes. Therefore, Se, vitamin E, and sulfur-containing amino acids,
through different biochemical mechanisms, are capable of preventing
some of the same nutritional diseases. Vitamin E prevents fatty acidhydroperoxide formation, sulfur amino acids are precursors of glutathione peroxidase, and Se is a component of glutathione peroxidase(Smith et al., 1974).
To some extent, vitamin E and Se are mutually replaceable, but there
are lower limits below which substitution is ineffective. In diets severely
deficient in Se, vitamin E does not prevent or cure exudative diathesis,
whereas addition of as little as 0.05 ppm Se completely prevents this disease (Scott, 1980).

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Other Functions
Scott et al. (1982) reported additional functions of vitamin E,
including (1) normal phosphorylation reactions, especially of high-energy phosphate compounds, such as creatine phosphate and adenosine
triphosphate; (2) a role in synthesis of vitamin C; (3) a role in synthesisof ubiquinone; and (4) a role in sulfur amino acid metabolism. Pappu etal. (1978) reported on the role of vitamin E in vitamin B12 metabolism.
A deficiency of vitamin E interferes with conversion of vitamin B12 to its
coenzyme 5′-deoxyadenosylcobalamin and concomitantly metabolismof methylmalonyl-CoA to succinyl-CoA. Turley and Brewster (1993)
suggested that in humans, cellular deficiency of adenosylcobalamin may
be one mechanism by which vitamin E deficiency leads to neurologic
injury. In rats, vitamin E deficiency has been reported to inhibit vitamin
D metabolism in the liver and kidneys with the formation of active
metabolites and decreases in the concentration of the hormone-receptor
complexes in the target tissue. Liver vitamin D hydroxylase activity
decreased by 39%, 25-OHD3 1-hydroxylase activity in the kidneys by
22%, and 24-hydroxylase activity by 52% (Sergeev et al., 1990).

REQUIREMENTS
Estimated vitamin E requirements for selected animals and humans
are presented in Table 4.3. Scott (1980), after reviewing the literature,concluded that the minimum vitamin E requirement of normal animals
and humans is approximately 30 ppm of diet. Vitamin E requirements
are exceedingly difficult to determine because of the interrelationships
with other dietary factors (see Functions and Deficiency sections). The
requirement may be increased with increasing levels of PUFA, oxidizing
agents, vitamin A, carotenoids, gossypol, or trace minerals anddecreased with increasing levels of fat-soluble antioxidants, sulfur aminoacids, or Se (Dove and Ewan, 1990; Nockels, 1990; Hidiroglou et al.,
1992b; McDowell et al., 1996; Franklin et al., 1998). On otherwise adequate diets containing sufficient cystine and methionine and containing
a minimum of PUFA, vitamin E requirements appear to be low. This isevidenced by difficulties in producing deficiency signs on such diets
under optimum environmental conditions.
The levels of PUFA found in unsaturated oils such as cod liver oil,corn oil, soybean oil, sunflower seed oil, and linseed oil increase vitamin
E requirements. This is especially true if these oils are allowed to undergo oxidative rancidity in the diet or are in the process of peroxidation
170

V I TA M I N E■ Table 4.3

Vitamin E Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Growing
Milk replacer
Growing
Lactating cows and bulls
All classes

15–60 IU/kg
40 IU/kg
25 IU/kg
15 IU/kg
100 IU/kg

Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100 g intake)
Broilers, 0–8 weeks
Growing, 0–7 weeks
All classes
All classes
Growing, pregnant, lactating,
and working
MaintenanceAll classes
Growing
All classes
Growing
All classes
CatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
Children
Adults10 IU/kg
5 IU/kg
5 IU/kg
10 IU/kg
10 IU/kg
10–12 IU/kg
15–20 IU/kg

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1989a)
Morand-Fehr(1981)NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)

80 IU/kg
50 IU/kg
11–44 IU/kg
25 IU/kg
30 IU/kg
22 IU/kg
40 IU/kg
25–50 IU/kg
30–50 IU/kg
15–100 IU/kg
18 IU/kg
22 IU/kg
3–4 mg/day
6–7 mg/day
8–12 mg/day

NRC (1989b)
NRC (1989b)
NRC (1998)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1977)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

Goat
Chicken

Duck
Turkey
Sheep
Horse
Swine
Mink
Cat
Dog
Rabbit
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg α-tocopherol equivalents/day.

when consumed by the animal. If they become completely rancid before
ingestion, the only damage is the destruction of the vitamin E present in
the oil and in the feed containing the rancidifying oil. But if they are
undergoing active oxidative rancidity at the time of consumption, they
apparently cause destruction of body stores of vitamin E as well (Scottet al., 1982). The vitamin E requirement for dogs is five times higher
under conditions of high PUFA intake (NRC, 1985a). The amount ofvitamin E required per gram of PUFA is dependent on experimental conditions, species differences, levels and kinds of PUFA, and test used
(Lopez-Bote et al., 1997; McGuire et al., 1997). Nevertheless, for anumber of species, 0.6 IU of vitamin E per gram of PUFA is inadequate,
and 1 IU is a realistic minimum (Hoffmann-La Roche, 1972). A combi171

CHAPTER FOUR

nation of stress of infection and presence of oxidized fats in swine diets
was reported to exaggerate vitamin E needs still further (Tiege et al.,1978). These researchers reported that supplements of 100 IU of vitamin
E per kilogram of diet and 0.1 ppm Se did not entirely prevent deficiency lesions in weanling pigs afflicted with dysentery and fed 3% cod liveroil.
Of all factors, the most important determinant of vitamin E requirements is the dietary concentrations of unsaturated fatty acids. A diet that
contains high levels of fish oil may cause a threefold to fourfold increasein a cat’s daily requirement for α-tocopherol. Early cases of steatitis (yellow fat disease) occurred almost exclusively in cats that were fed acanned, commercial fish-based cat food, of which red tuna was the principal type of fish.
Since the PUFA content of membranes can be altered by dietary fats,
it is not surprising that the dietary requirement for vitamin E is closely
related to the dietary concentration of PUFAs. When a large amount ofpolyunsaturated fat is fed after it has been stripped of tocopherols, asmuch as 100 mg of α-tocopherol per kilogram of diet may be insufficient to protect against lipofuscin formation (Hayes et al., 1969).
Harris and Embree (1963) proposed a dietary α-tocopherol:PUFA
ratio (mg/g) of 0.6:1 as a minimum to protect against PUFA peroxidation. This is a step in being more accurate for determining vitamin E
requirements for animals; however, it should be realized that some
PUFAs require much more vitamin E to counteract detrimental effects
than others. Fish-oil PUFAs, which contain arachidonic acid (20:4),
docosapentaenoic acid (22:5), and docosahexaenoic acid (22:6), require
much more vitamin E for stabilization than typical plant PUFAs, such as
linoleic acid (18:2) and linolenic acid (18:3). The longer the carbon
chain of a fatty acid and the more double bonds, the greater the vitamin
E requirement. As an example, when 5% tuna oil (long carbon chain
and greater unsaturation) was substituted for 5% of the lard (shorter
carbon chain and less unsaturation), steatitis was severe in cats unsupplemented with vitamin E. The severity of steatitis was diminished bysupplemental vitamin E, but was not entirely prevented by 34 IU/kg of
diet. When 136 IU/kg of diet were provided, no lesions were seen
(Gershoff and Norkin, 1962). By-product organs (e.g., liver) in the dietwould also have more of the longer-chained, more unsaturated fatty
acids.
The amount of vitamin E needed to maintain adequate growth andreproduction would not necessarily be enough to ensure optimal
immune function as noted previously (Weiss, 1998). For example, in the172V I TA M I N E

rat, Bendich et al. (1986) reported that 15 IU/kg vitamin E of diet prevented muscle abnormalities and 50 IU/kg of diet was necessary to prevent red blood cell breakdown, but for maximum lymphocyte stimulation, 200 IU of vitamin E per kilogram of diet was required. In pigs, 150
IU/kg of supplemental vitamin E resulted in a higher activity of lysozyme
and a higher rate of yeast lysis (Riedel-Caspari et al., 1986). Reduced
mortality and increased humoral immune titers were reported whenchicks infected with Escherichia coli were supplemented with 150 or
300 mg of dl-α-tocopherol/kg (Heinzerling et al., 1974).
Stress, exercise, infection, and tissue trauma all increase vitamin E
requirements (Nockels, 1991). Handling and bleeding heifers periodically in a 10-day period resulted in a large decrease in the vitamin E content of red blood cells and a 62% decrease in neutrophil vitamin E levels (Nockels, 1991). Vitamin E supplementation to stressed calves
increased immune response (Golub and Gershwin, 1985). Nockels et al.
(1996) noted that for most sampled tissues, stress did not affect α-tocopherol concentration, although other indicators confirmed a deficiency.
Requirements of both vitamin E and Se are greatly dependent on the
dietary concentrations of each other (see Functions and Deficiency sections). As noted earlier, they are mutually replaceable above certain limits. Chicks consuming a diet containing 100 IU/kg vitamin E required
0.01 ppm Se, while those receiving no added vitamin E required 0.05
ppm Se (Thompson and Scott, 1969).
Vitamin E is known to reduce the Se requirement in at least two
ways: (1) by maintaining body Se in an active form, or preventing bodyloss; and (2) by preventing destruction of membrane lipids within themembrane, thereby inhibiting the production of hydroperoxides andreducing the amount of glutathione peroxidase needed to destroy peroxides formed in the cell. Selenium is known to spare vitamin E in at least
three ways: (1) it is required to preserve the integrity of the pancreas,
which allows normal fat digestion and thus normal vitamin E absorption;(2) it reduces the amount of vitamin E required to maintain the integrityof lipid membranes via glutathione peroxidase; and (3) it aids in some
unknown way in retention of vitamin E in the blood plasma.
Determination of vitamin E requirements is further complicatedbecause the body has a fairly large ability to store both vitamin E and
Se. Sows maintained on a diet deficient in vitamin E and Se produced
normal piglets during the first reproductive cycle of the deficiency, andclinical deficiency signs occurred only after five such cycles (Glienke andEwan, 1974). A number of studies to establish requirements for bothnutrients have underestimated the requirements by failing to account for173

CHAPTER FOUR

their augmentation from both body stores as well as experimental
dietary concentrations.
It has been concluded that in humans, a daily intake of 3 to 15 mgof tocopherol is required from natural diets (RDA, 1989). However, theallowances will not be adequate in individuals who, for a variety of reasons, do not absorb fat efficiently or who have medical conditions that
result in an abnormal vitamin E status in the blood and tissues. As inother species, the human requirement for vitamin E is related to dietaryintake of PUFA. However, in normal diets in the United States, this relationship is probably of little significance, inasmuch as primary dietary
sources of PUFA—vegetable oils, margarine, and shortening—are alsorich sources of vitamin E. This situation is not true when foods consumed contain the longer-chained fatty acids (e.g., fish oils).

NATURAL SOURCES
Many vitamin E analyses of foods and feedstuffs have been reported using a variety of analytical techniques; however, there is a lack ofcharacterization of individual tocopherols in the majority of analyses.
Total tocopherol analysis of a food or feedstuff is of limited value inproviding a reliable estimate of the biological vitamin E value. The
occurrence of tocopherols (and tocotrienols) other than alpha (Table4.4), as well as the prevalence of non–tocopherol-reducing substances innatural products, has led to analytical examination of these materials bytechniques capable of precise quantitation of individual species.
Because α-tocopherol is the most active form of vitamin E, many

■ Table 4.4

Tocopherols in Selected Feedstuffs (ppm)

Feedstuff
Barley
Corn
Oats
Rye
Wheat
Corn oil
Cottonseed oil
Palm oil
Safflower oil
Soybean oil
Wheat germ oil

α

β

γ

δ

4
6
7
8
10
112
389
256
387
101
1,330

3
—a
2
4
9
50
—
—
—
—
710

0.5
38
3
6
—
602
387
316
174
593
260

0.1
Trace
—
—
0.8
18
—
70
240
264
271

Source: Modified from Ullrey (1981).
aNo value reported.

174

V I TA M I N E

nutritionists prefer listing this form in feeds versus the unreliable total
tocopherol values. Some of the less active tocopherols, particularly γtocopherol, are present in mixed diets in amounts two to four times
greater than that of α-tocopherol. Cort et al. (1983), utilizing HPLC
assay procedures, which allow separation of alpha and non-alpha formsof both tocopherol and tocotrienols, determined that corn, corn gluten
meal, oats, barley, and wheat contained significant amounts of γtocotrienol, which has minimal biological activity. For purposes of calculating total vitamin E activity of mixed diets, milligrams of β-tocopherol should be multiplied by 0.5, those of γ-tocopherol by 0.1, andthose of δ-tocopherol by 0.3 (RDA, 1989). These three forms, in addition to α-tocopherol, provide the only significant vitamin E activity in
typical diets. If only α-tocopherol in a mixed diet is reported, the value
in milligrams should be increased by 20% (multiply by 1.2) to accountfor other tocopherols that are present, thus giving an approximation of
total vitamin E activity as milligrams of α-tocopherol equivalents. The
α-tocopherol concentrations of various foods and feedstuffs are listed inTable 4.5.
Vitamin E is widespread in nature, with the richest sources being
vegetable oils, cereal products containing oils, eggs, and liver. In nature,
the synthesis of vitamin E is a function of plants, and thus, their products are by far the principal sources. It is abundant in whole cereal
grains, particularly in germ, and thus in by-products containing thegerm. There is wide variation in vitamin content of particular feeds, withmany feeds having a threefold to tenfold range in reported α-tocopherol
values.
Milk is highly variable in vitamin E content. As a source of vitamin
E, there can be a fivefold seasonal difference in the α-tocopherol content
of cow’s milk. The vitamin E content of colostrum is of special importance for the newborn, because at birth, many species have very small
amounts of vitamin E in their tissues. Hidiroglou (1989) reported thatcolostrum from dairy cows has a mean value of 1.9 μg α-tocopherol per
milliliter and declines at 30 days to 0.3 μg α-tocopherol per milliliter. In
this study, milk tocopherol was raised from 0.3 μg/ml to 1.6 μg/ml 12hours after an intraperitoneal injection of dl-α-tocopheryl acetate.
Supplemental vitamin E administered to ewes at graded levels of 0, 15,30, and 60 IU (as dl-α-tocopheryl acetate) increased colostral α-tocopherol at a linear rate of 3.3, 6.8, 8.1, and 9.6 μg/ml, respectively (Njeru
et al., 1994a). In sows, colostrum α-tocopherol concentration can be elevated by increasing the gestation dietary level of vitamin E or via injection during the last 14 days of pregnancy (Chung and Mahan, 1995).
175

CHAPTER FOUR
■ Table 4.5

α-Tocopherol Content of Feeds (ppm)

SourceMean

Alfalfa meal, dehydrated 17% protein
Alfalfa meal, sun cured 13% protein
Alfalfa hay
Barley, wholeBeef, meat
Brewer’s grains, dried
ButterChicken, meatCorn, whole
Cottonseed meal
Distiller’s grains, dehydrated
Eggs
Fat, animal
Fish, halibut
Fish (seafood product), shrimp
Fish meal, herring
Fish meal, Peruvian
Lard
Linseed meal
Meat and bonemeal
Milo
Molasses, cane
Oats, wholePork, meat
Poultry by-products meal
Rice, brown
Rice, bran
Sorghum, grain
Soybean meal, solvent process
Wheat, whole
Wheat, bran

73
41
53
36
6
27
24
3
20
9
30
11
8
9
9
17
2
12
8
1
12
5
20
5
2
13.5
61
12
3
11
17

Range
28–121
18–61
23–102
22–43
5–8
17–48
10–33
2–4
11–35
2–16
17–40
8–12
2–16
4–13
6–19
8–31
1–3
2–30
3–10
1–2
10–16
3–9
18–24
4–6
1–4
13–14
34–87
10–16
1–5
3–15
15–19

Source: Adapted from Bauernfeind (1980) and Ullrey (1981).
Note: When only α-tocopherol in a mixed diet is available, multiply value by 1.2 toaccount for the other tocopherols present.

Colostrum α-tocopherol concentration was approximately five times
higher than later milks (Mahan and Vallet, 1997).
Animal by-products supply only small amounts of vitamin E, andmilk and dairy products are often poor sources. Eggs, particularly theyolks, make a significant contribution depending on the diet of the hen.
Wheat germ oil is the most concentrated natural source, and various
other oils, such as soybean and peanut, and particularly cottonseed, arealso rich sources of vitamin E. Unfortunately, most of the oilseed meals
that are part of pet-food diets are almost devoid of these oils because of
their removal by solvent extraction (Maynard et al., 1979). Green forage and other leafy materials, including good-quality hay, are very goodsources; alfalfa is especially rich. Concentration of tocopherols per unit
176

V I TA M I N E

dry matter in fresh herbage is between 5 and 10 times as great as that insome cereals or their by-products (Hardy and Frape, 1983). Variability
in forage vitamin E content is so great, both between and within farms,
that one must have current results or representative samples to ensure
proper vitamin E fortification programs (Harvey and Bieber-Wlaschny,1988). These authors indicated that previously published values on vitamin E content of forages are unacceptable for use in feed formulation.
Stability of all naturally occurring tocopherols is poor, and substantial losses of vitamin E activity occur in feedstuffs when processed andstored, as well as in manufacturing and storage of finished feeds(Coelho, 1991; Dove and Ewan, 1991; McDowell et al., 1996). Vitamin
E sources in these ingredients are unstable under conditions that promote oxidation of feedstuffs—heat, oxygen, moisture, oxidizing fats,
and trace minerals. Vegetable oils that normally are excellent sources ofvitamin E can contain extremely little if oxidation has been promoted.
Not only does oxidized oil have little or no vitamin E, but it will destroy
the vitamin of other feed ingredients and deplete animal tissue stores ofvitamin E. Rats fed a diet containing 15% oxidized frying oil had significantly lower α-tocopherol in plasma and tissues (Liu and Huang,
1995, 1996).
For concentrates, oxidation increases following grinding, mixing
with minerals, addition of fat, and pelleting. When feeds are pelleted,destruction of both vitamins E and A may occur if the diet does not contain sufficient antioxidants to prevent their accelerated oxidation underconditions of moisture and high temperature. Iron salts (e.g., ferric chloride) can completely destroy vitamin E. Dove and Ewan (1991) observedthat the rate of oxidation of natural tocopherols is increased in swine
diets containing increased levels of copper, iron, zinc, and manganese.
High dietary copper in swine diets decreased serum tocopherols (Doveand Ewan, 1990). Both nitrogen trichloride and chlorine dioxide, atconcentrations usually used to bleach flour, will destroy much of thevitamin E activity in flour. According to Moore et al. (1957), baking
destroyed 47% of remaining tocopherols in treated flour.
Artificial dehydration or processing of forages and grains will
reduce availability of tocopherol as well as that of Se. For example, Kinget al. (1967) reported that 80% of the vitamin E is lost in hay making,whereas ensiling or rapid dehydration of forages retains most of thevitamin. Vitamin E content in forage is affected by stage of maturity at
the time of forage cutting and the period between cutting and dehydration. Storage losses can reach 50% in 1 month, and losses during drying
in the swath can amount to as much as 50% within 4 days. Vitamin E
177

CHAPTER FOUR

losses of 54 to 73% have been observed in alfalfa stored at 33oC for 12
weeks, and 5 to 33% losses have been obtained with commercial dehydration of alfalfa.
In a study testing vitamin E stability, Orstadius et al. (1963) reported that vitamin E content of corn was reduced from 30 to 50 mg/kg toabout 5 mg/kg dry weight as a result of artificial drying at 100oC for 24hours under a continuous flow of air. Similarly, Adams (1973) reported
that artificial drying of corn for 40 minutes at 74oC produced an average 19% loss of α-tocopherol and 12% loss of other tocopherols. Whencorn was dried for 54 minutes at 93oC, the α-tocopherol loss averaged
41%. Young et al. (1975) reported a concentration of 9.3 and 20 mg/kg
α-tocopherol in artificially dried corn versus undried, respectively.
Preservation of moist grains by ensiling caused almost complete loss ofvitamin E activity. Corn stored as acid-treated (propionic or acetic-propionic mixture), high-moisture corn contained approximately 1 mg/kg
dry matter of α-tocopherol, whereas similar corn artificially dried contained approximately 5.7 mg/kg (Young et al., 1978). Apparently, damage is not due to moisture alone but to the combined propionic acidmoisture effect (McMurray et al., 1980). Further decomposition of αtocopherol occurs over a more extended period, until the grain eventually has α-tocopherol levels of less than 1 mg/kg, which is commonly
found in propionic acid-treated barley.

DEFICIENCY
Vitamin E displays the greatest versatility of all vitamins in the range
of deficiency signs. Deficiency signs differ among species and even within the same species. The amount of vitamin E required in diets can varydepending on such factors as levels of PUFAs, Se, antioxidants, and sulfur amino acids in feed. Deficiency diseases and compounds preventing
them are shown in Table 4.6.
Blaxter (1962) reported that muscular dystrophy seemed to be theone syndrome commonly encountered in vitamin E deficiency in all
species. He cited references indicating that muscular degeneration thatcan be prevented with vitamin E occurs in all laboratory and farm animals, including camels, buffalos, kangaroos, and quokkas. In some 20
different animal species, tocopherol deficiency leads to muscular dystrophy. Fundamentally, this is Zenker’s degeneration of both skeletal andcardiac muscle fibers. Connective tissue replacement that follows isobserved grossly as white striations in the muscle bundles (Smith, 1970).
Occurrence of muscular dystrophy is worldwide, but its incidence,178

■ Table 4.6

Vitamin E Deficiency Diseases As Influenced by Other Factors
Prevented byDisease
Reproductive failure
Embryonic degeneration
Type A
Type B
Sterility (male)
Neuropathy
Liver, blood, brain,
capillaries, pancreas
Necrosis
Fibrosis
Erythrocyte hemolysis
Plasma protein loss
AnemiaEncephalomalacia
Exudative diathesis
Kidney degeneration
Steatitis (ceroid)
Depigmentation

Experimental Animal

Tissue Affected

Rat, hamster, mouse, hen,turkeyCow, eweRat, guinea pig, hamster, dog,cock, rabbit, monkey
Chick, human

Vascular systemof embryoRat, pig
Chick, mouse
Rat, chick, human (prematureinfant), calf, dog, monkey
Chick, turkey
MonkeyChick
Chick, turkey
Rat, mouse, monkey, mink
Mink, pig, chick
Rat

Liver
Pancreas
Erythrocytes

PUFA
Influence

Vitamin
E

X

X—a

Serum albumin
Bone marrow
Cerebellum
Vascular system
Kidney tubular
Adipose tissue
Incisors

Antioxidant

X
Xb

X

Male gonads
Brain

Se

Sulfur
Amino
Acids

X

X

X
X

X
X
X
X

X

X
X

X
X
X
X
X
X
X
X

X
X

X
X
X
X
X
X
X

179

■ Table 4.6

Continued

180

Prevented byDiseaseNutritional myopathies
Type A (nutritional
muscular dystrophy)
Type B (whitemuscle disease)
Type CType D
Retinopathy
Dermatosis
Immunodeficiency

PUFA
Influence

Vitamin
E

Experimental Animal

Tissue Affected

Rabbit, guinea pig, monkey,
duck, mouse, mink, dog
Lamb, calf, kid, foal

Skeletal muscle

X

Skeletal and heart
musclesGizzard, heart
Skeletal muscle
Retinal pigment
epithelium(photoreceptor cells)
Skin
Reticuloendothelial

—a

Xb

—c
X

—a
X
X

X

X

X

X

TurkeyChickenDog, monkey, rat
Dog
Dog, chick, mouse, sheep, pig

Sources: Modified from Scott (1980) and Sheffy and Williams (1981).

Se

Antioxidant

Sulfur
Amino
Acids

?

X

aNot effective in diets severely deficient in selenium.
bWhen added to diets containing low levels of vitamin E.
cA low level (0.5%) of linoleic acid is necessary to produce dystrophy; higher levels did not increase vitamin E required for prevention.

V I TA M I N E

or at least diagnosis, particularly in a mild or subclinical form, varies
widely in different countries and regions within countries (McDowell,1997). Considerable research has revealed the positive relationshipbetween Se content in soil and geographical occurrence of muscular dystrophy that responds to vitamin E and Se.

Effects of Deficiency
Ruminants
White muscle disease (WMD; also known as nutritional muscular
dystrophy), a serious muscle degeneration disease in young ruminants,
is caused by Se deficiency but is influenced by vitamin E status. Whitemuscle disease occurs with two clinical patterns; the first is a congenital
type of muscular dystrophy in which young ruminants are stillborn or
die within a few days of birth after sudden physical exertion, such as
nursing or running. The second pattern (delayed WMD) develops after
birth; it is observed most frequently in lambs within 3 to 6 weeks ofbirth but may occur as late as 4 months after birth. The condition in
calves is generally manifested at 1 to 4 months of age.
Typically, WMD is characterized by generalized weakness, stiffness, and deterioration of muscles; affected animals have difficulty
standing (Figs. 4.3 and 4.4). Affected animals have difficulty standing
and exhibit crossover walking and impaired suckling ability because
the tongue musculature is affected (Muth, 1955). Calves with WMD
have chalky white striations, degeneration, and necrosis in the skeletal
muscles and heart (Fig. 4.3). Often, death occurs suddenly from heart
failure as a result of severe damage to the heart muscle. In calves withmilder cases, in which the chief clinical signs are stiffness and difficulty standing, dramatic, rapid improvement can result with vitamin E-Se
injections.
An acute and chronic as well as a peracute form of the disease canbe distinguished in older calves, usually already in the finishing period.
In particular, stress situations such as transport, regrouping, or abruptchanges in feed composition are generally considered precipitating factors. Sudden death without previous unmistakable signs of disease is themain feature of the peracute condition (Bostedt, 1980). The cause is usually found in advanced degeneration of the myocardium, and motor disturbances such as an unsteady gait or stiff-calf disease; hard lumbar,
neck, and forelimb muscles; muscle tremor; and perspiration areencountered in the acute form.
In Florida, the condition is most common in “buckling” calves that181

CHAPTER FOUR

A

C

B

Fig. 4.3 White muscle disease in a calf about 3 months old. (A) Lameness andgeneralized weakness of muscles can be seen. (B and C) Abnormal white areasin the heart muscle. (Courtesy of O.H. Muth, College of Veterinary Medicine,
Oregon State University.)come off the truck or out of the processing chute with weakness of rearlegs, buckling of fetlocks, and frequently a generalized shaking or quivering of muscles (Figs. 4.5 and 4.6). Many calves become progressively
worse until they are unable to rise and may appear to be paralyzed.
Many animals will be down or continue to buckle for extended periods,
and death loss is high in severe cases. Calves with excitable temperaments appear to be most affected. Postmortem examination of affected
calves reveals pale chalky streaks in the muscles of the hamstring and182

V I TA M I N E

Fig. 4.4 Vitamin E-selenium deficiency in sheep is known as stiff-lamb diseaseor white muscle disease. This lamb is unable to stand as a result of tissue
degeneration. (Courtesy of O.H. Muth, College of Veterinary Medicine,
Oregon State University.)

back, and the heart, rib muscles, and diaphragm may also be affected(McDowell, 1985).
In lambs, WMD (also known as stiff-lamb disease) takes a course
similar to that found in calves. There are motor disturbances such as
unsteady gait (Fig. 4.4); stiffness in rear quarters, neck, and forelimbmuscles; and arched back. Muscle tremor and perspiration are encountered in the acute form. On necropsy, the disease is manifested as white
striations in cardiac muscles and is characterized by bilateral lesions inskeletal muscles. A gradual swelling of the muscles, particularly in the
lumbar and rear thigh regions, gives the erroneous impression of anespecially muscular young animal. In addition to the peracute form
encountered in calves (changes primarily in the myocardium), chronic
cardiac muscle degeneration is also found in the lamb. Despite good initial development, affected lambs quickly lose weight after the third weekof life and are unthrifty. They try to avoid any strain and usually standapart from the herd. Cardiac arrhythmia and increased heart rate can183

CHAPTER FOUR

Fig. 4.5 Vitamin E-selenium deficiency is seen as flexion of the hock and fetlock joints as a result of decreased support by the gastrocnemius muscle, which
is severely affected by myodegeneration. (Courtesy of Bob Mason andUniversity of Florida.)

result even after slight exercise. In the advanced stage, animals eat little
feed and rapidly waste away.
Other conditions responsive to vitamin E-Se are not restricted toyoung animals and relate to unthriftiness (“illthrift”), occurring inlambs and hoggets at pasture (Underwood, 1981). Yearling sheep canalso be affected by WMD. In sheep of 9 to 12 months of age, the disease
is frequently observed following driving, with the rapid onset of listlessness, stiffness, inability to stand, prostration, and, in the most acutecases, death within 24 hours (Andrews et al., 1968). Hartley and Grant(1961) reported that the incidence of barren ewes was reduced fromover 30 to 5% with Se administration. Farms in New Zealand have hadlamb losses as high as 40 to 50%. Vitamin E reduced losses by only
60%; Se, by 96%.
The immune response in sheep has been improved with supplemental vitamin E. Vitamin E improved disease resistance in lambs challenged
with chlamydia (Stephens et al., 1979). Reffett et al. (1988) reported
that vitamin E and Se independently increased the immune response oflambs challenged with a viral pathogen. Myopathic lambs exhibited low
lymphocyte responses when deficient in vitamin E and Se (Finch and184

V I TA M I N E

Fig. 4.6 Vitamin E-selenium deficiency in cattle is manifested as white muscle
disease or necrosis of the gastrocnemius muscle; chalky white streaks are evident in the belly of the muscle. (Courtesy of Bob Mason and University of
Florida.)Turner, 1989; Turner and Finch, 1990). The poor lymphocyte responsesof the lambs with nutritional myopathy was rapidly reversed by intramuscular administration of these nutrients, with the prophylaxis mosteffective during the first 5 weeks of life (Finch and Turner, 1989).
Deficiency of vitamin E and/or Se in the goat, as in other ruminants,
results mainly in WMD. Kids especially suffer from this disturbance ofmuscle metabolism, as they are born with little or no reserves of the fatsoluble vitamins A, D, and E. Sudden death of young kids under 2 weeks
of age may reveal postmortem evidence of muscle disease and degeneration in the heart or diaphragm. In older kids and mature animals, thedisease may occur after sudden exercise, and the animals show bilateral
stiffness, usually in their hind legs.
In preruminant calves, WMD has been easily induced by feeding
polyunsaturated oils; however, it was thought unlikely that unsaturated
fats were responsible for the disease in ruminating calves because of theapparent near 100% hydrogenation of all unsaturated fatty acids by therumen microflora (Noble et al., 1974). However, one study indicated185

CHAPTER FOUR

that unsaturated fatty acids from lipids in grasses can act as substitutes
for the peroxidative challenge in nutritional muscular dystrophy incalves (McMurray and Rice, 1984). Nutritional degenerative myopathy
in older calves occured most frequently at turnout to spring pasture(Anderson et al., 1976). McMurray et al. (1980) showed that PUFAs
were capable of escaping ruminal hydrogenation at turnout, resulting ina threefold increase of plasma linolenic acid within 3 days of turnout.
Rice et al. (1981) showed that linolenic acid, if protected from ruminalhydrogenation, rapidly reaches high levels in blood and is associatedwith a rise in plasma creatine phosphokinase, indicating musculardegenerative myopathy.
Most nutritional myopathy cases have involved young ruminants,
with effects less fully described for adult animals. However, degenerative
myopathy in adult cattle has been reported (Van Vleet et al., 1977;
Gitter and Bradley, 1978; Hutchinson et al., 1982), and a group of yearling Chianina heifers experienced abortion, stillbirth, and periparturient
recumbency (Hutchinson et al., 1982). Necropsy and tissue analyses
revealed myodegeneration and a combined deficiency of vitamin E and
Se. Rapid growth in these heifers, coupled with stresses of late pregnancy and parturition, may have contributed to this vitamin E deficiency. A
myopathic condition affecting yearling cattle was reported by Bartonand Allen (1973) and was associated with animals fed grains treatedwith propionic acid, which is known to destroy vitamin E.
Attempts to establish a practical role for vitamin E in ruminant
reproductive deficiencies of both males and females have been limited.
The relationship to reproduction is of special interest since early rat
research demonstrated that reproductive failure was a key feature ofvitamin E deficiency. In one experiment, four generations of female andmale dairy cattle were fed diets low in vitamin E (Gullickson et al.,1949). Although growth, reproduction, and milk production were normal, several cattle died suddenly of apparent heart failure between 21
months and 5 years of age. Large doses of vitamins A, D, E, and C werereported to favorably affect some characteristics of semen and sperm(Kozicki et al., 1981).
Recent research has shown that vitamin E supplementation is beneficial for male reproduction in bulls fed high concentrations of gossypol.
Velásquez-Pereira et al. (1998) reported that bulls receiving 14 mg offree gossypol per kilogram of body weight had a lower (P < 0.05) percentage of normal sperm than those that also received supplemental
vitamin E (31 and 55%, respectively; Table 4.7). Likewise, sperm pro-186

V I TA M I N E

duction per gram of parenchyma and total daily sperm production werehigher (P < 0.05) when gossypol-treated animals also received vitamin
E. Bulls receiving gossypol exhibited more sexual inactivity (P < 0.05)
than bulls receiving other treatment (Table 4.8). Vitamin E supplementation in bulls receiving gossypol improved number of mounts in thefirst test and time of first service in the second test. The final conclusion
based on these data is that vitamin E is effective in reducing or eliminating important gossypol toxicity effects for male cattle (VelasquezPereira et al., 1998, 1999).
From a different aspect of reproduction in dairy cattle, Harrison etal. (1984) reported that supplemental vitamin E was required in addition to Se for prevention of retained placenta. Groups administered vitamin E alone, Se alone, and controls had a retained-placenta incidence of
17.5% compared to 0% for animals receiving both vitamin E and Se.
Other research found that the incidence of retained placenta (22.1%)
was not reduced by a combination of vitamin E and Se or by Se alone(Hidiroglou et al., 1987). In high-producing dairy goats, deficiency manifested itself in poor involution of the uterus, with accompanying
retained placenta and metritis following kidding (Guss, 1977).
Adequate amounts of vitamin E in the diet are needed to prevent■ Table 4.7

Relationship of Gossypol and Vitamin E on Semen Characteristics of Dairy
Bulls
Treatment

Item

Controla+ Gossypolb+ Gossypol + Vitamin Ec

Normal, %64.7 + 6.4g

31.4 + 7.4h

54.6 + 6.4g

Abnormal (DIC)d, %4.4 + 1.3g

13.4 + 1.5h

4.8 + 1.2g

DSPGe (x106/g)

14.6 + 1.0g

10.2 + 1.0h

17.6 + 1.0g

DSPf (x109)

3.2 + 3.0g

2.2 + 3.0h

4.1 + 3.0g

Source: From Velásquez-Pereira et al. (1998).
Note: Least square means + SEM.
aDiet based on soybean meal, corn, and 30 IU vitamin E/kg of supplement.
bDiet containing 14 mg free gossypol/kg body weight per day and 30 IU vitamin E/kg of
supplement.
cDiet containing 14 mg free gossypol/kg body weight per day and 4,000 IU vitamin
E per bull per day.
dMidpiece abnormalities evaluated in isotonic formal saline using differential-phasemicroscopy.
eDaily sperm production per gram of parenchyma.
fDaily sperm production total.
g,hMeans in a row with different superscripts differ when P < 0.05.

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CHAPTER FOUR
■ Table 4.8

Effects of Gossypol and Vitamin E on Sex Drive of Holstein Bulls
Month

Controlb+ Gossypolc+ Gossypol + Vitamin Ed

Libido score

16

No. of mounts

1216

10.4 + 0.7

8.9 + 0.5

10.0 + .6

5.7a + 1.2
9.4 + 1.3

3.2b + 1.1
9.5 + 0.9

6.3a + 1.2
7.9 + 1.1

No. of services

16

2.4 + 0.5

1.7 + 0.4

2.3 + 0.4

Time of mounts
(sec)

16

38a + 7

29a,b + 5

21b + 5Time of firstservice (sec)1216

223 + 94
154a,b + 71

232 + 69
213a + 52

206 + 69
69b + 60

Sexual inactivity
(min)

12
16

1.2d + 0.8
0.1 + 0.2

3.9c + 0.7
0.1 + 0.1

1.7d + 0.8
0.2 + 0.2

Item

Source: From Velásquez-Pereira et al. (1998).
Notes: Each treatment represented eight animals.
Animals received 14 mg free gossypol per kg body weight (except controls).
Vitamin E–treated animals received 4,000 IU vitamin E per day as dl-αtocopheryl acetate.
a,bMeans with different superscripts in the same row differ (P < 0.01).
c,dMeans with different superscripts in the same row differ (P < 0.05).

oxidative flavors in milk. However, the cost is high, with efficiency oftransfer into milk less than 2% (NRC, 1989a).

Swine
In pigs, most signs of vitamin E deficiency have been associated withSe deficiency, and scientists usually refer to vitamin E and/or Se deficiency since it is not clear which is involved, and generally, dietary levels of both must be low to bring about deficiency signs and lesions. Since
the early 1950s, reports in the European literature have revealed tissuedegeneration signs in swine under field conditions associated with vitamin E deficiency; the significance of Se deficiency was not realized until
1957.
Muscular dystrophy and hepatosis dietetica (toxic liver dystrophy)
were particularly widespread in the swine industry in Sweden. Obel(1953) reported that records from the State Veterinary Medical Instituteof Stockholm from 1947 to 1952 revealed that of 4,382 pigs autopsied,over 10% suffered from hepatosis dietetica.
Vitamin E-Se deficiencies have been readily produced in swine diets
through use of both highly unsaturated fats (e.g., cod liver oil) and rancid fats. However, naturally occurring vitamin E-Se deficiencies were notreported in the United States until the late 1960s (Michel et al., 1969),
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V I TA M I N E

and in the 1970s, they became widespread. High incidence of vitamin ESe deficiencies in swine was believed to be due to a number of factors(Trapp et al., 1970), including (1) swine raised in complete confinement
without access to pasture, (2) low Se content in midwestern U.S. feeds,(3) solvent-extracted protein supplements low in vitamin E, (4) limited
feeding programs for sows, (5) loss of vitamin E and Se from corn due
to oxidation as a result of air and heat drying or storing high-moisturegrains, and (6) selection of meatier-type pigs that require more Se.
Evidence also suggests that moldy feed in bulk-holding bins may produce mycotoxins that either inhibit the uptake of vitamin E in the small
intestine or affect the antioxidant balance of cells.
An increase in confinement rearing of swine on concrete floors or
slats has been accompanied by a decrease in the utilization of pastureand forages. Such crops not only are excellent sources of vitamin E but
also provide the more highly available form of the vitamin, α- versus γtocopherol.
Vitamin E-Se deficiency in swine is often associated with sudden
death. In most cases, clinical signs of the condition were not observed
prior to death (Michel et al., 1969; Trapp et al., 1970), although occasionally, pigs were observed with clinical signs of icterus, difficult locomotion, reluctance to move, and weakness. Clinical signs also include
peripheral cyanosis (particularly of the ears), dyspnea (abdominal respiration), and weak pulse, all occurring shortly before death. In manycases, the faster-growing, more thrifty-appearing pigs died suddenly.
The most common pathological lesions include massive hepatic
necrosis (hepatosis dietetica) (Figs. 4.7 through 4.9), degenerative
myopathy of cardiac and skeletal muscles (Figs. 4.10 and 4.11), edema,esophagogastric ulceration, icterus, nephrosis, hemoglobinuria, acutecongestion, hemorrhaging (Fig. 4.12) in various tissues (Trapp et al.,1970; Piper et al., 1975), and yellowish discoloration of adipose tissue(yellow fat).
Many pathology reports of vitamin E-Se deficiency have noted thatthe most striking lesion was liver necrosis (Trapp et al., 1970); however,bilateral paleness of skeletal muscles was the gross lesion most commonly found. In some pigs, microscopic lesions in liver were either
absent or minimal, whereas changes in skeletal muscles were extensive.
In other cases, the reverse was true.
Other conditions reported in swine herds with vitamin E-Se deficiency include mastitis-metritis-agalactia syndrome (MMA) in sows,spraddled rear legs in newborn pigs, gastric ulcers, infertility, and poorskin condition. These conditions were believed initially to be unrelated
189

CHAPTER FOUR

Fig. 4.7 Lesions in growing pig fed
diet low in vitamin E (7.0 IU α-tocopherol/kg diet and selenium (0.061
ppm). Liver with severe acute lesions
is characteristic of hepatosis dietetica,
consisting of a mosaic pattern ofdeep red and yellow lobules of massive coagulation necrosis. (Courtesyof L.R. McDowell, R.C. Piper, andWashington State University.)

Fig. 4.8 Lesions in growing pig fed
diet low in vitamin E (7.0 IU α-tocopherol/kg diet) and selenium (0.061
ppm). In subacute dietary massive
hepatic necrosis, the lesions are morechronic than those seen in the liver of
pig in Fig. 4.7, in that many hepatic
lobules are atrophic and collapsed,causing a pitted appearance on thesurface of the liver. (Courtesy of L.R.
McDowell, R.C. Piper, andWashington State University.)

to pig deaths due to vitamin E-Se deficiency. However, after supplementation with dietary vitamin E or injections of Se and vitamin E, a noticeable reduction in these conditions occurred (Trapp et al., 1970).
Whitehair et al. (1983) provided evidence that the MMA syndrome may
be ameliorated by supplementation of the gestation-lactation diet withvitamin E and Se. In one experiment, vitamin E was shown to help
190

V I TA M I N E

Fig. 4.9 Lesions in liver of growing pig fed diet low in vitamin E (7.0 IU αtocopherol/kg diet) and selenium (0.061 ppm). Individual hepatic lobule has
undergone acute massive coagulation necrosis, and the necrosis cells are being
replaced by blood. Note the normal adjacent lobule. (Courtesy of L.R.
McDowell, R.C. Piper, and Washington State University.)

Fig. 4.10 Lesions in heart of growing pig fed diet low in vitamin E (7.0 IU αtocopherol/kg diet) and selenium (0.061 ppm). Note the degenerative nutritional cardiac myopathy and the large pale areas due to degeneration andnecrosis of myocardial fibers that are most severe along the inner border of theleft ventricle. (Courtesy of L.R. McDowell, R.C. Piper, and Washington State
University.)

reduce the incidence of MMA from 50 to 14% (Ullrey, 1969).
Occurrence of MMA was reduced from 39 to 24% in two studies
involving 191 farrowings (Ullrey et al., 1971). Diets low in Vitamin ESe for long periods will affect reproductive efficiency of boars (MarinGuzman et al., 1997). Sperm motility was found to decline, and the percentage of abnormal sperm increased.
191

CHAPTER FOUR

Fig. 4.11 Lesions in skeletal muscle of pig fed diet low in vitamin E (7.0 IU α-tocopherol/kg diet) and selenium (0.061 ppm). A muscle fiber (low power) of the biceps
fermoris is undergoing degeneration, surrounded by normal fibers. Cytoplasm isbreaking up into granules from enzymatic digestion as the fiber is invaded by macrophages. (Courtesy of L.R. McDowell, R.C. Piper, and Washington State University.)

Fig. 4.12 Skin lesion in growing pig fed diet low in vitamin E (7.0 IU α-tocopherol/kg diet) and selenium (0.061 ppm). The skin is from a pig with severecongestion and hemorrhage. The hemorrhage extended throughout the epidermis, dermis, and subcutaneous fat and down to the cutaneous musculature.
(Courtesy of L.R. McDowell, R.C. Piper, and Washington State University.)

Studies by Tiege et al. (1978) have shown that susceptibility of pigs
to dysentery resulting from exposure to the spirochete Treponema hyodysenteriae was greatly increased by the combined dietary deficiencies ofvitamin E and Se. In studies with young pigs, supplemental vitamin E has
been beneficial in increasing the humoral response against sheep red
blood cells (Peplowski et al., 1981; Morrow et al., 1987). Depressed
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V I TA M I N E

peripheral blood lymphocytes and polymorphonuclear cell immune functions were found in vitamin E-restricted sows (Wuryastuti et al., 1993).
The vitamin E-Se deficiency syndromes (nutrition-related microangiopathy; nutritional hepatic dystrophy; muscle degeneration in theback, pelvis, and upper thigh) can, however, also be found in the fattening and reproductive stages of pig production (Bostedt, 1980). At an
early age, it is particularly myocardial damage, also known as nutritional microangiopathy or mulberry heart disease, that may cause substantial losses within a litter. This is the most serious of disorders, since
when heart muscle tissue is damaged, the result is usually sudden death.
There may be hemorrhagic lesions within the heart that give the characteristic mulberry appearance of mulberry heart disease.
There is a low tolerance of vitamin E- and Se-deficient baby pigs to
intramuscular injections of iron dextrose for prevention of anemia. At 2
or 3 days, piglets die from iron shock if given routine treatment withiron, with death resulting from an iron-induced lipid peroxidation in tissues. Pretreatment with vitamin E, Se, or ethoxyquin was protective
against toxic effects of injectable iron (Tollerz and Lannek, 1964; Hill etal., 1999).
Maximum incidence of death due to vitamin E-Se deficiency generally occurs at 6 to 8 weeks of age, with the incidence declining up to thesixteenth week of life; however, conceptuses can be adversely affectedprior to parturition, resulting in stillborn pigs (Putnam, 1984). Vitamin
E deficiency can also occur in fattening and reproductive stages of swine
production (Bostedt, 1980). Clinical conditions characterized by cellular
damage most often occur after a period of stress, such as change of feedor housing, transportation, or weaning. A Michigan survey diagnosed
vitamin E-Se deficiencies in swine herds, with mortality ranging from 3
to 10% (Michel et al., 1969; Trapp et al., 1970). One producer, however, lost approximately 300 of 800 pigs weaned.
It has been realized for many years that vitamin E-deficient animals
are more subject to the effects of stress than normal animals. The concept of stress is difficult to define, but experience has shown that dietaryand environmental abnormalities of various kinds can lead to clinicalsigns of disease and death in animals deprived of vitamin E. Death isoften associated with unaccustomed muscular activity. Incidence ofdeath in baby pigs is greatly increased because of fighting when animals
are weaned and mixed with different litters. Castration, an additionalcause of stress, has been implicated as a cause of early death in pigs deficient in vitamin E and Se (Piper et al., 1975).

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CHAPTER FOUR

Fig. 4.13 Exudative
diathesis resulting fromvitamin E-selenium
deficiency in chicks.
Note profuse subcutaneous edema. (Courtesyof L.E. Krook, Cornell
University.)

Poultry
Vitamin E deficiency in poultry can result in at least three conditions: exudative diathesis (Fig. 4.13), with signs of subcutaneous edema
and, in severe cases, blackening of the affected parts, apathy, and inappetence; encephalomalacia (“crazy chick disease”) (Fig. 4.14), characterized by ataxia, head retraction, and “cycling” with legs; and muscular dystrophy (Fig. 4.15) (Scott et al., 1982).

CHICKENS
Exudative diathesis in chicks is a severe edema produced by amarked increase in capillary permeability. The subcutaneous edema
soon progresses to a hemorrhagic stage, producing a blue-green discoloration of the skin. Affected chicks show reduced spontaneous activity
and food intake; if not treated with vitamin E or Se, they survive usually no more than 2 to 6 days. Both vitamin E and Se are involved in prevention of exudative diathesis and nutritional muscular dystrophy. In
diets severely deficient in Se, however, vitamin E does not prevent orcure exudative diathesis, whereas addition of as little as 0.05 ppm of
dietary Se completely prevents this disease.
Encephalomalacia generally affects chicks from 2 to 6 weeks of age
and results from hemorrhages and edema within the cerebellum.
Degeneration of the Purkinje layer of cells in the cerebellum results in
nervous signs typified as sudden prostration, with toes and legs outstretched, toes flexed, and head outstretched (NRC, 1994). At least one
important function of vitamin E is to interrupt the production of free
radicals at the initial stage of encephalomalacia. The quantitative need194

V I TA M I N E

Fig. 4.14
Encephalomalacia in achick fed a vitamin Edeficient diet. This disease is caused by deficiency of vitamin E or
antioxidants. Selenium
supplementation of thediet will not prevent it.
(Courtesy of M.L.
Scott, Cornell
University.)
Fig. 4.15 Nutritional
muscular dystrophy inchick fed a vitamin Edeficient diet low in
sulfur amino acids. The
diet contained anantioxidant and 0.1
ppm of selenium to prevent encephalomalacia
and exudative diathesis.
Note the white degenerated muscle fibers inthe breast and thigh.
(Courtesy of M.L.
Scott, Cornell
University.)for vitamin E for this function depends on the amount of linoleic acid in
the diet. Selenium is ineffective in preventing encephalomalacia, while
synthetic antioxidants are partially effective. The fact that low concentrations of antioxidants are capable of preventing encephalomalacia inchicks but fail to prevent exudative diathesis or muscular dystrophy inthe same chicks strongly suggests that in preventing encephalomalacia
vitamin E acts as an antioxidant.
When vitamin E deficiency is accompanied by sulfur amino aciddeficiency, chicks show severe nutritional muscular dystrophy, especially of breast muscle, at about 4 weeks of age. Cystine is likewise effective
in preventing nutritional muscular dystrophy in vitamin E-deficient
chicks. Cystine, however, is apparently ineffective in preventing the dystrophic condition in other animals. Although vitamin E and Se are gen195

CHAPTER FOUR

erally both highly effective in preventing exudative diathesis, Se is only
partially effective in protecting against muscular dystrophy in chicks
when added in the presence of a low level of dietary vitamin E. Muchlarger quantities of Se are required to reduce the incidence of dystrophy
in chicks receiving a vitamin E-deficient diet low in methionine and cystine (Scott et al., 1982).
Prolonged vitamin E deficiency can result in permanent sterility.
Hatchability of eggs from vitamin E-deficient hens is reduced (NRC,
1994), and embryonic mortality may be high during the first 4 days of
incubation and during later stages as a result of circulatory failure.
Males become infertile because sperm become incompetent(Friedrichsen et al., 1980).

TURKEYS AND OTHER POULTRY SPECIESThe combined deficiency of vitamin E and Se in poults was found to
produce a mild type of exudative diathesis (Creech et al., 1957). Thiscondition was characterized by hemorrhaging on the inner margins of
the thighs and caudal breast muscles; in contrast to the exudative diathesis of the vitamin E- and Se-deficient chick, it involved only a mildedema. In ducklings, exudative diathesis would appear to be more similar to that of the chick, i.e., green edema of the subcutaneous tissues canbe seen most frequently on the thigh, with associated petechial hemorrhages of the thigh musculature (Combs and Combs, 1986). The appearance of exudative diathesis is infrequent and occurs in association with
only the more severe cases of nutritional muscular dystrophy in deficient
ducklings (Jager, 1977). In Japanese quail, the combined deficiency ofvitamin E and Se has only produced exudative diathesis in some animals.
Degeneration of the smooth muscle of the gizzard is the most characteristic sign of Se deficiency in the young turkey poult. In marked contrast to the skeletal myopathy of the vitamin E-deficient chick, gizzard
myopathy in vitamin E- and Se-deficient poults is not prevented bydietary sulfur containing amino acids but is completely prevented bysupplements of Se (Walter and Jensen, 1963). However, the dietary levelof vitamin E affects the amount of Se required for the prevention of thedisorder. It was necessary to use a basal diet low in methionine and vitamin E, as well as Se, in order to produce gizzard myopathy experimentally. Muscular dystrophy in ducklings is characterized by degenerationof the sarcoplasmic reticulum and mitochondria of the smooth muscleof the duodenum and gizzard, and are prevented with either vitamin E
or Se.
Vitamin E deficiency is also known to reduce hatchability in turkey
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V I TA M I N E

eggs (Jensen and McGinnis, 1957). Turkey embryos deficient in vitamin
E may have eyes that protrude with a bulging of the cornea. In Japanesequail, embryonic survival (i.e., egg hatchability) was markedly depressedamong females reared to maturity with vitamin E- and Se-deficient diets.
Many of the surviving progeny of vitamin E- and Se-deficient females
showed extreme generalized muscular weakness and prostration after
hatching (Jensen, 1968).

HorsesThe newborn foal that is born with nutritional muscular dystrophy
usually exhibits various clinical signs in this acute phase of vitamin E-Se
deficiency. Animals can hardly stand up and give the impression of general weakness. After laborious attempts to struggle to its feet, the foal
stands rather awkwardly and stiffly. If neck muscles are affected, suckling is substantially impaired, and the foal is unable to raise its head tothe mother’s udder, although it gives the impression of wishing to suckle (Bostedt, 1980). Changes affect predominantly pectoral, intercostal,and diaphragm muscles and result in accelerated, intermittent, primarily abdominal respiration that can be incorrectly diagnosed as bronchialpneumonia or dry pleurisy. Movement of the thorax appears to cause
pain. Dysphagia is a common finding (Dill and Rebhun, 1985), perhaps
because pain is especially prominent during extension and rotation of
the head and neck (Moore and Kohn, 1991). The dysphagia might also
be due to the weakening of the lingual and pharyngeal muscles and eventhe masticatory muscles. Cardiac arrhythmia is another clinical sign that
occurs as a result of muscle changes in the heart.
As with calves, the musculature of the tongue may be affected. In
spite of their obvious appetite, the animals are unable to swallow milk,
and the slightly opened mouth allows milk to leak out or trickle fromtheir nostrils. The stomach is empty and the belly drawn up, so that theperson in charge of the mare frequently consults a veterinarian for what
is believed to be a lack of milk. Foals affected in this manner usually die
during the first few days of life; the animal’s death is accelerated byhypostatic or deglutition pneumonia or by neonate infections arisingfrom immunological disorders caused by an inadequate intake ofcolostrum or from cardiovascular insufficiency (Bostedt, 1980).
In older foals (6 to 12 weeks), the progressive degeneration of motor
muscles rather than changes in the head and thoracic muscles are thefirst indications of the disease. Typical signs are an increasingly clumsy
gait and unsteady movements of the hindquarters. These foals also lie
down a great deal and can hardly be made to stand (Bostedt, 1980). The
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CHAPTER FOUR

urine may be coffee-colored because of myoglobin released from damaged muscle cells. As the condition worsens, the foal remains permanently in the lying position.
Combinations of vitamin E and Se have been used in the treatmentof the “tying-up” syndrome in horses (Hintz, 1996), which is characterized by lameness and rigidity of the loin muscles. However, no experimental evidence to confirm the value of vitamin E in the condition has
been provided. Likewise, published studies concerning the influence ofvitamin E on reproduction have been contradictory.
It has been suggested that vitamin E alone or with Se is essential forthe development of resistance to diseases such as influenza and tetanus
in horses (Hintz, 1996). Horses fed a diet containing 80 IU of vitamin E
per kilogram had greater immune responses than when fed diets containing 18 IU per kilogram (Baalsrud and Overnes, 1986).
Vitamin E supplementation has been claimed to enhance performance in horses, but few controlled studies have been conducted. Putnam(1986) concluded, apparently from field observations, that performance
could be enhanced by a daily intake of 2,000 mg of vitamin E. Dewes(1981) reported that the performance of horses might be improved byvitamin E-Se supplementation due to relief from pain in the musculatureof the neck.
It is hypothesized that vitamin E plays an oxidative role in equine
motor neuron disease (EMND) and equine degenerative myeloencephalopathy (EDM). Both are neurodegenerative diseases that result ina loss of motor neurons and, subsequently, muscle mass. The absence of
pasture, the feeding of marginal-quality hay, and high concentrates without additional vitamin E supplementation in part help to explain vitamin E-deficiency states in horses with EMND and EDM. Thus far, definitive causes for both EMND and EDM remain elusive. However, pathological evidence for each disorder presents a strong case for vitamin E
involvement (NRC, 1989b; Hintz, 1996).

Other Animal Species
DOGS
Definite clinical cases of vitamin E deficiency have been well documented in dogs. Particularly prominent indications of vitamin E deficiency were degeneration of skeletal muscle associated with muscle
weakness, degeneration of testicular germinal epithelium and failure of
spermatogenesis, failure of gestation, weak and dead pups, and brown
pigmentation (lipofuscinosis) of intestinal smooth muscle (NRC,198

V I TA M I N E

1985a).
Van Kruiningen (1967) reported a brown bowel in malabsorbing
boxer dogs with granulomatous colitis, and the condition was eventually reproduced experimentally by Hayes et al. (1969), who fed varying
levels of polyunsaturated fat to vitamin E-deficient weanling puppies
over a 16-week period. The puppies developed an increased susceptibility to hemolysis of red blood cells as well as brown bowel, mild muscle
degeneration, and neural-axonal dystrophy.
Retinal degeneration was described by Hayes and Rousseau (1970)
and reproduced experimentally (Riis et al., 1981) in puppies fed a dietcontaining stripped corn oil. In as few as 3 months, ophthalmoscopic
lesions were visible, which represented lipid peroxidation and disruptionof photoreceptors, with accumulation of lipofuscin pigment.
The effect of vitamin E deficiency on reproduction has been reported by Anderson et al. (1939; 1940) and Elvehjem et al. (1944), who
described a nutritional deficiency syndrome in puppies from bitches fed
evaporated or pasteurized milk diets for extended periods. Lactatingdams and puppies developed festering skin lesions, and the puppies
developed muscle weakness and hemorrhages of body cavities andbrain. Other pups were stillborn or died shortly after birth. The syndrome was alleviated by weekly feeding of 40 mg of vitamin E to thebitch during pregnancy.
A combined vitamin E-Se deficiency described by Van Vleet (1975)
included muscle weakness, subcutaneous edema, anorexia, depression,
dyspnea, and coma. Pathological examination revealed extensive skeletal muscle degeneration and regeneration, focal subendocardial necrosis,
lipofuscinosis, and renal mineralization.
Deficiency of vitamin E has been implicated in the development ofcertain dermatological disorders in dogs (Anderson et al., 1939;
Worden, 1958; Sheffy, 1979; Scott and Walton, 1985; Scott and Sheffy,
1987; Miller, 1989; Codner and Thatcher, 1993). The occurrence of
these skin disorders has been associated with decreased blood levels ofvitamin E. It has been postulated that subclinical vitamin E deficiency
causes suppression of the immune system, which in turn increases a
dog’s susceptibility to Demodex (skin lesions caused by the demodectic
mange mite, Demodex canis). When a group of dogs with demodicosis
was treated with supplemental vitamin E, significant levels of improvement were reported. However, other researchers have been unable toreproduce these results.
Scott and Sheffy (1987) experimentally produced vitamin E deficiency skin disorders. They were characterized clinically by an early ker199

CHAPTER FOUR

atinization defect (seborrhea sicca), a later inflammatory stage (erythroderma), and a tendency to develop secondary pyoderma. The vitamin E
anti-inflammatory effect may be related to stabilization of cell and lysosomal membranes against damage induced by free radicals and peroxides. By reducing oxidative damage to cells, vitamin E may inhibit
immune-mediated or neoplastic skin diseases associated with ultraviolet
irradiation.
Vitamin E therapy has been reported to be effective in dogs with discoid lupus erythematosus (Scott et al., 1983) and has been suggested astherapy for dogs with dermatomyositis (Hargis et al., 1985; Muller etal., 1989). Vitamin E supplementation has also been evaluated in dogs
with acanthosis nigricans. Eight dachshunds with primary acanthosis
nigricans showed improvement after 60 days of vitamin E therapy (200mg twice daily) (Scott and Walton, 1985). All dogs responded with gradual elimination of pruritus, inflammation, lichenification, greasiness,and odor.

CATS
Steatitis (yellow fat disease) results when sources of highly unsaturated fatty acids (e.g., tuna fish oil, cod liver oil, and unrefined herringoil) have been fed in the absence of adequate supplemental vitamin E.
The condition was known as yellow fat after the peroxidized ceroid in
adipose tissue. It was characterized by extreme hyperesthesia, fever, anda marked rise in leucocytes, primarily as neutrophils and eosinophils.
Anorexia, weight loss, listlessness or overt neurological dysfunction andmuscle spasms, harsh hair coat, and palpable lumps in the subcutaneous
fat were characteristic. Additional signs were focal interstitial myocarditis and, rarely, muscle fiber degeneration, focal myositis of the skeletal
muscle, and periportal mononuclear infiltration in the liver (Gershoffand Norkin, 1962). Affected cats are often lethargic and febrile andexhibit pain on gentle palpation. Subcutaneous and abdominal fat may
feel firm or lumpy, and draining tracts may develop. Microscopically,
the fat shows focal neutrophilic infiltration with some mononuclear
cells. Acid-fast ceroid pigment is present as globules and as peripheral
rings in the fat-cell vacuoles (NRC, 1986). The white blood cell count isusually elevated (24,000 to 70,000 μg/ml), primarily as a result of neutrophilia, and reflects the degree of fat necrosis.
Vitamin E deficiency in cats became a significant clinical problem
when fish products were first used in commercial cat foods that contained inadequate levels of tocopherol (Cordy, 1954; Munson et al.,1958). Later cases of the disease occurred in cats that were fed diets con200

V I TA M I N E

sisting wholly or largely of canned red tuna or fish scraps. Red tuna
packed in oil contains high levels of PUFA and low levels of vitamin E.
The addition of large amounts of fish products to a cat’s diet appears tobe the primary cause of this disease.
Stephan and Hayes (1978) induced severe vitamin E deficiency withhemolytic anemia and steatitis by feeding a diet containing 15%
stripped safflower oil without α-tocopherol. Clinical signs of deficiency
were prevented by supplementing with α-tocopheryl acetate at 100 IU
per kg.

FISH
Channel catfish fed a vitamin E-deficient diet containing oxidized
menhaden oil exhibited reduced growth, muscular dystrophy, fatty livers, anemia, exudative diathesis, and depigmentation in 16 weeks.
Sekoke disease is a condition in common carp characterized by a marked
loss of flesh. The disease in carp results from feeding oxidized silk worm
pupae and is completely prevented with supplemental vitamin E.
Vitamin E deficiency signs have been described for chinook salmon,
Atlantic salmon, channel catfish, common carp, rainbow trout, and yellowtail (NRC, 1993). The signs of vitamin E deficiency in various fishes are similar and include muscular dystrophy, involving atrophy andnecrosis of white muscle fibers; edema of heart, muscle, and other tissues due to increased capillary permeability, which allows exudates toescape and accumulate (these are often green as a result of hemoglobin
breakdown); anemia and impaired erythropoiesis; depigmentation; andceroid pigment in the liver. The incidence and severity of these deficiency signs were shown to be enhanced when diets deficient in both vitamin
E and Se were fed to Atlantic salmon (Poston et al., 1976), rainbow
trout (Bell et al., 1985), and channel catfish (Gatlin et al., 1986).

FOXES AND MINK
Diets containing rancid fats or high in unsaturated fat cause “yellow
fat” disease (steatitis) in foxes and mink as well as in cats. The most frequent clinical sign in animals with uncomplicated vitamin E deficiency
was sudden death due to minor stress (NRC, 1982a). Selenium had
some, but not complete, vitamin E-sparing effect.
Signs of steatitis in both foxes and mink are most evident among
fast-growing male pups. Acute and subacute cases frequently occur. In
fur-bearing animals, anemia is often found in chronic cases of yellow fat
disease. A pronounced fragility of the red blood cells and an increased
number of leukocytes and thrombocytes are frequent. In the post201

CHAPTER FOUR

mortem examination, the musculature often appears light and musty,
and fatty degeneration is usually discovered in the parenchymatousorgans and in the musculature. The skin is of poor quality and loses haireasily, both during and after tanning (Helgebostad and Ender, 1973).

LABORATORY ANIMALSIntrauterine death and resorption of fetuses in rats was the first evidence that led to the discovery of vitamin E. Other signs of vitamin E
deficiency in rats are increased red blood cell hemolysis, skeletal muscle
degeneration, accumulation of yellow pigment in smooth muscles; irreversible degeneration of the seminiferous epithelium of the testis in themale, which occurs by age 40 to 50 days; humped-back condition; roughcoat, skin ulcers; neural lesions; and impaired learning ability (Sarter
and Van Der Linde, 1987; NRC, 1995).
Pappenheimer (1942) reported muscular dystrophy and hyaline
degeneration in vitamin E-deficient mice but at a lower incidence than
was observed in rats. Spermatogenesis remained active in vitamin E-deficient mice for up to 439 days (Pappenheimer, 1942). Feeding a vitamin
E-deficient diet increased pathology in hearts of mice infected with avirus (Beck et al., 1994), indicating an immune response for vitamin E
in mice.
Diet-induced muscular dystrophy was produced in the guinea pig
when 5 to 20 g of cod liver oil per kilogram was included in the diet.
The research of Shimotori et al. (1940) related vitamin E deficiency tomuscular dystrophy. Vitamin E-deficient guinea pigs lost weight withdegeneration of skeletal muscle. In males, testes atrophied and developed degenerative changes in the seminiferous tubules, with clumping orcomplete disappearance of spermatozoa and spermatids. Fetal malformations, resorption, and death occurred in pregnant females (NRC,1995).

NONHUMAN PRIMATES
When deprived of vitamin E for prolonged periods, usually a year or
more, rhesus and cebus monkeys developed a characteristic anemia andmuscular dystrophy (NRC, 1978). Anemia resulted in the production of
defective erythrocytes, with muscular dystrophy closely resembling vitamin E deficiency in other species.

RABBITS
Muscular dystrophy in rabbits is caused primarily by vitamin E defi202

V I TA M I N E

ciency. The rabbit appears to be unusual in that Se has neither a protective nor sparing effect on muscular dystrophy that is preventable by vitamin E (Hove et al., 1958). Signs of this syndrome include degenerationof the skeletal and cardiac muscles, paralysis, and fatty liver (Bragdon
and Levine, 1949). Ringer and Abrams (1970) encountered widespreadsigns of vitamin E deficiency in a commercial herd of rabbits fed a natural diet that provided 16.7 ppm of vitamin E. Tissues from rabbits fed
ω-3 fatty acids rather than ω-6 and ω-9 were more susceptible to oxidation, with α-tocopherol lowering lipid oxidation (Lopez-Bote et al.,1997).

EXOTIC ANIMALS
Numerous cases of vitamin E deficiency have been diagnosed in zoo
and wildlife species (Dierenfeld and Traber, 1992). Increased longevityof species in zoos, possibly limited genetic variability, and various stressfactors contribute to the increasing number of clinical reports; however,dietary deficiency of vitamin E is likely the single most important determinant underlying current observations (Dierenfeld, 1994). Cervids(deer) and bovids (antelope) with vitamin E deficiency display cardiac
and skeletal muscle dystrophy (Liu et al., 1985; Dierenfeld, 1989).
Zebras suffer deterioration of the myelin sheath and spinal cord withvitamin E deficiency, clinically displayed as asymmetric weakness andataxia (Liu et al., 1983). Both the elephant and rhinoceros appear tohave limited absorption of dietary vitamin E, perhaps due to minimal
dietary lipid levels in zoo feeds. Elephants with deficiency have heartlesions similar to those of swine with microangiopathy (Dierenfeld andDolensek, 1988), whereas rhinos display skeletal and cardiac muscle
myopathies. Among all exotic birds examined, species with carnivorousor granivorous feeding habits are most severely affected by vitamin E
deficiency. Deficiency signs include decreased hatchability through pipping and cardiac muscle degeneration (Liu et al., 1985; Dierenfeld,
1989).

HumansAt one time, it was believed that in the United States and otherdeveloped countries, vitamin E intake of most adult human populations
was considered adequate for maintenance, growth, and reproduction innormal individuals. This view is radically changing as supplemental vitamin E has shown benefits for improving the immune system; for prevention of cancer, thrombosis and cardiovascular diseases, cataracts,
arthritis, and Parkinson’s disease; and as an antidote to toxic substances203

CHAPTER FOUR

and other pathological processes and diseases (Tengerdy, 1989; Padh,
1991; Consumer Report on Health, 1993; Langseth, 1995a,b; Chew,
1995). The various signs of vitamin E deficiency in humans often are
believed to be manifestations of membrane dysfunction resulting fromthe oxidative degradation of polyunsaturated membrane phospholipids
and/or the disruption of other critical cellular processes. Free radicals
can be extremely damaging to biological systems (Padh, 1991).
Antioxidants stabilize these highly reactive free radicals, thereby maintaining the structural and functional integrity of cells (Chew, 1995).
Therefore, antioxidants are very important to immune defense andhealth of both humans and animals. Tissue defense mechanisms against
free-radical damage include vitamin E, vitamin C, and β-carotene as the
major vitamin antioxidant sources.
New findings on vitamin E and cardiovascular diseases were summarized by Langseth (1995a) as follows:
1. The use of vitamin supplements—especially vitamin E—was associated with a significantly reduced risk of coronary disease in Canadian men.
2. Low vitamin E intake was associated with elevated coronary mortality in Finnish men and women.
3. Vitamin E enhanced the benefits of aspirin in reducing the incidenceof stroke in high-risk patients.
4. Vitamin E supplements slowed progression of coronary artery
lesions, both in men taking cholesterol-lowering drugs and in those taking
inactive placebos.
5. Vitamin E supplements lowered the risk of recurrence of coronary
blockage in patients who had been treated for previous blockages.
6. Two studies of diabetics (a group at high risk for cardiovascular disease) showed beneficial changes in lipid peroxidation and other biochemicalfactors after vitamin E supplementation.

Jialal and Grundy (1993) reported that combined supplementation
with vitamin C, β-carotene, and α-tocopherol is not superior to highdose α-tocopherol alone in inhibiting low-density lipoprotein oxidation.
Hence, α-tocopherol therapy should be favored in future coronary prevention trials involving antioxidants.
Cancer-prevention findings related to vitamin E have been summarized (Consumer Report on Health, 1993; Langseth, 1995a). In addition
to possibly helping prevent cancer by keeping cells from becoming oxidized and turning cancerous, vitamin E might fight the disease indirectly by boosting the immune system. People who had taken vitamin E
204V I TA M I N E

supplements for at least 6 months were half as likely to develop cancersof the mouth and throat. The evidence suggests that people who consume the most vitamin E may have a slightly lower risk of cancers of thecolon, rectum, esophagus, and lung. One study reported a 34% decreasein prostate cancer and a 16% decrease in colorectal cancer. Preliminary
research has hinted that certain forms of vitamin E applied to the skinmight help protect against skin cancer caused by sun exposure.
In animal species, there is considerable evidence of beneficial effectsof vitamin E on the immune response. In humans, however, there are
only limited data on the relation between vitamin E status and theimmune system. In a French study of healthy elderly subjects, the levelof plasma vitamin E was inversely correlated with the number of infections the subjects experienced in the 3 years previous to the study(Chavance et al., 1989). In a double-blind, placebo-controlled trial,
healthy older adults (60 years or older) improved some in vivo and invitro parameters of immune function as a result of receiving supplemental vitamin E (Meydani, 1995).
A myriad of neurological abnormalities can be found in the presenceof vitamin E deficiency. Cavalier and Gambetti (1981) reported dystrophic axons in the nucleus gracilis and demyelination of the fasciculus
gracilis in patients with cystic fibrosis. In a review, Sokol (1989) reported that vitamin E deficiency in humans results in loss of deep tendon
reflexes, truncal and limb ataxia, reduced perception of vibration andposition, ophthalmoplegia, muscle weakness, dysarthria, and scoliosis.
Additional human diseases that are alleviated by vitamin E include
thrombophlebitis and intermittent claudication, both of which involve
blood flow, particularly in the extremities of elderly persons (Haeger,1974). Low levels of vitamin E also have been found in the plasma ofmany persons with absorptive defects (Sitrin et al., 1987). Patients suffering from sprue, cystic fibrosis of the pancreas with accompanying
steatorrhea, or any other disease that causes malabsorption of fat also
show a marked decrease in plasma tocopherol levels.
Plasma vitamin E concentration in the newborn infant is about onethird that of adults, with that in the low-birth-weight (LBW) infant evenlower. This is primarily a reflection of lower concentration of blood
lipids in newborn infants that is due to inefficient placental transfer ofthe vitamin. Serum vitamin E levels increase during pregnancy, while
fetal levels remain low, at about 25% of the maternal level. In the UnitedStates, edema and anemia attributed to vitamin E deficiency have beenreported in LBW infants fed low-vitamin E commercial formulas madewith polyunsaturated fat (Oski and Barness, 1967). Premature infants
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CHAPTER FOUR

fed these formulas with inadequate vitamin E develop hemolytic anemia,
resulting in shortened life span of red blood cells.
Numerous researchers have reviewed the effects of vitamin E on
human diseases and presented two extreme points of view: (1) One
group claims that vitamin E is a cure for almost every known disease;the other group holds that vitamin E has not been proved scientificallyto have any of the effects attributed to it. (2) The second viewpoint—that vitamin E is of little value—is no longer valid, as many studies haveshown the positive benefits of vitamin E supplementation for the prevention and cure of many disease conditions.

Assessment of Status
Confirmation of a low vitamin E and/or Se status in animals is
obtained when specific deficiency diseases associated with lack of thesenutrients are present. Likewise, gross lesions and histopathologicalexaminations provide definite evidence of vitamin E and/or Se deficiency.
Muscular damage as a result of vitamin E and/or Se deficiency causes leakage of intercellular contents into the blood. Thus, elevated levelsof selected enzymes (above the normal concentrations for particularspecies) serve as diagnostic aids in detecting tissue degeneration. Serum
enzyme concentrations used to follow incidence of nutritional muscular
dystrophy include serum glutamic-oxaloacetic-transaminase (SGOT),
aspartate amino transferase (ASPAT), lactic dehydrogenase (LDH), creatine phosphokinase (CPK), and malate dehydrogenase (MDH). A distinction regarding type of tissue degeneration can sometimes be made.
For example, in swine, SGOT and ornithine carbamyl transferase (OCT)
are useful indicators of muscular dystrophy and liver necrosis, respectively (Wretlind et al., 1959). Enzyme tests are very sensitive, and an elevation of enzyme activity in serum is usually discovered before any
pathological changes or clinical signs appear (Tollersrud, 1973).
In addition to serum enzyme determinations, other laboratory tests
devised to assist in diagnosis of vitamin E and Se deficiency include (1)vitamin E and Se analyses of feeds, blood, and tissues and (2) electrocardiogram changes that reflect heart muscle injury. Low tissue concentrations of glutathione peroxidase, an enzyme that contains Se, is a relatively good status indicator of this element. However, an in vitro
hemolysis test is an indicator of vitamin E but not Se status.
Nutritional status with respect to vitamin E is commonly estimatedfrom plasma (or serum) concentration. There is a relatively high correlation between plasma and liver levels of α-tocopherol (also between
206

V I TA M I N E

amount of dietary α-tocopherol administered and plasma levels). Thishas been observed in rats, chicks, pigs, lambs, and calves within rather
wide ranges of intake. There is a much higher correlation between bloodand liver concentrations for vitamin E than for vitamin A. Plasma tocopherol concentrations of 0.5 to 1 μg/ml are considered low in most animal species, with less than 0.5 μg/ml generally considered a vitamin E
deficiency. Thus, plasma α-tocopherol concentrations can be used forassay purposes without the necessity of liver biopsy or animal slaughter(Ullrey, 1981; Njeru et al., 1994b).
In normal adult human populations of the United States, the rangeof total plasma tocopherols is 0.5 to 1.2 μg/ml, with values for α-tocopherol 10 to 15% lower (Bieri and Prival, 1965). It is generally accepted that a plasma level of total tocopherols below 0.5 μg/ml is undesirable, although it has not been shown that lower concentrations inadults, unless they continue a year or longer, are associated with inadequate tissue concentrations. Plasma tocopherol concentrations are highly correlated with total lipid, with less than 0.6 to 0.8 μg/g lipid considered deficient in humans (Machlin, 1991).
For a number of disease conditions, it is important to know the status of Se as well as vitamin E. For many species, Se concentrations inliver, renal cortex, and blood (as well as other tissues) each adequately
portray Se status. For example, swine hepatic, renal cortex, and blood
Se concentrations of 0.25, 2.5, and 0.1 ppm (dry basis), respectively,
were determined to be critical levels below which clinical illness, death,or lesions of vitamin E-Se deficiency could be expected (McDowell et al.,1977). Serum or plasma Se is considered a good status indicator, withless than 0.03 ppm being critical for cattle (McDowell, 1985).

SUPPLEMENTATION
Methods of providing supplemental vitamin E are (1) as part of aconcentrate or liquid supplement, (2) included with a free-choice mineral mixture, (3) as an injectable product, and (4) in drinking water preparations.
To offset losses of vitamin E activity in feedstuffs, diets should be
adequately fortified with vitamin E. The principal commercially available forms of vitamin E used in food, feed, and pharmaceutical industries are acetate and hydrogen succinate esters of RRR α-tocopherol andthe acetate ester of all-rac-α-tocopherol (Table 4.9). During commercial
synthesis of dl-α-tocopherol, it is esterified to acetate to stabilize it, withthe ester extremely resistant to oxidation. Thus, dl-α-tocopherol acetate207

CHAPTER FOUR
■ Table 4.9

Forms of α-Tocopherol Commercially Available

Form

IU/mg

dl-α-Tocopheryl acetate (all-rac)dl-α-Tocopherol (all-rac)d-α-Tocopheryl acetate (RRR)
d-α-Tocopherol (RRR)
dl-α-Tocopheryl acid succinate (all-rac)d-α-Tocopheryl acid succinate (RRR)

1.00
1.10
1.36
1.49
0.89
1.21

does not act as an antioxidant in the feed and has antioxidant activity
only after it is hydrolyzed in the intestine and free dl-α-tocopherol isreleased and absorbed.
The acetate forms of α-tocopherol are commercially available fromtwo basic sources (Hoffmann-La Roche, 1972): (1) d-α-tocopheryl
acetate is made by extraction of natural tocopherols from by-productsof vegetable oil refining, molecular distillation to obtain the alpha form,
and then acetylation to form the acetate ester, and (2) dl-α-tocopheryl
acetate is made by complete chemical synthesis, producing a racemic
mixture of equal parts of d and l isomers. The d and l forms differ onlyin spatial placement of the isoprenoid side chain.
Commercially, the dl- and d-α-tocopheryl acetates are available inpurified form or in various dilutions, including (Hoffmann-La Roche,1972) (1) a highly concentrated oily form for further processing; (2)emulsions incorporated in powders for use in dry premixes or water-dispersible preparations; (3) beadlets or powders consisting of the tocopheryl acetate incorporated in oil, or in emulsifiable form mixed withgelatin and sugar, gum acacia, soy grits, or dextrin as carriers (suchbeadlets or powders may be further diluted to lower potencies in a feed
ingredient or water-soluble material and are primarily for use in feed);and (4) adsorbates of the oily tocopheryl acetate on selected carriers, infree-flowing “dry” powder, meal, or granules. This type is for use in
feeds only. Vitamin E as the acetate form is highly stable in vitamin premixes, with 98% retention after 6 months, but in the alcohol form iscompletely destroyed during that time (Coelho, 1991).
Even though the ester is more stable than the natural free or alcohol
form, it is desirable to further stabilize it by a gelatin coating or adsorption technique that reduces it to a beadlet, granule, or powder form tobe added more easily and uniformly to animal feeds (Bauernfeind,1969). Both types of products are quite stable. Stabilized dl-α-tocopheryl acetate gelatin beadlets have been blended in mash feed that also208

V I TA M I N E

has been pelleted and stored with satisfactory stability results.
Injectable vitamin E preparations are also available, which containfree d- or dl-α-tocopherol or their esters. Liquid emulsions of appropriate types are used in drinking water and liquid feeds, in regular feed, andfor injection. In this form, α-tocopherol is more efficiently absorbed
from intramuscular injection sites than a water-miscible preparation
containing α-tocopheryl acetate or either form dissolved in an oily base
(Machlin, 1991). Products containing vitamin E and Se are often given
intramuscularly to animals exhibiting clinical signs of muscular degeneration. Response to treatment of this condition is extremely variable
depending on degree of muscular degeneration. However, intramuscular
injection of α-tocopherol and Se to young ruminants usually provides a
rapid means of alleviating deficiencies of these nutrients (McDowell,1992). For cattle, animals that are down are unlikely to survive, while
animals showing moderate weakness of the rear limbs or slight bucklingof fetlocks may respond. Total recovery may require several days to 1
month (Mason et al., 1985). However, affected animals are particularly
responsive to treatment, generally gaining ability to walk unassisted 3 to
5 days following vitamin E-Se therapy.
Some cattle ranchers make it a practice to inject newborn calves
intramuscularly with a combination of vitamin E and Se. For dystrophic
lambs, an oral therapeutic dose of 500 mg of dl-α-tocopherol followedby 100 mg on alternate days until recovery was successful (Rumsey,1975). Reddy et al. (1987a) concluded that supplementation of conventional dairy calf diets with 125 to 250 IU vitamin E per animal daily
increases performance compared to controls. Stuart (1987) recommended that weaned calves with transit shrink receive 400 IU of vitamin E
daily during arrival to a feedlot finishing program, while those with minimal shrink receive 100 to 200 IU. The recommended amount for yearling cattle was 200 to 400 IU vitamin E, depending on previous dietaryhistory in relation to vitamin E and Se. Most preventive preparations for
WMD in ruminants contain a combination of both vitamin E and Se. As
a preventive measure, they should be administered to pregnant cows during the second trimester and again 30 days prior to calving.
Supplementing vitamin E in well-balanced diets has been shown toincrease humoral immunity for ruminants (Hoffmann-La Roche, 1994)
and monogastric species (Langweiler et al., 1983; Wuryastuti et al.,1993). These results suggest that the criteria for establishing requirements based on overt deficiencies or growth do not consider optimal
health (Hoffmann-La Roche, 1991). The effects of oral vitamin E supplementation in young calves was evaluated by Cipriano et al. (1982).
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CHAPTER FOUR

Calves were fed skimmed colostrum and supplemented with either 0 or
1,000 mg dl-α-tocopheryl acetate for 6 weeks in a vitamin E-deficientdiet. Conventionally managed calves were included as positive controls.
Vitamin E-supplemented calves had greater plasma α-tocopherol concentrations as well as mean lymphocyte blastogenesis response to phytohemagglutinin (PTH) expressed as mean lymphocyte stimulation
indices (LSIs) at 6 weeks. These authors suggested that the enhancing
effect of vitamin E on immune response of cattle could have been partially masked in this study by feeding of diets high in emulsified fats.
Vitamin E administration to calves enhanced immune response andweight gain, while enzymes of muscle origin (e.g., creatine kinase andserum glutamic oxaloacetic transaminase) and plasma cortisol concentration were decreased (Reddy et al., 1987b). Vitamin E also positively
influenced neutrophil-mediated antibody-dependent cellular cytotoxicity and phagocytosis as well as lymphocyte stimulation in calves fed milk
replacer (Pruiett et al., 1989).
In a series of 28-day feedlot receiving trials, Lee et al. (1985)
observed an improvement in early performance of newly arrived growing cattle supplemented with 450 IU vitamin E (as dl-α-tocopheryl
acetate) per head per day that were stressed by long-distance shipment
and changes from green forages to high-grain feedlot diets. Depressionof circulating cortisol concentrations may explain the improved gain
and feed efficiency in this trial.
Gill et al. (1986) supplemented newly received feedlot cattle with1,600 IU of vitamin E (as dl-α-tocopheryl acetate) per head per day forthe first 21 days and 800 IU for the remaining 7 days of a 28-day trial.
Average daily gain and gain-feed ratios were improved by 23.2 and28.6%, respectively, for vitamin E-supplemented stressed cattle (Table
4.10). The number of sick pen days per head was reduced by 15.6%,
and morbidity was reduced by 13.4% with vitamin E supplementation.
The growth response to vitamin E could be related to the fact thatyoung, rapidly growing animals are in a metabolically demanding state
resulting from overall tissue growth, which has a high energy demand.
Vitamin E is an integral part of this response via its ability to quench free
radicals, which are generated during the course of metabolism.
Supplemental levels of vitamin E higher than those recommended
for dairy cattle by the NRC (1989a) have been beneficial in the controlof mastitis. Smith and Conrad (1987) reported that intramammary
infection was reduced 42.2% in vitamin E- and Se-supplemented versus
unsupplemented controls. The duration of all intramammary infections
in lactation was reduced 40 to 50% in supplemented heifers. Weiss et al.
210

V I TA M I N E■ Table 4.10

Effects of Vitamin E Supplementation on Performance,
Morbidity, and Mortality of Stressed Cattle

Item

Control

Vitamin Ea

No. of head

252

250

Average daily gain (kg)

0.43b

0.53c

Feed conversion

18.56

15.06

No. of sick days

3.2

2.7

Morbidity (%)

43.2

37.5

Mortality (%)

1.8

l.6

Source: From Gill et al. (1986).
a1,600 IU of vitamin E per head per day for first 21 days and 800 IU for
the last 7 days.
b,cMeans with different superscripts differ (P < 0.01).

(1990) reported that clinical mastitis was negatively related to plasma Se
concentration and concentration of vitamin E in the diet.
Many new intramammary infections (IMIs) occur in the 2 weeks
before and after calving. Deficiencies of either vitamin E or Se have been
associated with increased incidence and severity of IMI, increased clinical mastitis cases, and higher somatic cell counts in individual cows andbulk tank milk. Somatic cell counts are a primary indicator of mastitis
and milk quality in dairy herds. The polymorphonuclear neutrophil(PMN) is a major defense mechanism against infection in the bovine
mammary gland. A known consequence of vitamin E and Se deficiency
is impaired PMN activity, and postpartum vitamin E deficiency is frequently observed in dairy cows. Dietary supplementation with vitamin
E and Se results in a more rapid PMN influx into milk following intramammary bacterial challenge and increased intracellular kill of ingested
bacteria by PMN. In one study, subcutaneous injections of vitamin E
approximately 10 and 5 days before calving successfully elevated PMN
α-tocopherol concentrations during the periparturient period and negated the suppressed intracellular kill of bacteria by PMN that commonly
is observed around calving (Smith et al., 1997).
Diets of multiparous dairy cows were supplemented with either 0 or
1,000 IU of vitamin E (as dl-α-tocopheryl acetate) during the dry period(Smith et al., 1984). Cows were additionally administered Se at the rateof 0 or 0.1 mg/kg body weight via intramuscular injection 21 days
prepartum. No vitamin E or Se was supplemented during lactation.
Incidence of new clinical cases of mastitis was reduced by 37% in both211

CHAPTER FOUR

groups receiving vitamin E compared to controls. The reduction in clinical mastitis was only 12% when cows were injected with Se but not
supplemented with dietary vitamin E. These authors also reported thatclinical cases in the vitamin E-supplemented, Se-injected cows were consistently of shorter duration than those occurring in all other groups.
Erskine et al. (1989) investigated specific effects of Se status of dairy cattle on the induction of mastitis by E. coli. Bacterial concentrations weresignificantly higher in Se-deficient than in Se-adequate cows, and Se supplementation reduced both severity and duration of clinical mastitis.
Plasma concentrations of α-tocopherol decreased at calving forcows fed dietary treatments with low or intermediate concentrations ofvitamin E, but not for cows fed the high vitamin E treatment (Weiss etal., 1997). High dietary vitamin E increased concentrations of α-tocopherol in blood neutrophils at parturition. The high vitamin E treatment
was 1,000 IU/day of vitamin E during the first 46 days of the dry period, 4,000 IU/day during the last 14 days of the dry period, and 2,000
IU/day during lactation.
The percentage of quarters with new infections at calving was notdifferent (32.0%) between cows receiving treatments that contained low
and intermediate concentrations of vitamin E but was reduced (11.8%)
in cows receiving the high vitamin E treatment. Clinical mastitis affected 25.0, 16.7, and 2.6% of quarters during the first 7 days of lactation
for cows receiving the low, intermediate, and high vitamin E treatments,
respectively. Cows with plasma concentrations of α-tocopherol less than
3.0 μg/ml at calving were 9.4 times more likely to have clinical mastitis
during the first 7 days of lactation than were cows with concentrations
greater than 3.0 μg/ml (Weiss et al., 1997).
Many attempts have been made to control lipid oxidation in meats
through the use of antioxidants. One such approach is through dietary
supplementation of vitamin E, which functions as a lipid-soluble antioxidant in cell membranes (Linder, 1985), thus protecting phospholipids
and even cholesterol against oxidation. Increased dietary levels of vitamin E result in higher tissue α-tocopherol concentrations and greater
stability of these tissues toward lipid oxidation and thus, for example,
increased shelf-life of pork (Soler-Velásquez et al., 1998; Corino et al.,
1999; Lauridsen et al., 1999). Buckley and Connolly (1980) reduced therate of rancidity development in frozen pork by including vitamin E in
the feed (80 mg per day per animal) for 7 days before slaughter of thepigs. Likewise, meat from turkeys raised on vitamin E-supplemented
diets was more resistant to rancidity development than meat from birds
receiving control diets (Uebersax et al., 1978; Bartov et al., 1983).
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Dramatic effects of vitamin E supplementation (500 IU per headdaily) to finishing steers on the stability of beef color have been observed(Faustman et al., 1989a). Loin steaks of control steers discolored 2 to 3
days sooner than those of steers supplemented with vitamin E.
Supplemental dietary vitamin E extended the color shelf-life of loinsteaks from 3.7 to 6.3 days. This was most likely due to the increased αtocopherol content of the loin tissue of the supplemented animals, which
was approximately fourfold greater than that of the controls (Faustman
et al., 1989a). Color is an extremely critical component of fresh red meat
appearance and greatly influences the customer’s perception of meat
quality. Steaks from cattle supplemented with vitamin E were preferredover those from controls by 91% of Japanese survey participants (N =10,941), and 58% of all participants identified muscle color as the mostimportant factor in selecting beef products (Sanders et al., 1997).
In a subsequent report, Faustman et al. (1989b) observed that vitamin E stabilized the pigments and lipids of meat from the supplemented
steers. Perhaps the vitamin E-supplemented steers were able to incorporate a greater amount of vitamin E into cellular membranes, where it canperform its antioxidant function. The effects of vitamin E as an in vivo
lipid stabilizer and its effect on flavor and storage properties of various
meats have been reviewed. Supplementing cattle with vitamin E resulted
in steaks that exhibited superior lean color, less surface discoloration,
more desirable overall appearance, and less lipid oxidation during retaildisplay than control steaks (Sanders et al., 1997). Vitamin E also plays
a role in controlling the color of veal calf meat. Combined feeding ofmonosodium phosphate and 100 IU of vitamin E per calf daily produced
a light-colored veal without making calves anemic (Agboola et al.,1990).
Feeding supplemental vitamin E at levels of 1,000 to 2,000 mg ofnaturally occurring mixed tocopherols per cow per day increased thevitamin E content of milk and its stability against oxidized flavor(Krukovsky and Loosli, 1952; Neilsen et al., 1953). The vitamin E content of milk from cows fed stored feeds was lower than that of milk fromcows on pasture, and their milk was more susceptible to development ofoxidized flavor (Krukovsky et al., 1950).
Feeding supplemental vitamin E as dl-α-tocopheryl acetate, providing an equivalent of 500 mg of dl-α-tocopherol per cow per day,increased the vitamin E content and oxidative stability of milk (Dunkley
et al., 1967). Nicholson et al. (1991) suggested that adequate Se
improves the transfer of dietary tocopherol to milk.
The need for supplementation of vitamin E is dependent on the213CHAPTER FOUR

requirement of individual species, conditions of production, and available vitamin E in food or feed sources (see Natural Sources). The primary factors that influence the need for supplementation include (1)vitamin E- and/or Se-deficient concentrates and roughages; (2) excessively dry ranges or pastures for grazing livestock; (3) confinement feeding where vitamin E-rich forages are not included, or only forages ofpoor quality are provided; (4) diets that contain predominantly non–αtocopherols and thereby are less biologically active; (5) diets that include
ingredients that increase vitamin E requirements (e.g., unsaturated fats,waters high in nitrates); (6) harvesting, drying, or storage conditions offeeds that result in destruction of vitamin E and/or Se (see Natural
Sources); (7) accelerated rates of gain, production, and feed efficiency
that increase metabolic demands for vitamin E; and (8) intensified production that also indirectly increases vitamin E needs of animals by elevating stress, which often increases susceptibility to various diseases(McDowell and Williams, 1991; McDowell, 1992). After stress, livestock may have reductions in α-tocopherol concentrations in certain tissues. Supplemental vitamin E may be required after stress to restore αtocopherol in tissues (Nockels et al., 1996). Based on health data, Weiss(1998) suggested the vitamin E requirement should be increased at least
500% and perhaps as much as 700% above NRC recommendations fordry cows and lactating cows.
Vitamin E-Se deficiencies are found in specific world regions and arecharacterized by low concentrations of feedstuffs. Regions that rely on
concentrate importation from these areas deficient in vitamin E and/or
Se (e.g., midwestern United States) likewise must provide these nutrients
to livestock. Adverse conditions, such as poor weather (drought andearly frost), molds, and insect infestation, will reduce the vitamin E
value of feedstuffs. The vitamin E activity in blighted corn was 59%
lower than that in sound corn, and the vitamin E activity in lightweight
corn averaged 21% below that in sound corn (Hoffmann-LaRoche,1994). Feed spoilage will also promote vitamin E-Se deficiencies; therefore, to prevent loss of vitamin E in diets, the producer should use fresh
feed at all times because the vitamin is rapidly destroyed under hot,humid conditions. Also, the producer should use an antioxidant in thediet to prevent the destruction of the vitamin E (as well as other vitamins, e.g., vitamin A). Losses during storage increase as the durationand temperature of storage increase. Rate of oxidation of natural tocopherols is increased in diets containing increased levels of copper, iron,zinc, or manganese (Dove and Ewan, 1991).
In pet foods, the most important feed ingredient that will dictate the214V I TA M I N E

need for vitamin E supplementation is fat source and the stability of the
fat source. Large amounts of PUFA can quickly precipitate vitamin E
deficiency. Fats from oilseeds are normally not highly detrimental
because the oils, in addition to containing high amounts of predominantly linoleic acid (18:2), likewise are sources of vitamin E. Fish oils,
however, contain little vitamin E and are highly destructive to vitamin E
stores and greatly increase the requirements of the vitamin. This can bea problem for cats, since their diets can be very high in fish and fish oils.
The long-chain, highly unsaturated fatty acids in fish oil can quickly
deplete the cat’s body stores of vitamin E. A diet that contains high levels of fish oil may cause a threefold to fourfold increase in a cat’s dailyrequirement for vitamin E. Pets should never be fed rancid fats as they
will quickly bring about destruction of fat-soluble vitamins.
The ability of vitamin E to affect growth, health, and reproductionof animals is documented. A vitamin E supplementation program utilizing both parenteral and oral administration is often suggested, particularly when fresh green pasture is lacking. Mahan (1991) assessed theinfluence of low supplemental vitamin E (< 16 IU/kg) to sows and offspring in three parities. Smaller litter size, sow agalactia, and pig mortality during the first week after birth resulted from inadequate supplemental vitamin E to breeding sows.
Exercise has an influence on vitamin E requirements and needed
supplementation (Davies et al., 1982; Valberg et al., 1993). Dietary levels of vitamin E greater than the 80 IU/kg dry matter, and potentiallyapproaching 300 IU/kg dry matter, are required to maintain blood andmuscle vitamin E concentrations in horses undergoing exercise conditioning. The level of vitamin E recommended by the NRC for working
horses, 80 IU/kg dry matter, will not maintain serum vitamin E levels.
In humans, vitamin E supplements may protect overworked muscles
during exercise. In a pair of studies, sedentary men took either a placebo or 530 mg of vitamin E every day for 7 weeks. They then performed
a vigorous 45-minute workout on the treadmill. After the workout,
blood samples from the men who took vitamin E contained smaller
amounts of chemicals that promote inflammation. That effect, theresearchers speculated, should limit both soreness and injury after strenuous exercise (Consumer Report on Health, 1993).
Good experimental evidence indicates a need for supplemental vitamin E in the diets of pregnant and lactating women, newborn infants(particularly premature infants), and older persons suffering from circulatory disturbances and intermittent claudication. Chronic diarrhea isalso associated with reduced absorption of vitamin E (Liuzzi et al.,215

CHAPTER FOUR

1998). Higher levels may be indicated for persons exposed to oxygenand environmental pollutants, such as ozone, nitrites, and heavy metals.
Feeding LBW infants entails problems somewhat different from those of
normal-weight infants. Because of their reduced absorption of fat, utilization of α-tocopherol is impaired; thus special effort is required toensure adequate intake. The American Academy of Pediatrics
Committee on Nutrition (1977) recommended that formulas for these
infants should provide 5 IU of water-soluble α-tocopherol daily.
There are human health benefits of consumption of higher levels ofvitamin E (Vitamin E Research and Information Service, 1993;
McDowell et al., 1996). Typical human vitamin E supplementation levels are 200 to 800 IU daily. The antioxidant nutrients, vitamin E, vitamin C, and β-carotene, have become the focus for their protective rolein disease prevention, particularly cardiovascular disease (atherosclerosis) and cancer. Vitamin E also has a beneficial effect in preventing eye
disorders, skin diseases, ulcers, and intermittent claudication (inadequate blood flow). Vitamin E may reduce atherosclerosis by preventing
low-density lipoproteins from oxidizing and causing arterial injury.
Antioxidants influence the virulence of virus infections. In a healthy
state, a virus would be benign or less virulent. Vitamin E deficiency may
allow viruses to cause disease and may have important implications forenterovirus-induced inflammatory heart disease in adult human populations (Beck et al., 1994).

TOXICITY
Compared with vitamin A and vitamin D, both acute and chronic
studies with animals have shown that vitamin E is relatively nontoxic
but not entirely devoid of undesirable effects. Hypervitaminosis E studies in rats, chicks, and humans indicate maximum tolerable levels in therange of 1,000 to 2,000 IU/kg of diet (NRC, 1987).
Massive doses of vitamin E (5,000 mg of dl-α-tocopherol per kilogram of diet) caused reduced packed-cell volumes in trout (NRC, 1993).
Administration of vitamin E to vitamin K-deficient rats, dogs, chicks,
and humans exacerbated the coagulation defect associated with vitamin
K deficiency (Corrigan, 1982). In chickens, the effects of vitamin E toxicity are depressed growth rate, reduced hematocrit, reticulocytosis,
increased prothrombin time (corrected by injecting vitamin K), andreduced calcium and phosphorus in dry, fat-free bone ash (NRC, 1987).
In humans, isolated and inconsistent reports have appeared on theadverse effects from high intakes of dl-α-tocopheryl acetate, but most216

V I TA M I N E

adults appear to tolerate these doses. Negative effects in human subjects
consuming up to 1,000 IU of vitamin E per day included headache,fatigue, nausea, muscular weakness, and double vision. Large doses of
α-tocopherol in anemic children suppress normal hematologicalresponse to parenteral iron administration (RDA, 1989).

■ REFERENCES
Adams, C.R. (1973). Effect of Processing on the Nutritional Value of Feeds, p.
142. National Adacemy of Sciences, Washington D.C.
Adamstone, F.B., Krider, J.L., and James, M.F. (1949). Ann. N.Y. Acad. Sci. 52,
260.
Agboola, H.A., Cahill, V.R., Conrad, H.R., Ockerman, H.W., Parker, C.F.,
Parrett, N.A., and Long, A.R. (1990). J. Anim. Sci. 68, 117.
American Academy of Pediatrics, Committee on Nutrition (1977). Pediatrics 60,
19.
Anderson, H.D., Elvehjem, C.A., and Gonce Jr., J.E. (1939). Proc. Soc. Exp.
Biol. Med. 42, 750.
Anderson, H.D., Elvehjem, C.A., and Gonce Jr., J.E. (1940). J. Nutr. 20, 433.
Anderson, P.H., Berrett, S., and Patterson, D.S. (1976). Vet. Rec. 99, 316.
Andrews, E.D., Hartley, W.J., and Grunt, A.B. (1968). N.Z. Vet. J. 16, 3.
Baalsrud, K.J., and Overnes, G. (1986). Eq. Vet. J. 18, 472.
Badwey, J.A., and Karnovsky, M.L. (1980). Ann. Rev. Biochem. 49, 695.
Barton, C.R., and Allen, W.M. (1973). Vet. Rec. 92, 288.
Bartov, I., Basker, D., and Angel, S. (1983). Poult. Sci. 62, 1224.
Bauernfeind, J.C. (1969). World Rev. Anim. Prod. 5, 20.
Bauernfeind, J.C. (1980). In Vitamin E: A Comprehensive Treatise, p. 99 (L.J.
Machlin, Ed.). Marcel Dekker, New York.
Beck, M.A. (1997). J. Nutr. 127, 966S.
Beck, M.A., Kolbeck, P.C., Kohr, L.H., Shi, Q., Morris, V.C., and Levander,
O.A. (1994). J. Nutr. 124, 345.
Bell, J.G., Cowey, C.B., Adron, J.W., and Shanks, A.M. (1985). Br. J. Nutr. 53,
149.
Bendich, A. (1987). In Proc. Roche Technical Symposium: The Role of Vitamins
on Animal Performance and Immune Response. RCD 7442. Hoffmann-La
Roche Inc., Nutley, New Jersey.
Bendich, A., Gabriel, E., and Machlin, L.J. (1986). J. Nutr. 116, 675.
Bieri, J.G., and Prival, E.L. (1965). Proc. Soc. Exp. Med. 120, 554.
Blaxter, K.L. (1962). Proc. Nutr. Soc. 21, 211.
Bostedt, H. (1980). Collegium Vet. 61, 45.
Bragdon, J.H., and Levine, D.H. (1949). Am. J. Pathol. 25, 265.
Brownlee, N.R., Huttner, J.J., and Panganamala, R.V. (1977). J. Lipid Res. 18,
635.
Buckley, D.J., and Connolly, J.F. (1980). J. Food Protect. 43, 265.
Burkholder, W.J., and Swecker, W.S. (1990). Semin. Vet. Med. Surg. (Small

217

CHAPTER FOUR

Anim.) 5(3), 154.
Cavalier, S.J., and Gambetti, P. (1981). Neurology 31, 714.
Charmley, E. Hidiroglou, N., Ochoa, L., McDowell, L.R., and Hidiroglou, M.
(1992). J. Dairy Sci. 75, 804.
Chavance, M., Herbeth, B., Fournier, C., Janot, C., and Vernhes, G. (1989). Eur.
J. Clin. Nutr. 43, 827.
Chew, B.P. (1995). J. Nutr. 125, 1804S.
Chow, C.K. (1979). Am. J. CLin. Nutr. 32, 1066.
Chung, Y.K., and Mahan, D.C. (1995). Korean J. Anim. Sci. 37, 616.
Cipriano, J.E., Morrill, J.L., and Anderson, N.V. (1982). J. Dairy Sci. 65, 2357.
Codner, E.C., and Thatcher, C.D. (1993). Compend. Cont. Educ. Small Anim.
15, 411.
Coelho, M.B. (1991). Vitamin Stability in Premixes and Feeds: A practical
approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Cohen, H., and Lapointe, M.R. (1980). J. Assoc. Off. Anal. Chem. 63, 1254.
Combs Jr., G.F., and Combs, S.B. (1986). The Role of Selenium. Academic Press,
Inc., New York.
Consumer Report on Health. (1993). 5(4), 33.
Cordy, D.R. (1954). Cornell Vet. 44, 310.
Corino, C., Oriani, G., Pantaleo, L., Pastorelli, G., and Salvatori, G. (1999). J.
Anim. Sci. 71, 1755.
Corrigan, J. (1982). Ann. N.Y. Acad.Sci. 393, 361.
Cort, W.M., Vincente, T.S., and Waysek, E.H. (1983). J. Agric. Food Chem. 31,
1330.
Creech, B.G., Feldman, G.L., Ferguson, I.M., Reid, R.L., and Couch, J.R.
(1957). J. Nutr. 62, 83.
Davies, J.A., Quintanilha, A.T., Brooks, G.A., and Packer, L. (1982). Biochem.
Biophys. Res. Comm. 107, 1198.
Dewes, H.F. (1981). N.Z. Vet. J. 29, 83.
Dierenfeld, E.S. (1989). J. Zoo. Wildl. Med. 20, 3.
Dierenfeld, E.S. (1994). J. Nutr. 124, 2579S.
Dierenfeld, E.S., and Dolensek, E.P. (1988). Zoo Biol. 7, 165.
Dierenfeld, E.S., and Traber, M.G. (1992). In Vitamin E in Health and Disease(L. Packer and J. Fuchs, eds.), p. 345. Marcel Dekker, New York.
Dill, S.G., and Rebhun, W.C. (1985). Compend. Cont. Educ. Pract. Vet. 7, S627.
Dove, C.R., and Ewan, R.C. (1990). J. Anim. Sci. 68, 2407.
Dove, C.R., and Ewan, R.C. (1991). J. Anim. Sci. 69, 1994.
Drouchner, W. (1976). Ubers Terernahrung 4, 93.
Dunkley, W.L., Ronning, M., Franke, A.A., and Robb, J. (1967). J. Dairy Sci.
50, 492.
Elvehjem, C.A., Gonce Jr., J.E., and Newell, G.W. (1944). J. Pediatr. 24, 436.
Erskine, R.J., Eberhart, R.J., Grasso, P.J., and Scholz, R.W. (1989). Proc. Am.
Assoc. Bovine Pract. p. 119, Calgary, Alberta, Canada.
Evans, H.M. (1925). Proc. Natl. Acad. Sci. U.S.A. 11, 373.
Faustman, C., Cassens, R.G., Schaefer, D.M., Buege, D.R., and Scheller, K.K.
(1989a). J. Food Sci. 54(2), 485.
Faustman, C., Cassens, R.G., Schaefer, D.M., Buege, D.R., Williams, S.N., andScheller, K.K. (1989b). J. Food Sci. 54(4), 838.
218

V I TA M I N E

Finch, J.M., and Turner, R.J. (1989). Vet. Immunol. Immunopathol. 23, 245.
Franklin, S.T., Sorenson, C.E., and Hammell, D.C. (1998). J. Dairy Sci. 81,
2623.
Friedrichsen, J.V., Arascott, G.H. and Willis, D.L. (1980). Nutr. Rep. Int. 22, 41.
Fritsche, K.L., and McGuire, S.O. (1996). J. Nutr. Biochem. 7, 623.
Gallo-Torres, H.E. (1980). In Vitamin E. A Comprehensive Treatise (L.S.
Machlin, ed.), p. 193. Marcel Dekker, New York.
Gardner, H.W. (1989). Free Radic. Biol. Med. 7, 65.
Gatlin, D.M. III, Poe, W.E., and Wilson, R.P. (1986). J. Nutr. 116, 1061.
Gershoff, S.N., and Norkin, S.A. (1962). J. Nutr. 77, 303.
Gill, D.R., Smith, R.A., Hicks, R.B., and Ball, R.L. (1986). Ok. Agra. Exp. Sta.
Res. Rep. MP 118, 240.
Gitter, M., and Bradley, R. (1978). Vet. Rec. 103, 24.
Glienke, L.R., and Ewan, R.C. (1974). J. Anim. Sci. 39, 975(Abstr).
Golub, M.S., and Gershwin, M.E. (1985). In Animal Stress (G.P. Moberg, ed.)Am. J. Physiol. Soc. Bethesda, Maryland.
Gullickson, T.W., Palmer, L.S., Boyd, W.L., Nelson, J.W., Olson, F.C., Calverley,
C.E., and Boyer, P.D. (1949). J. Dairy Sci. 32, 495.
Guss, S.B. (1977). Management and Diseases of Dairy Goats, Dairy Goat
Journal Publ. Corp., Scottsdale, Arizona.
Hadden, J.W. (1987). Ann. N.Y. Acad. Sci. 496, 39.
Haeger, K. (1974). J. Clin. Nutr. 27, 1179.
Hardy, B., and Frape, D.L. (1983). Micronutrients and Reproduction, No. 1895.
Hoffmann-La Roche, Basel.
Hargis, A.M., Haupt, K.H., and Prieur, D.J. (1985). Compend. Contin. Educ.
Pract. Vet. 7(4), 306.
Harris, P.L., and Embree, N.D. (1963). Am. J. CLin. Nutr. 13, 385.
Harrison, J.H., Hancock, D.D., and Conrad, H.R. (1984). J. Dairy Sci. 67, 123.
Hartley, W.J., and Grant, A.B. (1961). Fed. Proc. Fed. Am. Soc. Exp. Biol. 20,
679.
Harvey, J.D., and Bieber-Wlaschny, M. (1988). Feedstuffs 60(12), 15.
Hayes, K.C., and Rousseau, J.E. (1970). Lab. Anim. Care 20, 48.
Hayes, K.C., Nielsen, S.W., and Rousseau Jr., J.E. (1969). J. Nutr. 99, 196.
Heinzerling, P.H., Rollin, C.F., Nockels, C., Quarles, L., and Tengerdy, R.P.
(1974). Proc. Exp. Biol. Med. 146, 279.
Helgebostad, A., and Ender, F. (1973). Acta Agric. Scand. Suppl. 19, 79.
Herdt, T.H., and Stowe, H.D. (1991). Vet. Clin. North Am. Food Anim. Pract.
7, 391.
Hidiroglou, M. (1989). J. Dairy Sci. 72, 1067.
Hidiroglou, M., and Charmley, E. (1991). Am. J. Vet. Res. 52, 640.
Hidiroglou, M., and Singh, K. (1991). J. Dairy Sci. 74, 2718.
Hidiroglou, M., Hoffman, I., and Jenkins, K.J. (1969). Can. J. Physiol.
Pharmacol. 47, 953.
Hidiroglou, M., McAllister, A.J., and Williams, C.J. (1987). J. Dairy Sci. 70,
1281.
Hidiroglou, N, and McDowell, L. (1987). Int. J. Vitam. Res. 57, 261.
Hidiroglou, N., Laflamme, L.F., and McDowell, L.R. (1988a). J. Anim. Sci. 66,
3227.
219

CHAPTER FOUR

Hidiroglou, N., McDowell, L.R., and Pastrana, R. (1988b). Int. J. Vitam. Nutr.
Res. 58, 189.
Hidiroglou, N., McDowell, L.R., and Balbuena, O. (1989). J. Dairy Sci. 72,
1793.
Hidiroglou, N., Butler, G., and McDowell, L.R. (1990). J. Anim. Sci. 68, 782.
Hidiroglou, N., McDowell, L.R., Papas, A.M., Antapli, M., and Wilkinson, N.S.
(1992a). J. Anim. Sci. 70, 2556.
Hidiroglou, N., Cave, N., Atwal, A.S., Farnsworth, E.R., and McDowell, L.R.
(1992b). Ann. Rech. Vet. 23, 337.
Hidiroglou, N., McDowell, L.R., Batra, T.R., and Papas, A.M. (1994). Reprod.
Nutr. Dev. 34, 133.
Hill, G.M., Link, J.E., Meyer, L., and Fritsche, K.L. (1999). J. Anim. Sci. 77,
1762.
Hintz, H.F. (1996). In Vitamin E in Animal Nutrition and Management, (M.B.
Coelho, ed.), 2nd Ed., p. 449, BASP Corporation, Mount Olive, New
Jersey.
Hoekstra, W.G. (1975). Fed. Proc. 34, 2083.
Hoffmann-La Roche. (1972). Vitamin E, No. 1206. F. Hoffmann-La Roche &Co. Ltd., Basel, Switzerland.
Hoffmann-La Roche. (1991). Vitamin E for Ruminants. RCD 8361/191. Hoffmann-La Roche Inc., Nutley, New Jersey.
Hoffmann-La Roche. (1994). Vitamin Nutrition for Ruminants. RCD 87751/
894. Hoffmann-La Roche Inc., Nutley, New Jersey.
Hove, E.L., Fry, G.S., and Schwarz, K. (1958). Proc. Soc. Exp. Biol. Med. 98,
27.
Hutchinson, L.J., Scholz, W., and Drake, T.R. (1982). J. Am. Vet. Med. Assoc.
181, 581.
Jacob, R.A. (1995). Nutr. Res. 15, 755.
Jager, F.C. (1977). Nutr. Metabol. 14, 210.
Jensen, L.S. (1968). Proc. Soc. Exp. Biol. Med. 128, 970.
Jensen, L.S., and McGinnis, J. (1957). Poult. Sci. 36, 212.
Jensen, S.K., Engberg, R.M., and Hedemann, M.S. (1999). J. Nutr. 129, 1355.
Jialal, I., and Grundy, S.M. (1993). Circulation 88, 2780.
King, R.L., Burrows, F.A., Hemken, R.W., and Bashore, D.L. (1967). J. Dairy
Sci. 50, 943.
Kozicki, L., Silva, R.G., and Barnabe, R.C. (1981). Vet. Med. 28, 538.
Krukovsky, V.N., and Loosli, J.K. (1952). J. Dairy Sci. 35, 834.
Krukovsky, V.N., Whiting, F., and Loosli, J.K. (1950). J. Dairy Sci. 33, 791.
Langseth, L. (1995a). Antiox. Vitam. Newsl. 11, 1.
Langseth, L. (1995b). Antiox. Vitam. Newsl. 12, 1.
Langweiler, M., Sheffy, B.E., and Schultz, R.D. (1983). Am. J. Vet. Res. 44, 5.
Lauridsen, C., Nielsen, J.H., Henckel, P., and Sørensen, M.T. (1999). J. Anim.
Sci. 77, 105.
Lee, R.W., Stuart, R.L., Perryman, K.R., and Ridenour, K.W. (1985). J. Anim.
Sci. 61(Suppl. 1), 425.
Linder, M.C. (1985). In Nutritional Biochemistry and Metabolism (M.C. Linder,ed.). Elsevier Sci. Publ. Co., New York.
Liu, J.F., and Huang, C.J. (1995). J. Nutr. 125, 3071.
220

V I TA M I N E

Liu, J.F., and Huang, C.J. (1996). J. Nutr. 126, 2227.
Liu, S.K., Dolensek, E.P., Adams, C.R. and Tappe, J.P. (1983). J. Am. Vet. Med.
Assoc. 183, 1266.
Liu, S.K., Dolensek, E.P. and Tappe, J.P. (1985). Heart Vessels 1(Suppl.), 282.
Liuzzi, J.P., Cioccia, A.M., and Hevia, P. (1998). J. Nutr. 128, 2467.
Lopez-Bote, C.J., Rey, A.I., Sanz, M., Gray, J.I., and Buckley, D.J. (1997). J.
Nutr. 127, 1176.
Machlin, L.J. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 99.
Marcel Dekker, Inc., New York.
Mahan, D.C. (1991). J. Anim. Sci. 69, 2904.
Mahan, D.C., and Vallet, J.L. (1997). J. Anim. Sci. 75, 2731.
Marin-Guzman, J., Mahan, D.C., Chung, Y.K., Pate, J.L., and Pope, W.F.
(1997). J. Anim. Sci. 75, 2994.
Marin-Guzman, J., Mahan, D.C., Jones, L.S., and Pate, J.L. (1989). Ohio Swine
Res. Ind. Rep. The Ohio State Univ., Anim. Sci. Dept. Series 89-1, p. 20.
Mason, R.M., Ostrum, P., and McDowell, L.R. (1985). Florida Cattlemen,
March, 1993.
Maynard, L.A., Loosli, J.K., Hintz, H.F., and Warner, R.G. (1979). In Animal
Nutrition, 7th Ed., p. 283. McGraw-Hill, New York.
McCay, P.B., Gibson, D.D., and Hornbrook, K.R. (1981). Fed. Proc. 40, 199.
McDowell, L.R. (1985). In Nutrition of Grazing Ruminants in Warm Climates(L.R. McDowell, ed.), p. 339. Academic Press, Orlando, Florida.
McDowell, L.R. (1992) Minerals in Animals and Human Nutrition. Academic
Press, San Diego, California.
McDowell, L.R. (1997). Minerals for Grazing Ruminants in Tropical Regions,
3rd Ed. University of Florida, Gainesville.
McDowell, L.R., and Williams, S.N. (1991). In 2nd Annual Florida Ruminant
Nutrition Symposium, p. 46. Gainesville, Florida.
McDowell, L.R., Froseth, J.H., Piper, R.C., Dyer, I.A., and Kroening, G.H.
(1977). J. Anim. Sci. 45, 1326.
McDowell, L.R., Williams, S.N., Hidiroglou, N., Njeru, C.A., Hill, G.M.,
Ochoa, L., and Wilkinson, N.S. (1996). Anim. Feed Sci. Tech. 60, 273.
McGuire, S.O., Alexander D.W., and Fritsche, K.L. (1997). J. Nutr. 127, 1388.
McIntosh, G.H., Lawson, C.A., Bulman, F.H., and McMurchie, E.J. (1985).
Proc. Nutr. Soc. Australia 10, 207.
McMurray, C.H., and Blanchflower, W.J. (1979). J. Chromatogr. 178, 525.
McMurray, C.H., and Rice, D.A. (1984). Vitamin E and Selenium Deficiency
Diseases, No. 1953. Hoffmann-La Roche, Basel.
McMurray, C.H., Rice, D.A., and Blanchflower, W.J. (1980). Proc. Nutr. Soc.
39, 65(Abstr).
Meydani, S.N. (1995). Nutr. Rev. 53, 552.
Michel, R.L., Whitehair, C.K., and Keahey, K.K. (1969). J. Am. Vet. Med. Assoc.
155, 50.
Miller, W.H. Jr. (1989). Vet. Clin. North Am. Small Anim. Pract. 19(3), 497.
Moore, R.M., and Kohn, C.W. (1991). Compend. Cont. Educ. Pract. Vet. 13,
476.
Moore, T., Sharman, I.M., and Ward, R.J. (1957). J. Sci. Food Agric. 8, 97.
Morrow, J.L., McGlone, J.J., Tribble, L.F., and Stansbury, W.F. (1987). Anim.
221

CHAPTER FOUR

Sci. Res. Rpt. Texas Tech., p. 14, Lubbock, Texas.
Morand-Fehr, P. (1981). In Goat Production, p. 193 (G. Gall, Ed.). Academic
Press, New York.
Muller G.H., Kirk, R.W., and Scott, D.W. (1989). In Small Animal Dermatology,
4th Ed., p. 658. W.B. Saunders Co., Philadelphia.
Munson, T.O., Holzworth, J., Small, E., Witzel, S., Jones, T.C., and Luginbuhl,
H. (1958). J. Am. Vet. Med. Assoc. 133, 563.
Muth, O.H. (1955). J. Am. Vet. Med. Assoc. 126, 355.
Neilsen, J., Fisher, A.N., and Pedersen, A.H. (1953). J. Dairy Res. 20, 333.
Nicholson, J.W., St. Laurent, A.M., McQueen, R.E., and Charmley, E. (1991).
Can. J. Anim. Sci. 71, 143.
Njeru, C.A., McDowell, L.R., Wilkinson, N.S., Linda, S.B., Williams, S.N., andLentz, E.L. (1992). J. Anim. Sci. 70, 2562.
Njeru, C.A., McDowell, L.R., Wilkinson, N.S., Linda, S.B., and Williams, S.N.
(1994a). J. Anim. Sci. 72, 1636.
Njeru, C.A., McDowell, L.R., Wilkinson, N.S., Linda, S.B., Rojas, L.X., andWilliams, S.N. (1994b). J. Anim. Sci. 72, 739.
Noble, R.E., Moore, J.H., and Harfoot, C.G. (1974). Br. J. Nutr. 31, 99.
Nockels, C.F. (1990). In Proc. National Feed Ingredients Association Nutrition
Institute: Developments in Vitamin Nutrition and Health Applications,Kansas City, Missouri. National Feed Ingredients Association (NFIA), Des
Moines, Iowa.
Nockels, C.F. (1991). Vitamin E requirement of beef cattle: Influencing factors.
BASF Technical Symposium, p. 40. Bloomington, Minnesota.
Nockels, C.F., Odde, K.G., and Craig, A.M. (1996). J. Anim. Sci. 74, 672.
NRC. Nutrient Requirements of Domestic Animals. National Academy of
Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of SciencesNational Research Council, Washington, D.C.
Obel, A.L. (1953). Acta Pathol. Microbiol. Scand. Suppl. 94, 5.
Ochoa, L., McDowell, L.R., Williams, S.N., Wilkinson, N., Boucher, J., andLentz, E.L. (1992). J. Anim. Sci. 70, 2568.
Orstadius, K., Nordstrom, G., and Lannek, N. (1963). Cornell Vet. 53, 60.
Oski, F.A., and Barness, L.A. (1967). J. Pediatr. 70, 211.
Padh, H. (1991). Nutr. Rev. 49, 65.
222

V I TA M I N E

Panganamala, R.V., and Cornwell, D.C. (1982). Ann. N.Y. Acad. Sci. 393, 376.
Pappenheimer, A.M. (1942). Am. J. Pathol. 18, 169.
Pappenheimer, A.M., and Goettsch,M. (1931). J. Exp. Med. 53, 11.
Pappu, A.S., Fatterpacker, P., and Sreenivasan, A. (1978). Biochem. J. 172, 115.
Peplowski, M.A., Mahan, D.C., Murray, F.A., Moxon, A.L., Cantor, A.H., andEkstrom, K.E. (1981). J. Anim. Sci. 51, 344.
Piper, R.C., Froseth, J.A., McDowell, L.R., Kroening, G.H., and Dyer, I.A.
(1975). Am. J. Vet. Res. 36, 273.
Poston, H.A., G.F. Combs Jr., and Leibovitz, L. (1976). J. Nutr. 106, 892.
Pruiett, S.P., Morrill, J.L., Blecha, F., Reddy, P.G., Higgins, J., and Anderson,
N.V. (1989). J. Anim. Sci. 67(Suppl 1), 243.
Putnam, M.E. (1984). Pig Vet. Soc. Proc. 9, 178.
Putnam, M.E. (1986). Mod. Vet. Pract. 67, 121.
Rajaraman, V., Nonneche, B.J., Franklin, S.T., Hammell, D.C., and Horst, R.L.
(1998). J. Dairy Sci. 81, 3278.
RDA. (1989). Recommended Dietary Allowances, 9th Ed., National Academyof Sciences-National Research Council, Washington, D.C.
Reddy, P.G., Morrill, J.J., and Frey, R.A. (1987a). J. Dairy Sci. 70, 123.
Reddy, P.G., Morrill, J.L., Minocha, H.C., and Stevenson, J.S. (1987b). J. Dairy
Sci. 70, 993.
Reffett, J.K, Spears, J.W., and Brown, Jr., T.T. (1988). J. Anim. Sci. 66, 1520.
Rice, D.A., Blanchflower, W.J., and McMurray, C.H. (1981). Vet. Rec. 109, 161.
Riedel-Caspari, G., Schmidt, F.W., Gunther, K.D., and Wagner, K. (1986). J. Vet.
Med. 33, 650.
Riis, R.C., Sheffy, B.E., and Loew, E. (1981). Am. J. Vet. Res. 42, 74.
Ringer, D.H., and Abrams, G.D. (1970). J. Am. Vet. Med. Assoc. 157, 1228.
Roquet, J., Nockels, C.F., and Papas, A.M. (1992). J. Anim. Sci. 70, 2542.
Rumsey, T.S. (1975). Feedstuffs 47, 30.
Sanders, S.K., Morgan, J.B., Wulf, D.M., Tatum, J.D., Williams, S.N., andSmith, G.C. (1997). J. Anim. Sci. 75, 2634.
Sarter, M., and Van Der Linde, A. (1987). Neurobiol. Aging 8, 297.
Scott, D.S., and Sheffy, B.E. (1987). Companion Anim. Pract. 1(4), 42.
Scott, D.W., and Walton, D.K. (1985). J. Am. Anim. Hosp. Assoc. 21, 345.
Scott, D.W., Walton, D.K., Manning, T.O., Smith, C.A., and Lewis, R.M.
(1983). J. Am. Anim. Hosp. Assoc. 19, 481.
Scott, M.L. (1980). Fed. Proc. Fed. Am. Soc. Exp. Biol. 39, 2736.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Sergeev, I.N., Aarkhapchev, Y.P., and Spirichev, V.B. (1990). Biokhimiya-Engl.
Tr. 55, 1483.
Sheffy, B.E. (1979). Compend. Cont. Ed. 1, 759.
Sheffy, B.E., and Schultz, R.D. (1979). Cornell Vet. 68, 48.
Sheffy, B.E., and Williams, A.J. (1981). Animal Nutrition Events, p. 1. 1781
Hoffmann-La Roche, Basel, Switzerland.
Shimotori, N., Emerson, G.A., and Evans, H.M. (1940). J. Nutr. 19, 547.
Sitrin, M.D., Liberman, F., Jensen, W.E., Noronha, A., Milburn, C., andAddington, W. (1987). Ann. Intern. Med. 107, 51.
Smith, K., Hogan, J.S., and Weiss, W.P. (1997). J. Anim. Sci. 75, 1659.
223

CHAPTER FOUR

Smith, K.L., and Conrad, H.R. (1987). Proc. Roche Technical Symposium. The
Role of Vitamins on Animals Performance and Immune Response, p. 47.
Daytona Beach, Florida.
Smith, K.L., Harrison, J.H., Hancock, D.D., Todhunter, D.A., and Conrad, H.R.
(1984). J. Dairy Sci. 67, 1293.
Smith, P.J., Tappel, A.L., and Chow, C.K. (1974). Nature (London) 247, 392.
Smith, S.E. (1970). In Duke’s Physiology of Domestic Animals, (M.J. Swenson,ed.), 8th Ed., p. 634. Cornell Univ. Press Ithaca, New York.
Sokol, R.J. (1989). Free Radic. Biol. Med. 6, 189.
Soler-Velásquez, M.P., Brendemuhl, J.H., McDowell, L.R., Sheppard, K.A.,
Johnson, D.D., and Williams, S.N. (1998). J. Anim. Sci. 76, 110.
Stephan, Z.F., and Hayes, K.C. (1978). Fed. Proc. 73, 706(Abstr).
Stephens, L.C., McChesney, A.E., and Nockels, C.F. (1979). Br. Vet. J. 135, 291.
Stuart, R.L. (1987). Proc. The Role of Vitamins on Animal Performance and
Immune Response, p. 67. Hoffmann-La Roche, Nutley, New Jersey.
Sure, B. (1924). J. Biol. Chem. 58, 693.
Tanaka, K., Hashimoto, T., Tokumaru, S., Iguchi, H., and Kojo, S. (1997). J.
Nutr. 127, 2060
Taylor, S.I., Lambden, M.P., and Tappel, A.L. (1976). Lipids 11, 530.
Tengerdy, R.P. (1980). In Vitamin E: A Comprehensive Treatise (L.J. Machlin,ed.). Marcel Dekker, New York.
Tengerdy, R.P. (1989). Ann. N.Y. Acad. Sci. 570, 335.
Thompson, J.N., and Scott, M.L. (1969). J. Nutr. 97, 335.
Tiege, J., Saxegaard, F., and Froslie, A. (1978). Acta Vet. Scand. 19, 133.
Tollersrud, S. (1973). Acta Agric. Scand. Suppl. 19, 124.
Tollerz, G., and Lannek, N. (1964). Nature (London), 201, 846.
Trapp, A.L., Keahey, K.K., Whitenack, D.L., and Whitehair, C.K. (1970). J. Am.
Vet. Med. Assoc. 157, 289.
Turley, C.P., and Brewster, M.A. (1993). J. Nutr. 123, 1305.
Turner, R.J., and Finch, J.M. (1990). J. Comparative Pathol. 102, 16.
Uebersax, M.A., Uebersax, M.L., and Dawson, L.E. (1978). Poult. Sci. 57, 937.
Ullrey, D.E. (1969). Michigan State University Bulletin AH-SW-695, p. 16.
Ullrey, D.E. (1981). J. Anim. Sci. 53, 1039.
Ullrey, D.E. (1982). Proc. Distill. Feed Conf. Distill. Feed Res. Counc.
Cincinnati 37, 81.
Ullrey, D.E., Miller, E.R., Ellis, D.J., Orr, D.E., Hitchcock, J.P., Keahey, K.K.,
and Trapp, A.L. (1971). Mich. State Univ. Agr. Exp. Sta. Rep. Swine Res.
148, 48.
Underwood, E.J. (1981). The Mineral Nutrition of Livestock. Commonwealth
Agricultural Bureaux, London.
Valberg, S., Jonsson, L., Lindholm, A., and Holgrem. N. (1993). Equine Vet. 25.
11.
Van Kruiningen, J.J. (1967). Gastroenterology 53, 114.
Van Saun, R.J., Herdt, T.H., and Stowe, H.D. (1989). J. Nutr. 119, 1156.
Van Vleet, J.F. (1975). J. Am. Vet. Med. Assoc. 166, 769.
Van Vleet, J.F., Crawley, R.R., and Amstutz, H.E. (1977). J. Am. Vet. Med.
Assoc. 171, 443.
Van Vleet, J.F., Runnels, L.J., Cook Jr., J.R., and Scheidt, A.B. (1987). Am. J.
224

V I TA M I N E

Vet. Res. 48, 1520.
Velásquez-Pereira, J., Risco, C.A., Chenoweth, P.J., McDowell, L.R., Prichard,D., Martin, F.G., Wilkinson, N.S., Williams, S.N., and Staples, C.R. (1998).
J. Anim. Sci. 76, 2894.
Velásquez-Pereira, J., Risco, C.A., McDowell, L.R., Staples, C.R., Prichard, D.,
Chenoweth, P.J., Martin, F.G., Williams, S.N., Rojas, L.X., Calhoun, M.C.,
and Wilkinson, N.S. (1999). J. Dairy Sci. 82, 1240.
Vitamin E Research and Information Service. (1993). An Overview of Vitamin
E Efficacy in Humans. VERIS, LaGrange, Illinois, 24 pp.
Walter, W.D., and Jensen, L.S. (1963). J. Nutr. 80, 327.
Weiss, W.P. (1998). J. Dairy Sci. 81, 2493.
Weiss, W.P., Hogan, J.S., Smith, K.L., and Hoblet, K.H. (1990). J. Dairy Sci. 73,
381.
Weiss, W.P., Hogan, J.S., Todhunter, D.A., and Smith, K.L. (1997). J. Dairy Sci.
80, 1728.
Whanger, P.D. (1981). In Selenium in Biology and Medicine, (J.E. Spallholz, J.L.
Martin, and H.E. Ganther, eds.), p. 230. AVI, Westport, Connecticut.
Whitehair, C.K., Vale, O.E., Loudenslager, M., and Miller, E.R. (1983). Mich.
State Univ. Agr. Exp. Sta. Res. Swine Rep. 456, 9.
Worden, A.N. (1958). Vet Rec. 70, 189.
Wretlind, B.K., Orstadius, K., and Lindberg, P. (1959). Zentralbl. Vet. Med. 6,963.
Wuryastuti, H., Stowe, H.D., Bull, R.W., and Miller, E.R. (1993). J. Anim. Sci.
71, 2464.
Young, L.G., Lun, A., Pos, J., Forshaw, R.P., and Edmeades, D.E. (1975). J.
Anim. Sci. 40, 495.
Young, L.G., Miller, R.B., Edmeades, D.M., Lun, A., Smith, G.C., and King, G.J.
(1978). J. Anim. Sci. 47, 639.

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chapter fiveV I TA M I N K

INTRODUCTION
Vitamin K was the last fat-soluble vitamin to be discovered. For
many years after its discovery, vitamin K appeared to be limited in its
function to only the normal blood-clotting mechanism. However, vitamin K-dependent proteins have been identified that suggest roles for thevitamin in addition to that of blood coagulation. Because of the bloodclotting function, vitamin K was previously referred to as the “coagulation vitamin,” “antihemorrhagic vitamin,” and “prothrombin factor.”
Vitamin K is indispensable for maintaining the function of the blood
coagulation system in humans and all investigated animals. Even thoughvitamin K is synthesized by intestinal microorganisms, deficiency signs
have been observed under field conditions. Poultry, and to a lesser degree pigs, are susceptible to vitamin K deficiency. In ruminants a deficiency can be caused by ingestion of spoiled sweet clover hay, which isa natural source of dicumarol (a vitamin K antagonist). In human nutrition, vitamin K is most required in infants because of insufficient intestinal synthesis and in adults in whom fat absorption is impaired.

HISTORY
This history of vitamin K discovery has been reviewed (McCollum,
1957; Almquist, 1975; Olson and Suttie, 1978; Loosli, 1991; Suttie,
1991). Hemorrhagic signs and symptoms frequently had been seen in infants, in patients with obstructive jaundice, in experimental animals
maintained on purified diets, and in cattle eating moldy sweet clover
hay. The presence of a dietary antihemorrhagic factor was first suspected
in 1929, when Henrik Dam of Denmark fed chickens a purified low-fat
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diet in an attempt to determine whether they were able to synthesize
cholesterol. He noted that chickens became anemic and developed subcutaneous and intermuscular hemorrhages and that blood taken fromthese animals clotted slowly. Hemorrhagic signs were reported by otherworkers using ether-extracted fish meal for chicks (McFarlane et al.,1931). Since the condition was prevented by unextracted fish meal, thecurative factor was fat soluble. When the ether extracts were added backto the feed, the animals returned to normal. However, studies in a number of laboratories soon demonstrated that this disease could not be
cured by administration of any of the known fat-soluble vitamins (A, D,
and E) or other known physiologically active lipids.
Dam continued to study the distribution and lipid solubility of theactive component in vegetable and animal sources, and in 1935 he proposed that the antihemorrhagic vitamin of the chick was a new fat-soluble vitamin that he called vitamin K, from the Danish word for coagulation (koagulation). At the same time, Almquist and Stokstad (1935) of
California, described their success in curing the hemorrhagic disease
with ether extracts of alfalfa and clearly pointed out that microbial action in fish meal and bran preparations could lead to development of antihemorrhagic activity. Of historical note, it is interesting that research
journals of Almquist and Stokstad actually discovered what were later
referred to as vitamins K1 and K2 in 1928. The paper reporting their results was delayed by university administrators, and when it was finallysubmitted to the journal Science, it was rejected. About that time, Dam’s
research was published. When Dam (with Doisey) later received theNobel prize for the discovery of vitamin K, he had reportedly expected
Almquist to share the prize. Doisey also contributed to the knowledgeof the role of vitamin K in blood clotting (Loosli, 1991). Dam et al.
(1936) demonstrated that prothrombin activity decreased in vitamin Kdeficient chicks. At about the same time, the hemorrhagic condition resulting from obstructive jaundice (deficiency of bile) was shown to be
due to poor absorption of vitamin K, and bleeding episodes were attributed to lack of plasma prothrombin (Suttie, 1991). This hemorrhagic
condition was originally thought to be due solely to a lowered concentration of plasma prothrombin (factor II), but during the 1950s it was
shown that the synthesis of three other clotting factors—VII, IX, andX—were also depressed in the deficient state.
A number of groups were involved in attempts to isolate and characterize this new vitamin, and Dam’s collaboration with Karrer, of the
University of Zurich, resulted in isolation of the vitamin from alfalfa asa yellow oil (Suttie, 1991). Research proceeded to show that a large
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number of chemical compounds possess some degree of vitamin K activity. The main source present in plants was referred to as K1, and theform synthesized by microflora as K2. The simplest source of vitamin K3
is menadione, produced by laboratory synthesis. Vitamin K1 was synthesized by three laboratories in 1939, while the structure of K2 was elucidated in 1940 but was not synthesized until 1958 by Isler and coworkers in Switzerland.
It was only in 1974 that the metabolic role of vitamin K was elucidated, when γ-carboxyglutamate was found to be present in the vitamin
K-dependent proteins but absent from the abnormal precursors that circulate in deficiency (Nelsestuen et al., 1974; Stenflo, 1974). In more recent years, four additional vitamin K-dependent proteins have been
identified in plasma (C, S, Z, and M) and a major protein of the bonematrix (osteocalcin).

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTSThe general term vitamin K is now used to describe not a single
chemical entity but a group of quinone compounds that have characteristic antihemorrhagic effects. Vitamin K is a generic term for a homologous group of fat-soluble vitamins consisting of a 2-methyl-1,4-naphthoquinone, with the various isomers differing in the nature and lengthof the side chain (Fig. 5.1). When synthesized in the laboratory, it yields
a mixture of cis- and trans-isomers; the cis-form has little, if any, vitamin K activity. Vitamin K extracted from plant material was named
phylloquinone or vitamin K1. Phylloquinone has a phytyl side chain
composed of four isoprene units, the first of which contains a doublebond.
Vitamin K-active compounds from material that had undergone bacterial fermentation were named menaquinones or vitamin K2. The simplest form of vitamin K is the synthetic menadione (K3), which is composed of the active nucleus (2-methyl-1,4 naphthoquinone) and has noside chain. The menaquinone family of K2 homologs is a large series ofvitamins containing unsaturated side chains that differ in number of isoprene units. Numerous natural analogs have been isolated, almost all ofwhich are variations of the side chain at position 3. Some of these chainsare quite long, with as many as 65 carbon atoms in some bacterial vitamin K forms, but none is specifically required for activity. Menaquinone4 is synthesized in liver from ingested menadione or changed to a biologically active menaquinone by intestinal microorganisms.
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Fig. 5.1 Chemical structures of the vitamin K compounds.

Vitamin K is a golden yellow viscous oil. Natural sources of vitamin
K are fat soluble, stable to heat, and labile to oxidation, alkali, strong
acids, light, and irradiation. Vitamin K1 is slowly degraded by atmospheric oxygen but fairly rapidly destroyed by light. In contrast to natural sources of vitamin K, some of the synthetic products, such as saltsof menadione, are water soluble.
A number of vitamin K antagonists exist that increase the need forthis vitamin. Vitamin K deficiency is brought about by ingestion of
dicumarol (Fig. 5.2), an antagonist of vitamin K, or by feeding sulfonamides (in monogastric species) at levels sufficient to inhibit intestinalsynthesis of vitamin K. Mycotoxins, toxic substances produced bymolds, are also antagonists causing vitamin K deficiency. A hemorrhagic
disease of cattle that was traced to consumption of moldy sweet clover
hay was described in the 1920s. The destructive agent was found to be
dicumarol, a substance produced from natural coumarins. Dicumarols
are produced by molds, particularly those that attack sweet clover, thus
giving rise to the term sweet clover disease (see Deficiency). During theprocess of spoiling, harmless natural coumarins in sweet clover are converted to dicumarol (bis-hydroxycoumarin), and when toxic hay orsilage is consumed by animals, hypoprothrombinemia results, presumably because dicumarol combines with the proenzyme to prevent formation of the active enzyme required for the synthesis of prothrombin.
It probably also interferes with synthesis of factor VII and other coagulation factors.
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Fig. 5.2 Antagonists of vitamin K include coumarin derivatives.

Dicumarol serves as an anticoagulant in medicine to slow blood coagulation in people with cardiovascular disease to avoid intravascularblood clots, just as vitamin K under other conditions increases coagulation time. Thus additional vitamin K will overcome this action by
dicumarol. Goplen and Bell (1967) showed that in cattle, vitamin K1 ismuch more potent than vitamin K3 as an antidote to dicumarol. The
most successful dicumarol for long-term lowering of the vitamin K-dependent clotting factors is warfarin (Fig. 5.2), which is widely used as arodenticide. Concern has been expressed because of the identification of
anticoagulant-resistant rat populations. Spread of this resistance led tosynthesis of new and more effective coumarin derivatives (Hadler andShadbolt, 1975; Suttie, 1991).
Because of the use of dicumarol derivatives as clinical anticoagulants, investigation of the mechanism of action of dicumarol has been ofinterest to vitamin K researchers (Suttie, 1991). Investigations have centered on interconversion of vitamin K and its 2,3-epoxide as the site of
coumarin action. One hypothesis is that metabolic effects of these compounds are the consequence of their inhibition of the microsomal epoxide reductase (Bell, 1978). The epoxide apparently acts as a competitive
inhibitor of vitamin K at its site of action, and a coumarin such as warfarin is an inhibitor of vitamin K action only to the extent that it increases the cellular ratio of oxide to the vitamin.

ANALYTICAL PROCEDURES
Vitamin K can be analyzed by a variety of color reactions or by direct spectroscopy. The chemical reactivity is a function of the naphthoquinone nucleus, and other quinones also react with many of the avail231

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able colorimetric assays. For spectroscopy to be successful, a significant
amount of separation is often required to eliminate interfering substances present in crude extracts.
Improved procedures for vitamin K analyses have used high-pressure liquid chromatography (HPLC) as an analytical tool to investigate
the vitamin as well as the interconversion of vitamin K to its 2,3-epoxide. The HPLC method is highly suitable for vitamin K analysis becauseof its high sensitivity, specificity, and accuracy (Manz and Maurer, 1982;
Shino, 1988). In addition to feeds, phylloquinone concentrations inblood, milk, and selected tissues are now more commonplace.
The classic biological assay for the amount of vitamin K in an unknown source is a determination of the whole-blood clotting time ofchicks. This assay measures prothrombin time since prothrombin is themost limiting vitamin K-dependent blood-clotting factor in chicks receiving vitamin K-deficient diets. Young chicks are raised on a vitamin
K-deficient diet until their whole-blood clotting time is increased to fourto seven times normal. The usual one-stage method for measuring prothrombin time consists of adding excess thromboplastin (obtained fromchick brain or another source) and calcium to the blood and then measuring the time for the blood to clot. The response of chicks receiving astandard vitamin K preparation is compared to the prothrombin time
obtained when a supplement containing an unknown quantity of vitamin K is added to the vitamin K-deficient basal diet (Scott et al., 1982;
Weiser and Tagwerker, 1982).

METABOLISM
Absorption and Transfer
Like all fat-soluble vitamins, vitamin K is absorbed in association
with dietary fats and requires the presence of bile salts and pancreatic
juice for adequate uptake from the alimentary canal. Absorption of vitamin K depends on its incorporation into mixed micelles, and optimal
formation of these micellar structures requires the presence of both bileand pancreatic juice. Thus, any malfunction of the fat-absorption mechanism, for example, biliary obstruction, will reduce availability of vitamin K. Unlike phylloquinone and the menaquinones, menadione bisulfites and phosphates are relatively water soluble and therefore areabsorbed satisfactorily from low-fat diets. Male animals are more susceptible to dietary vitamin K deprivation than females, apparently as aresult of stimulation of phylloquinone absorption by estrogens. The ad232

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ministration of estrogens increases absorption in both male and female
animals (Jolly et al., 1977). The lymphatic system is the major route oftransport of absorbed phylloquinone from the intestine in mammals but
by portal circulation in birds, fishes, and reptiles. Shearer et al. (1970)
demonstrated the association of phylloquinone with serum lipoproteins,
but little is known of the existence of specific carrier proteins. The absorption of various forms of vitamin K has been studied and found todiffer significantly. Ingested phylloquinone is absorbed by an energy-dependent process from the proximal portion of the small intestine(Hollander, 1973). In contrast to the active transport of phylloquinone,
menaquinone is absorbed from the small intestine by a passive non–carrier-mediated process. Menadione can be absorbed from both the small
intestine and the colon by a passive process and can be alkylated to a biologically active form (Hollander and Truscott, 1974).
Efficiency of vitamin K absorption has been measured at 10 to 70%,
depending on the form in which the vitamin is administered. Some reports have indicated that menadione is completely absorbed, but phylloquinone only at a rate of 50%. Rats were found to excrete about 60%
of ingested phylloquinone in the feces within 24 hours of ingestion but
only 11% of ingested menadione (Griminger and Donis, 1960;
Griminger, 1984a). However, 38% of ingested menadione but only asmall amount of phylloquinone were excreted via the kidneys in thesame period. The conclusion was that although menadione is well absorbed, it is poorly retained, while just the opposite is true for phylloquinone. Poor retention of menadione can be explained by the need toadd, in the liver, a difarnesyl chain and thus transform it into
menaquinone (vitamin K2) with a 20-carbon chain (MK-4). Apparently
there are quantitative limitations in this biosynthetic step. The menadione not converted is rapidly detoxified and excreted. The turnover ofliver phylloquinone was found to be 2 to 4 times more rapid thanmenaquinone initially, but no difference was found between the sources
at 48 hours (Will and Suttie, 1992). However, phylloquinone was much
more effective than menaquinone in maintaining normal vitamin K status in rats at low dietary concentrations (0.2 μmol/kg diet), whereas at
high dietary concentrations (5.6 μmol/kg diet) they were equally effective.
Konishi et al. (1973) administered radioactive menadione, phylloquinone, or menaquinone-4, to rats and found that radioactive menadione was spread over the whole body much faster than the other two
compounds, but the amount retained in the tissues was low. Martius andAlvino (1964) utilized radioactive menadione to establish that it could
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be converted to a more lipophilic compound that, on the basis of theirlimited characterization, appeared to be menaquinone-4. Therefore,
they concluded that the vitamin K form of animal tissues was
menaquinone-4. They found that when either a menaquinone or a phylloquinone was given to animals, the side chain was removed, probably
by the microorganisms in the gut.
On the contrary, Griminger and Brubacher (1966) showed that amajor portion of the phylloquinone they fed to chicks was absorbed anddeposited in the liver intact, and that as such it had as good biologicalactivity upon prothrombin synthesis as the menaquinone-4, which theyfound in the chick’s liver following feeding of menadione. Therefore,
menaquinone-4 is most likely produced only if menadione is fed, or if
the intestinal microorganisms degrade the dietary K1 or K2 to menadione. The formation of menaquinone-4 is not obligatory for metabolic
activity, since phylloquinone is equally active in bringing about synthesis of the K-dependent blood-clotting proteins (Scott et al., 1982).
Newborn infants have low vitamin K body stores, indicating poor
placental transfer. Correlation of vitamin K between mothers and babies
have suggested that the vitamin passes the placenta only in small quantities (Suzuki et al., 1989). The vitamin is frequently not detectable in thecord blood from mothers with normal plasma levels, and sometimes,
levels from neonates are only half those of their mothers.

Storage and Excretion
About half of the total body pool of vitamin K is in the liver. A number of studies have shown that phylloquinone and menaquinones are
specifically concentrated and retained in the liver. Menaquinone concentrations exceed those of phylloquinone in organs other than liver(Thijssen et al., 1996). Menadione is found to be widely distributed in
all tissues and very rapidly excreted. Although phylloquinone is rapidly
concentrated in liver, it does not have a long retention time (17 hours
half-life) in this organ (Thierry et al., 1970; Shearer et al., 1996).
Therefore, the inability to rapidly develop vitamin K deficiency in most
species results from the difficulty in preventing absorption of the vitaminfrom the diet or from intestinal synthesis rather than from a significant
storage of the vitamin.
Some breakdown products of vitamin K are excreted in the urine.
One of the principal excretory products is a chain-shortened and oxidized derivative of vitamin K, which forms a γ-lactone and is probably
excreted as a glucuronide. Vitamin K oxide has also been identified as ametabolite of vitamin K in rats (Matschiner et al., 1970). The principal234

V I TA M I N K

metabolites of menadione are the sulfate and glucuronide of dihydromenadione (Losito et al., 1967). Some vitamin K is re-excreted into the intestine with bile, part of which is excreted in the feces. In humans, 20%
of injected phylloquinone was excreted in the urine, and 40 to 50% was
excreted in the feces via the bile (Shearer and Barkhan, 1973).

FUNCTIONS
Coagulation time of blood is the major function ascribed to vitamin
K. Four vitamin K-dependent proteins involved in blood coagulation
were discovered early in the investigations of the vitamin as a result ofhemorrhagic disease caused by deficiency. The vitamin is required forthe synthesis of the active form of prothrombin (factor II) and otherplasma clotting factors (VII, IX, and X). These four blood-clotting proteins are synthesized in the liver in inactive precursor forms (zymogens)and then converted to biologically active proteins by the action of vitamin K (Suttie and Jackson, 1977). In deficiency, administration of vitamin K brings about a prompt response and return toward normal of depressed coagulation factors in 4 to 5 hours. In the absence of the liver,this response does not occur.
The blood-clotting mechanism can apparently be stimulated by either an intrinsic system, in which all the factors are in the plasma, or an
extrinsic system. In the extrinsic system of coagulation, injury to the skinor other tissue frees tissue thromboplastin that, in the presence of various factors and calcium, changes prothrombin in the blood to thrombin.
The enzyme thrombin facilitates the conversion of the soluble fibrinogen
into an insoluble fibrin clot (Fig. 5.3). Fibrin polymerizes into strands
and enmeshes the formed elements of the blood, especially the red blood
cells, to form the blood clot (Griminger, 1984a). The final active component in both the intrinsic and extrinsic systems appears to activate theStuart-Prower factor, which in turn leads to activation of prothrombin.
The principal steps involved in blood clotting are presented in Fig. 5.3,
illustrating the need for vitamin K at four different sites in these reactions.
It is recognized that vitamin K-deficient animals synthesize vitamin
K-dependent proteins but in an inactive form, and vitamin K is needed
to convert these inactive protein precursors to biologically active proteins (Suttie, 1980). Investigations related to vitamin K mechanisms revealed that prothrombin contained 10 residues of a previously unrecognized amino acid, γ-carboxyglutamic acid (Gla). Likewise, the Gla
residues were found in the three other vitamin K-dependent proteins.
235

CHAPTER FIVE

Fig. 5.3 Scheme involving blood clotting. The vitamin K-dependent factors(synthesis of each is inhibited by dicumarol) include factor IX, Christmas factor; factor X, Stuart-Prower factor; factor VII, proconvertin; and factor II, prothrombin.

The amino-terminal regions of these proteins are homologous, and the
Gla residues are in essentially the same position in all of these clotting
factors (Suttie and Olson, 1990).
The metabolic function of vitamin K is as the coenzyme in the carboxylation of protein-incorporated glutamate residues to yield γ-carboxyglutamate (Fig. 5.4), thus converting inactive precursor proteins tobiological activity. Carboxylation allows prothrombin and the otherprocoagulant proteins to participate in a specific protein-calcium-phospholipid interaction that is necessary for their biological role (Suttie andJackson, 1977). To participate in this reaction, vitamin K is converted tohydroquinone and is then reconstituted to the quinone form via vitamin
K-epoxide.
Liver microsomes contain enzymes that oxidize the vitamin to its
236V I TA M I N K

Fig. 5.4 The vitamin K-dependent carboxylation reaction. The enzyme carboxylates specific glutamic acid residues of a limited number of proteins to γcarboxyglutamic acid residues.

2,3-epoxide and reduce the epoxide back to the reduced vitamin. The
carboxylase activity and epoxidase activity appear to share a common
oxygenated intermediate, and available data suggest that this may be ahydroperoxide of the vitamin. Evidence indicates that the role of vitamin
K is to labilize the γ-hydrogen of the substrate for CO2 attack rather
than to activate or transfer the CO2 (Suttie, 1980). Warfarin and otheranticoagulants of the coumarin type interfere with 2,3-epoxide reductase and therefore with the reconstitution of the active form of vitamin
K (see Chemical Structure, Properties, and Antagonists).
Four other vitamin K-dependent proteins have been identified inplasma (C, S, Z, and M). Proteins C and S play an anticoagulant rather
than a procoagulant role in normal hemostasis (Suttie, 1990; Combs,
1992). Protein C inhibits coagulation, and stimulated by protein S, it
promotes fibrinolysis. Also, a protein C-S complex can partially hydrolyze the activated factors V and VIII and thus inactivate them.
Individuals with inherited deficiency of factor C are at high risk for
thromboses. Protein S also has the potential to be involved in the regulation of bone turnover (Binkley and Suttie, 1995). Functions for proteins M and Z are unknown.
Continuing research has revealed that vitamin K-dependent reactions are present in most tissues and not just blood, and that a reasonably large number of proteins are subjected to this posttranslational car237

CHAPTER FIVE

boxylation of specific glutamate residues to γ-carboxyglutamate residues(Vermeer, 1986; Ferland, 1998). Gla-containing proteins have been purified from kidney, liver mitochondria, spermatozoa, urine, chickchorioallantoic membrane, and snake venoms. Atherocalcin is a vitamin
K-dependent protein in atherosclerotic tissue. A vitamin K-dependent
carboxylase system has been identified in skin, which may be related tocalcium metabolism in skin (de Boer-van den Berg et al., 1986). Two of
the best characterized vitamin K-dependent proteins not involved in hemostasis are osteocalcin or bone Gla protein (BGP) and matrix Gla protein, which were initially discovered in bone. Osteocalcin is a protein
containing three Gla residues that give the protein its mineral-binding
properties. Osteocalcin appears in embryonic chick bone and rat bonematrix at the beginning of mineralization of the bone (Gallop et al.,1980). It accounts for 15 to 20% of the non-collagen protein in the boneof most vertebrates and is one of the most abundant proteins in the
body. Osteocalcin is produced by osteoblasts, with synthesis controlled
by 1,25-dihydroxy vitamin D. Matrix Gla protein-deficient mice haveabnormal calcification leading to osteopenia, fractures, and premature
death due to arterial calcification (Booth and Mayer, 1997).
As is true for other non-blood vitamin K-dependent proteins, thephysiological role of osteocalcin remains largely unknown (Price, 1993).
However, reduced osteocalcin content of cortical bone (Vanderschueren
et al., 1990) and alteration of osteocalcin distribution within osteons(Ingram et al., 1994) are associated with aging. It remains unknown
whether any of these findings are related to the age-related increased riskof fracture. Osteocalcin may play a role in the control of bone remodeling because it has been reported to be a chemoattractant for monocytes,
the precursors of osteoclasts (Mundy and Poser, 1983). Serum osteocalcin has been shown to be a good predictor of bone turnover in pigs
(Carter et al., 1996). This suggests a possible role for osteocalcin in bone
resorption (Binkley and Suttie, 1995).
Several reports have indicated that warfarin treatment alters thefunctional properties of bone particles prepared from rats. However, vitamin K-deficient chick embryos were able to mobilize sufficient quantities of calcium for normal skeletal development, although they exhibited severe reduction in blood clotting and bone osteocalcin
concentration (Lavelle et al., 1994).
Observations that vitamin K could be involved in the pathogenesisof bone mineral loss have been summarized (Binkley and Suttie, 1995):(1) low blood vitamin K in patients with bone fractures; (2) concentration of circulating under-γ-carboxylated osteocalcin associated with age,238

V I TA M I N K

low bone-mineral density, and risk for hip fracture; (3) anticoagulant
therapy associated with decreased bone density; and (4) vitamin K supplementation, which decreases bone loss and calcium excretion.
Vitamin K has been suggested as a protective factor against gastricmucosal lesions, but the mechanism of protection is unclear (Yonaga,
1990). The protective effect of vitamin K against the development oftaurocholate-induced gastric lesions in mice increased with the dosage ofthe vitamin, but the vitamin’s protective effect varied according to thedifferences in its chemical structure. Phylloquinone was the most effective, followed by menaquinone and then menadione.
Vitamin A activity has been related to the vitamin K-dependent protein matrix Gla protein. The consistent induction of matrix Gla protein
in rat cultured cells and in animals administered retinoic acid suggestthat this vitamin K-dependent protein may mediate some of the known
actions of retinoic acid in differentiation and development (Cancela etal., 1993).
In animals, discoveries of active proteins that depend on vitamin K
for their synthesis are becoming common. Discovery of a vitamin K-dependent protein in skeletal tissue suggested that distribution of γ-carboxyglutamic acid was much more widespread than was once realized.
A number of pathological states result in an ectopic calcification of various tissues. Residues of Gla have been found in these lesions as well as
in calcium-containing kidney stones. Microsomes from kidney cortex
possess a vitamin K-dependent carboxylase activity. Proteins containing
Gla residues all interact with Ca2+ or other divalent cations, but their
physiological roles are not well understood. Although much is unknown
concerning vitamin K function apart from blood coagulation, it is apparent that a reasonably large number of proteins may be involved.

REQUIREMENTS
Vitamin K requirement of mammals is met by a combination of dietary intake and microbial biosynthesis in the gut, which may involve intestinal microorganisms (such as Escherichia coli) as well as ruminal microbes. Ruminal microorganisms in particular synthesize large amountsof vitamin K, explaining why ruminants do not appear to need a dietary
source of the vitamin. Animals that practice coprophagy, such as the
rabbit, can utilize much of the vitamin K that is eliminated in the feces.
In rats, most menaquinone absorbed results from fecal ingestion rather
than dietary sources, or from direct synthesis and absorption from theintestine (Kindberg et al., 1987). Animal feces contain substantial
239

CHAPTER FIVE

amounts of the vitamin even when none is present in feed. Despite theintestinal synthesis, animals can be rendered deficient when fed vitamin
K-free diets, and coprophagy is prevented if animals are maintained
germ-free, or if a vitamin K antagonist is given. Difficulties in demonstrating dietary requirement in many species include the varying degrees
to which they utilize vitamin K synthesized by intestinal bacteria and thedegree to which different species practice coprophagy. Contrary to mostanimals, poultry have a limited ability for intestinal synthesis, so adequate dietary supplies are of greater importance (see Deficiency).
Chicks were also found to contain less hepatic vitamin K epoxide reductase (Will et al., 1992). Activity of this enzyme in chicks was about
10% of that in rats fed the same diets; the inability of chicks to effectively recycle the epoxide of vitamin K (phylloquinone 2,3 epoxide)
seems to be the major factor in its high requirement for the vitamin.
Rats caged on litter obtain vitamin K, as well as B vitamins, by eating feces. Thus the practice was established of keeping experimental animals on wire screens to prevent this complication in vitamin feeding experiments. This practice was assumed to be successful until Barnes andcoworkers (1957) showed, using a new technique, that rats may eat 50
to 65% of their feces even when maintained on wire screens. In a later
experiment, in which rats were prevented by this new technique fromeating feces, vitamin K deficiency developed uniformly with a vitamin Kfree diet.
Because of microbial synthesis, a precise expression of vitamin K requirements is not feasible. However, attempts to determine the contribution of microbial synthesis have been made. In conventional rats, thevitamin requirement is 0.05 to 0.10 ppm, whereas in germ-free rats, therequirement is more than doubled, to about 0.25 ppm (Suttie and Olson,
1990). In humans, vitamin K homologs stored in liver indicate thatabout 40 to 50% of the daily requirement is derived from diet (mostly
vitamin K1), and the remainder from microbiological synthesis.
Estimated vitamin K requirements for various animals and humans
are presented in Table 5.1. The daily requirement for most species
ranges from 2 to 200 μg of vitamin K per kilogram of body weight. Thisrequirement can be altered by age, sex, strain, anti-vitamin K factors,
disease conditions, and any condition influencing lipid absorption or altering intestinal flora (see Deficiency and Supplementation sections).
Rapid rate of feed passage (e.g., diarrhea) through the digestive tract
may also influence vitamin K synthesis in animals. It is well known that
antibiotic intake will reduce synthesis of intestinal vitamin K. In mice, a
direct effect of antibiotic on vitamin K requirements has been shown.
240

V I TA M I N K■ Table 5.1

Vitamin K Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle
Chicken

Adult
Adult
Leghorn, 0–18 weeks
Laying (100-g intake)
Broilers
All classes
0–4 weeks
8–12 weeks
Breeding
Adult
All classes
Adult
Adult
Adult
Adult
Growing
Lake trout
All classes
Growing
Infants
Children
AdultsMicrobial synthesis
Microbial synthesis
0.5 mg/kg
0.5 mg/kg
0.5 mg/kg
4.0 mg/kg
1.75 mg/kg
1.0 mg/kg
1.0 mg/kg
Microbial synthesis
0.5 mg/kg
Microbial synthesis
Microbial synthesis
2 mg/kg
0.1 mg/kg
1 mg/kg
0.5–1 mg/kg
1.0 mg/kg
5 mg/kg
5–10 μg/day
15–30 μg/day
45–80 μg/day

NRC (1996)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1989b)
NRC (1981)
NRC (1977)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

Duck
Turkey
Sheep
Swine
Horse
Goat
Rabbit
Cat
Dog
Fish
Rat
Guinea pig
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry
matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as μg/day.

Even in the absence of intestinal flora, administration of an antibiotic
exacerbated vitamin K deficiency (Shirakawa et al., 1990).
Excess of vitamin A and calcium have been shown to influence vitamin K requirements. Rats fed excess retinol had two to three times the
carboxylase activity of endogenous prothrombin precursors, which is anindicator of vitamin K deficiency (Lee and Morre, 1992). Hall et al.
(1991) reported a hemorrhagic condition in pigs fed 2.7% dietary calcium. The condition was cured with vitamin K supplementation and was
not produced in treatment with less dietary calcium.
The adult human requirement for vitamin K is extremely low, andthere seems little possibility of a simple dietary deficiency developing in
the absence of complicating factors (Suttie, 1991). In humans, since placenta transfer of vitamin K is poor, intestinal flora are insufficiently developed during the first days after birth to provide the vitamin and, because of the low vitamin K content of the mother’s milk (e.g., 1 to 2μg/L), prothrombin level is very low in the newborn. Breast-fed infants
may ingest only 1 μg/day, which is only 20% of the requirement.
Recommended daily requirements for adults range from 45 to 80 μg
241CHAPTER FIVE

(RDA, 1989). Depletion studies that included a clotting assay, undercarboxylated prothrombin, and a decrease in urinary γ-carboxyglutamic
acid suggest vitamin K requirements for humans of about 1 μg/kg bodyweight daily (Suttie et al., 1988).

NATURAL SOURCES
There are two major natural sources of vitamin K: phylloquinones(vitamin K1) in plant sources and menaquinones (vitamin K2) producedby bacterial flora in animals. Vitamin K concentrations of various foods
and feedstuffs are presented in Table 5.2. Green leaves are the richest
natural source of vitamin K1. Light is important for its formation, andparts of plants that do not normally form chlorophyll contain little vitamin K. However, natural loss of chlorophyll, as in the fall yellowing ofleaves, does not bring about a corresponding change in vitamin K. In
most green leafy vegetables, phylloquinone increased during plant maturation, and cabbage was found to contain three to six times more of
the vitamin in the outer than the inner leaves (Ferland and Sadowski,
1992b).
Vitamin K is present in fresh, dark green vegetables and in an extract of pine needles. Cauliflower, broccoli, spinach, lettuce, and brussels■ Table 5.2

Vitamin K Concentration of Various Foods and Feedstuffs (asfed basis)

Food or Feedstuff
Alfalfa hay, sun cured
Alfalfa meal, dehydrated (20% protein)
Barley, grain
Cabbage (green)
Carrots
Corn, grain
Eggs
Fish meal, herring (mechanically extracted)
Liver (cattle)
Liver (swine)
Meat (lean)
Peas
Potatoes
Sorghum, grain
Soybean, protein concentrate (70.0% protein)
SpinachTomatoes
Sources: From NRC (1982b) and Marks (1975).

242

Vitamin K Level (ppm)
19.4
14.2
0.2
4.0
0.1
0.2
0.2
2.2
1–2
4–8
1–2
0.1–0.3
0.8
0.2
0.0
6.0
4.0

V I TA M I N K

sprouts are excellent sources. Edible oils are good sources of vitamin K1,
and human requirements of the vitamin are met when soybean or rapeseed oil exceed 15% of the caloric content (Ferland and Sadowski,
1992a).
Most feeds containing very high levels of vitamin K are not usually
fed to domestic animals, with the exception of alfalfa leaf meal, which
is sometimes included in small amounts in the diets of poultry and otheranimals. Cereals and oil cakes contain only small amounts of vitamin K,
while liver and fish meal are good animal sources of the vitamin. All
feeds or foods of animal origin, including fish meal and fish liver oils,
are much higher in vitamin K after they have undergone extensive bacterial putrefaction. Milk from cattle has been reported to be 10 times asrich as that of the human, and vitamin K content of cow’s milk is known
to vary significantly with the month of sampling. Both human colostrum(3.4 μg/L) and later produced milk (2.9 μg/L) were insufficient to meet
infant requirements (Canfield et al., 1991). The effect of food processing and cooking on vitamin K content has not been carefully considered;however, limited data suggest that food vitamin K is relatively stable,with the greatest loss resulting from oxidation. Irradiation of beef has
resulted in destruction of the vitamin. Ferland and Sadowski (1992a) reported that vitamin K1 content of edible vegetable oils was stable to processing, decreased slightly with heat, and was rapidly destroyed by bothdaylight and fluorescent light. Amber glass containers protected the oils
from the destructive effects of light.
The menaquinones (vitamin K2) are produced by the bacterial flora
in animals and are especially important in providing the vitamin K requirements of humans and most other mammals. However, the chickdoes not receive sufficient vitamin K from intestinal microbial synthesis(Scott et al., 1982). Type of diet, independent of vitamin K concentration, will influence total menaquinone synthesis. Rats fed a diet basedon boiled white rice had less menaquinone production and more severe
vitamin K deficiency than animals consuming diets based on autoclaved
black-eye beans (Mathers et al., 1990).
Vitamin K production in the rumen of ruminants and subsequentpassage along the small intestine, a region of active absorption, makesuch synthesized vitamin K highly available to the host. In nonruminants, the site of synthesis is in the lower gut, an area of poor absorption; thus availability to the host is limited unless the animal practices
coprophagy, in which case the synthesized vitamin K is highly available.

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CHAPTER FIVE

DEFICIENCY
The major clinical sign of vitamin K deficiency in all species is impairment of blood coagulation. Clinical signs include low prothrombin
levels, increased clotting time, and hemorrhaging. In its most severe form,
vitamin K deficiency will cause subcutaneous and internal hemorrhages,
which can be fatal. Deficiency can result from insufficient vitamin K in
the diet, lack of microbial synthesis within the gut, inadequate intestinal
absorption, or inability of the liver to use the available vitamin K.

Effects of Deficiency
Ruminants
Intestinal microorganisms provide vitamin K to preruminant animals. Nestor and Conrad (1990) studied vitamin K in preruminant veal
calves and concluded that intestinal microorganisms synthesized sufficient menaquinone-4 to meet needs of the vitamin prior to rumen development.
Microorganisms in the rumen synthesize large amounts of vitamin
K, and a deficiency is seen only in the presence of a metabolic antagonist, such as dicumarol from moldy sweet clover. Dicumarol is a fungus
metabolite produced from substrates in sweet clover hay. The coumarinderivatives are not active in the fresh plant because they are bound toglycosides, but are active when sweet clover is improperly cured(Vermeer, 1984). This condition, referred to as sweet clover poisoning or
hemorrhagic sweet clover disease, has been responsible for a large number of animal losses. Ruminants may die from hemorrhage following aminor injury, or even from apparently spontaneous bleeding. Dicumarol
passes through the placenta in pregnant animals, and newborn animals
may become affected immediately after birth.
All clinical signs of dicumarol poisoning relate to the hemorrhagescaused by failure of blood coagulation. The first appearance of clinicaldisease after consumption of spoiled sweet clover varies greatly and depends to a large extent on dicumarol content of the particular sweet
clover fed and animal age. If dietary dicumarol is low or variable, animals may consume it for months before signs of disease appear. In an experiment with calves, dicumarol poisoning was produced by feeding naturally spoiled, sweet clover hay that contained a minimum of 90 mg
dicumarol per kilogram of hay (Alstad et al., 1985). The minimum timerequired to develop clinical signs of vitamin K deficiency in these calves244

V I TA M I N K

was 3 weeks. In veterinary practice, another common cause of induced
vitamin K deficiency is the accidental poisoning of animals with warfarin (a synthetic coumarin used as a rodent poison). Initial clinical signs
may be stiffness and lameness from bleeding into muscles and articulations. Hematomas, epistaxis (nose bleed), or gastrointestinal bleeding
may be observed. Death may occur suddenly, with little preliminary evidence of disease, and is caused by spontaneous massive hemorrhage or
bleeding after injury, surgery, or parturition. DeHoogh (1989) reported
that two possible early embryonic deaths occurred and one cow aborted
from sweet clover poisoning.
Measurement of clotting time or prothrombin is considered a fairly
good measure of vitamin K deficiency. In experimentally induced
dicumarol poisoning (hemorrhagic sweet clover disease), Alstad et al.
(1985) reported that normal prothrombin time is equal to or less than
20 seconds. Deficiency of vitamin K was characterized by prothrombin
times longer than 40 to 60 seconds; with severe deficiency, prothrombin
time can be as long as 5 to 6 minutes.

Swine
Schendel and Johnson (1962) were able to produce a vitamin K deficiency in baby pigs by using a sulfa drug and an antibiotic and by carefully minimizing coprophagy by cleaning the feces from wire bottom
cases where the pigs were housed. Likewise, hemorrhagic disease occurred in piglets weaned at 5 to 6 weeks of age within 2 weeks after
transfer onto flat decks (Hoppe, 1987). This vitamin K-responsive outbreak had occurred because (1) the diet was virtually free of vitamin K,(2) sulfonamide, a known vitamin K-antagonist, was included in thediet, and (3) flat deck-housing was used, which precludes intake of microbial vitamin K from feces or litter.
Hall et al. (1991) accidentally produced vitamin K deficiency characterized by increased blood clotting times and high death losses when
feeding excess calcium (2.7%) without supplemental vitamin K. Either
supplemental vitamin K or reduced dietary Ca prevented the disease
condition.
Clinical and subclinical signs of vitamin K deficiency include increased prothrombin and blood-clotting time, internal hemorrhage, andanemia due to blood loss (Cunha 1977; Newsholme et al., 1985).
Newborn pigs may be pale with loss of blood from the umbilical cord.
Until recently, vitamin K deficiency under natural conditions was not expected, as it was thought that pigs synthesized most if not all of the vitamin that was required. However, in the late 1960s and early 1970s,
245CHAPTER FIVE

there were prevalent reports of a bleeding disease of young pigs on commercial diets that was successfully treated with vitamin K supplementation. Observations from Australia and New Zealand were of hemorrhaging in the navel of newborn pigs (Cunha, 1977).
A number of field trials in the United States have reported a hemorrhagic syndrome in growing pigs. In one study, hemorrhagic syndrome
occurred 9 days after pigs were fed a standard diet, while those receiving either 2.5% dehydrated alfalfa meal or supplemental vitamin K remained in good health (Fritschen et al., 1970). Gross visible signs for hemorrhagic syndrome were large subcutaneous hemorrhages, blood in
urine, and abnormal breathing. (Note: Field observation may not be thesame as experiments.) Additional clinical signs from field observation
are that some pigs will develop enlarged blood-filled joints and becomelame, whereas others may have swelling along the body wall that arefilled with unclotted blood. Hematomas (or blood swellings) in the ears
also occur (Cunha, 1977). Hemorrhagic conditions in the growing pig
have in some cases been associated with ingestion of molds, such as aspergillus or moldy materials, and have usually responded to vitamin K
therapy.
Exactly why vitamin K supplementation is needed is not definitely
known. Cunha (1977) and Scott et al. (1982) have summarized likely
reasons for vitamin K deficiency under field conditions:
1. As confinement feeding has increased, less pasture and alfalfa, bothof which are good sources of vitamin K, are used for production of higher
energy and more efficient diets. Likewise, there has been a trend toward theuse of solvent-extracted soybean meal and other oil seed meals, and higherquality (less putrefied) fish meals, which are lower in vitamin K than theoriginal expeller meals and somewhat putrid fish meals.
2. Hemorrhaging gastric ulcers, which occur frequently, may increase
vitamin K needs.
3. Mycotoxins produced by certain molds may be present in the feed.
4. Antimetabolite (antivitamin K) may be in the feed and thus increase
vitamin K needs.
5. Use of slatted floors lessens the opportunity for coprophagy since feces are an excellent source of vitamin K.
6. Use of sulfa drugs, antibiotics, and other medications may reduce intestinal synthesis of the vitamin.
7. Breed or strains of pigs may require more vitamin K. Increased litter
size and rate of gain may also increase the need for vitamin K.

246

V I TA M I N K

Poultry
Vitamin K deficiency causes a reduction in the prothrombin content
of the blood and, in the chick, may reduce the quantity in the plasma toless than 2% of normal. Since the prothrombin content of the blood of
normal, newly hatched chicks is only about 40% that of adult birds, very
young chicks are readily affected by a deficiency of vitamin K. A carryover from the parent hen to the chick has been demonstrated (Almquist,
1971). Therefore, breeder hen diets should be supplemented with vitamin
K to ensure good chick health. To this point, laying hens fed a diet deficient in vitamin K produce eggs low in the vitamin, and when the eggs
are incubated, the chicks produced have low reserves and prolonged
blood-clotting time. Adverse effects on blood clotting are not apparent
until after hatching, when hemorrhaging and mortality occur if trauma
occurs. As a consequence, the chicks may bleed to death from an injury
as slight as that caused by debeaking or wing banding (Fig. 5.5).
In very young chicks deficient in vitamin K, blood coagulation time
begins to increase after 5 to 10 days of age, with clinical signs occurring
most frequently in chicks 2 to 3 weeks after they begin consuming a vi-

Fig. 5.5 Generalized hemorrhage due to severe vitamin K deficiency in a young
chick. (Courtesy of M.S. Scott, Cornell University.)

247

CHAPTER FIVE

tamin K-deficient diet. Hemorrhages often occur in any part of the body,either spontaneously or as the result of an injury or bruise. Postmortem
examination usually reveals accumulations of blood in various parts ofthe body; sometimes there are petechial hemorrhages in the liver, and almost invariably there is erosion of the gizzard lining.
Even though inadequate dietary vitamin K alters bone osteocalcin,
signs associated with the skeletal system are not as apparent as bloodclotting problems. One study found that although blood clotting was
impaired and there was a reduction in bone γ-carboxyglutamic acid concentration, vitamin K deficiency did not functionally impair skeletal metabolism of laying hens and their progeny (Lavelle et al., 1994).
Borderline deficiencies of vitamin K often cause small hemorrhagic
blemishes on the breast, legs, and wings; in the abdominal cavity and on
the surface of the intestine (Fig. 5.6). Chicks show anemia that may be
due in part to loss of blood, but also to the development of hypoplastic
bone marrow. Even borderline deficiency of vitamin K is of economic
importance in broiler production because the hemorrhagic areas that occur in the legs or throughout the body may result in a high percentage
of condemnations during inspection at the processing plant (Scott et al.,1982). A condition manifested by numerous small hemorrhages scattered throughout all tissues has been reported frequently in the commercial broiler industry (Almquist, 1978).
A number of considerations influence the likelihood of vitamin K
deficiency in poultry, including dietary sources of the vitamin, level ofthe vitamin in the maternal diet, intestinal synthesis, coprophagy, presence of sulfa drugs and other nonnutrients in the diet, and disease conditions. Chicks with coccidiosis, a disease that causes severe damage
along the intestinal tract, may bleed excessively or fatally. When sulfaquinoxaline or certain other drugs are present in the feed or drinking
water, or when coccidiosis is being treated, up to 10 times as much supplementary vitamin K is needed as when these drugs are absent (Scott etal., 1982). Antimicrobial agents suppress intestinal bacteria that synthesize vitamin K, and in their presence, the bird may be entirely dependent
on dietary vitamin K (NRC, 1994). Arsanilic acid increases the need for
dietary vitamin K in both breeder and chick diets.
In poultry, little intestinal synthesis occurs because of the short digestive tract. The young chicken’s large intestine or colon, a major areaof bacterial activity, comprises less than 6% of the total length of the intestinal tract, while the figure for the adult of the same species is 7%(Griminger, 1984b). In other domestic animals, the relative length varies
from 13% for the dog to 28% for the rabbit. Also, poultry cannot uti248

V I TA M I N K

Fig. 5.6 Hemorrhagic blemishes in the muscle of a chicken fed a diet deficientin vitamin K. (Courtesy of M.L. Scott, Cornell University.)

lize the vitamin K synthesized by intestinal flora because the synthesis istaking place too close to the distal end of the intestinal tract to permit
significant absorption.
Rapid rate of food passage through the digestive tract may also influence vitamin K synthesis in poultry. First defecation in pigs, for a specific portion of diet, may occur about 15 hours after feeding, but mostof a given meal will be retained in the tract appreciably longer. A comparable period for chickens would be approximately 3 hours (Griminger
1984b).

HorsesThere are no conclusive reports in the literature relating to vitamin
K deficiency in the equine species (NRC, 1989b). Clinical responses ofsome “bleeders” to vitamin K therapy suggest that some horses may require additional supplies of vitamin K (Wakeman et al., 1975). However,
it is generally assumed that phylloquinone in pasture or in good-quality
hay and that synthesized by microorganisms of the cecum and colon are
in sufficient quantities to meet requirements.

Other Animal Species
As a result of adequate dietary supplies and intestinal synthesis, vitamin K deficiency has not been reported in the guinea pig, and the mink
249

CHAPTER FIVE

has been little studied. Other species, including trout and channel catfish, have been reported to have limited response to supplemental vitamin K, with requirements determined only for trout (NRC, 1993) andjuvenile marine shrimp (Shiau and Liu, 1994).

DOGS AND CATS
Vitamin K deficiency signs are only occasionally described for dogs;
however, accidental intake of dicumarol types of rat poison, such as
warfarin and diphenadione (vitamin K antagonist), will result in a hemorrhagic condition in dogs (Kerr, 1986; Mount and Kass, 1989). Clinicalsigns in dogs include paleness and evidence of slow but persistent bleeding from a number of sites, including gums, bowel, and several injection
punctures (Kerr, 1986). Vitamin K deficiency has been demonstrated in
adult dogs following diversion of bile from the intestine by means of acholecystonephrostomy (NRC, 1985a). Vitamin K absorption fromboth diet and intestinal bacterial synthesis was apparently reduced.
Some reports indicate that newborn pups suspected of vitamin K deficiency occasionally respond to vitamin K therapy. Vitamin K deficiency
is rare in cats. However, vitamin K antagonism attributable to ingestionof rodenticides containing warfarin or related compounds is a cause ofhemorrhaging in cats. Clinical signs of vitamin K deficiency could include hematomas in the elbows; hemorrhage in the conjunctiva; and extensive hemorrhage in and around the stifle joint, with necropsy revealing extensive hemorrhage in the bladder, sublumbar area, pelvic canal,and perineum (Maddison et al., 1990).

FISH
Information on intestinal vitamin K-synthesizing microflora in fish
is lacking. However, cultured sardines fed a menadione-supplemented
feed were rich in menaquinone-4, which indicates that menadione wasconverted to menaquinone-4 in the body. Vitamin K deficiency results inanemia and prolonged coagulation time in fish. Hemorrhages in channel
catfish fed a vitamin K-deficient diet have been reported, while otherstudies of this species have found no benefit with supplemental dietary
vitamin K (NRC, 1993). For trout, no growth or survival data for vitamin K deficiency have been reported. However, dietary sulfaguanidine,
as well as cold water temperatures, have caused prolonged blood coagulation times. In shrimp, weight gain and feed efficiency were significantly higher in those fed menadione-supplemented diets than in controls.

250

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FOXESOn farms where silver and blue foxes were born with subcutaneous
hemorrhages, enrichment of diets of pregnant females with vitamin K
was beneficial (NRC, 1982a).

LABORATORY ANIMALS
Vitamin K deficiency (manifested as prolonged blood-clotting timeand hemorrhage) deveoped within 2 to 3 weeks in rats fed low dietary
quantities of the vitamin; the process was accelerated by feeding sulfonamides or preventing coprophagy (NRC, 1995). Rats given a low-fiber
diet based on boiled white rice developed signs of severe vitamin K deficiency within 23 days (Mathers et al., 1990). In mice, supplemental vitamin K corrected vaginal hemorrhages and fetal resorptions caused byfeeding a vitamin K antagonist. Hypoprothrombinemia can be produced
in adult male hamsters either by feeding a vitamin K-deficient diet or bytreatment with vitamin K antagonists. Guinea pigs did not develop hemorrhages or abnormal clotting times when fed a purified diet without
vitamin K (NRC, 1995). Deficiency was not in evidence due to the active intestinal flora and coprophagic habits.

NONHUMAN PRIMATES
An increase in prothrombin time of rhesus monkeys was demonstrated when vitamin K-deficient diets (soy protein based) were fed (Hillet al., 1964). Evidently, the vitamin K requirement is small (NRC, 1978),but vitamin K deficiency bleeding disorders are a particular risk formonkeys whose intestinal tracts do not harbor bacteria that can synthesize vitamin K (newborn animals and those receiving long-term, broadspectrum antibiotic therapy).

RABBITS
When a vitamin K-deficient diet was fed to pregnant rabbits, the result was placental hemorrhage and abortion of young (NRC, 1977).

HumansPrimary vitamin K deficiency is uncommon in healthy adult humans
because of the widespread distribution of vitamin K in plant and animal
tissues and the microbial flora of the normal gut, which synthesize themenaquinones in amounts that may supply the bulk of the vitamin K requirement. Cases of acquired vitamin K deficiency do, however, occur inthe adult population and, although relatively rare, they present a signif251

CHAPTER FIVE

icant problem for some individuals. Hazell and Baloch (1970) observedthat a relatively high percentage of older hospitalized adults have hypoprothrombinemia that responds to orally administered vitamin K.
Deficiency of vitamin K may result from inadequate intake, absorption, or utilization, or as a result of drugs that interfere with its activity.
Deficiency can occur rapidly in patients with poor food intake who areon antibiotics, particularly in the postoperative period (Pineo et al.,1973).
The most common condition known to result in vitamin K deficiency occurs in patients who have a low dietary intake of vitamin K and
are taking antibiotics. Patients with restricted food intake or who are on
total parenteral nutrition and are taking antibiotics should be closely observed for vitamin K deficiency.
Vitamin K as a fat-soluble vitamin requires the presence of bile salts
for its absorption in the upper small intestine; thus deficiency can becaused by biliary obstruction. In sprue, idiopathic steatorrhea, ulcerative
colitis, and other conditions in which fats are not effectively absorbed,
bleeding due to deficiency of vitamin K may occur. Severe bleeding during or, more frequently, a day or two after an operation for the relief ofjaundice due to obstruction of the common bile duct was once a complication much feared by surgeons. Today it is recognized that this danger can be circumvented by giving vitamin K by injection prior to the operation.
Hemorrhagic disease of the human newborn can result from dietarylack of vitamin K. The condition can affect infants as old as 3 to 4
months. It is more common among breast-fed infants than those fed formulated diets. The disease involves intracranial hemorrhage and has
high rates of associated death and central nervous system damage.
Tulchinsky et al. (1993) reviewed hemorrhagic disease of the newborn
in New York State and found that in 65% of 34 deaths, vitamin K had
not been administered or was given only after onset of hemorrhage. As
a result of this review, prophylactic vitamin K was made a mandatory
newborn care procedure in the State Public Health Code.
Newborn infants represent a special case of vitamin K nutrition because (1) the placenta is a relatively poor organ for maternal-fetal transmission of lipids, and (2) the newborn baby has a sterile intestinal tract
and, furthermore, is apt to be fed foods relatively free from bacterial
contamination. Breast milk and clean cow’s milk are very poor sourcesof the vitamin. Historically, it was once a mystery why cleanliness attimes led to hemorrhage due to an apparent vitamin K deficiency in infants receiving breast milk. Hemorrhages did not occur in children who252

V I TA M I N K

received milk under less sanitary conditions because many bacteria arecapable of producing vitamin K. There is an increasing incidence of hemorrhagic disease in newborns due to maternal drug ingestion and rising popularity of breast-feeding.
Infants in the first week of life have less prothrombin in their bloodthan normal adults and have a prolonged prothrombin time. In normal
infants, plasma prothrombin concentration and that of other vitamin Kdependent factors may decrease to levels as low as 30% of adult levelsin the second and third days of life. Then, as food is taken and the gastrointestinal tract is colonized, these levels gradually climb to normal
adult values over a period of weeks (Anonymous, 1985). Mild vitamin
K deficiency is seen in all newborn infants and is particularly severe inpremature infants. It is due to inadequate liver synthesis and is aggravated by decreased dietary intake and inadequate production by intestinal bacterial flora. The tendency to correlate the general low levels ofprothrombin seen in all infants, and particularly in premature or lowbirth-weight infants, with an insufficient hepatic level of vitamin K is
only partially correct as part of the prothrombin deficiency is due to an
inability of the immature liver to synthesize sufficient clotting factors(Suzuki, 1979).
Breast-fed infants are at higher risk of hemorrhage than are formula-fed babies because human milk contains only 1 to 2 μg of vitamin
K per liter, whereas cow’s milk contains 5 to 17 μg/L (Haroon et al.,1982). Furthermore, breast milk is sterile and delays (through the flora
it encourages) colonization of the gut with vitamin K-synthesizing bacteria.

Assessment of Status
Measurement of blood-clotting time has been used to evaluate thebody status of vitamin K (see Analytical Procedures), and althoughblood-clotting time is a fairly good measure of vitamin K deficiency, amore accurate measure is obtained by determining the prothrombin
time. Prothrombin time is indicative of efficiency of conversion of prothrombin to thrombin and fibrinogen to fibrin. Prothrombin time in severely deficient chicks, for example, may be extended from normal, 17
to 20 seconds, to 5 to 6 minutes or longer.
Prolongation of the prothrombin time in the absence of liver disease
indicates vitamin K deficiency, with further clarification by specific vitamin K-dependent factor assays or by the rapid response to administration of vitamin K. There are a number of one- and two-stage modified
assays for prothrombin determination. Also, snake venom preparations
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(e.g., Oxyuranus echis) have been used to develop one-stage clotting assays for prothrombin (Carlisle et al., 1975).
Inadequate vitamin K results in lowered plasma concentrations of
the vitamin K-dependent clotting factors—prothrombin (factor II), factor VII, factor IX, and factor X. Measurement of the plasma concentration of one of the vitamin K-dependent clotting factors is a good way toassess vitamin K status.
Recent advances in methodology have made it possible to routinely
measure circulating phylloquinone as a method of evaluating vitamin K
status. Levels below 0.5 ng/ml have been associated with impaired clotting functions. Assessing adequacy by using the rather insensitive prothrombin time required a large decrease in vitamin K activity in order toconclude that an apparent deficiency existed.
The ability to immunochemically detect circulating γ-carboxyglutamic acid residues is a basis for evaluating vitamin K status (Suttie,
1991). Restricted vitamin K for humans results in a functional clotting
assay that detects undercarboxylated prothrombin in plasma and a decrease in urinary γ-carboxyglutamic acid (Suttie et al., 1988). Urinary
excretion of γ-carboxyglutamate reflects functional vitamin K status,
since γ-carboxyglutamate released by catabolism of proteins is neither
reutilized nor metabolized. An increase in the extent of γ-carboxylationof the vitamin K-dependent bone protein (osteocalcin) by administeringvitamin K to subjects with no other signs of vitamin K deficiency suggests that this might be one of the more sensitive criteria for determining vitamin K status (Bach, 1994; Binkley and Suttie, 1995). More sensitive methods than blood coagulation provide an opportunity tomonitor much milder forms of vitamin K deficiency.

SUPPLEMENTATIONAs long as natural dietary vitamin K sources (e.g., green leafy
plants) are sufficiently high and/or bacterial synthesis in the intestinal
tract remains functional, supplemental dietary vitamin K is not necessary. However, high sources of vitamin K, such as green leafy plants, are
not usually fed to nongrazing domestic animals. An exception is alfalfa
meal, which is sometimes included in small amounts in chicken andother animal feed. Therefore, a source of vitamin K needs to be added
to the diets of animals that are not getting fresh greens, or their driedequivalent, and are not synthesizing sufficient amounts of this vitaminin their gastrointestinal tract.
Scott et al. (1982) reported that natural ingredients used in poultry254

V I TA M I N K

diets a few years earlier had probably contained sufficient vitamin K,
while their current diets did not. The common feedstuffs used in past
years, such as high levels of alfalfa meal, high-fat soybean and otheroilseed meals, and putrefied fish meals, supplied ample vitamin K.
Trends toward (1) reducing levels of alfalfa for production of higher-energy, higher-efficiency diets; (2) solvent extraction of soybean meals andother oilseed meals; (3) improved processing of fish meals, resulting inlower menaquinone levels because of less putrefaction; and (4) use of vitamin K-inhibiting drugs in feed and drinking water have had a combined effect that necessitates supplementation of most poultry feeds.
Inadequate vitamin K under practical circumstances for poultry is most
likely to occur during the starting period, and supplementation of thefeed at this time is advantageous. Dietary changes affecting poultry vitamin K needs are similar for swine in confinement (see Deficiency); however, it is generally accepted that swine better utilize vitamin K from microbial synthesis.
Vitamin K antagonists will increase the vitamin K needs of livestock.
In adjusting dietary vitamin K fortification levels, an appropriate margin of safety is needed to prevent deficiency and low optimum performance in livestock. Vitamin K antagonists include the following:
1. Use of certain antibiotics and sulfa drugs. Sulfonamides and broadspectrum antibiotic drugs can virtually sterilize the lumen of the intestine.
Nelson and Norris (1961) showed that the inclusion of 0.1% sulfaquinoxaline increased the chick’s need for supplemental vitamin K by four to seventimes. By altering the ruminal and/or intestinal microflora, an excellent
source of vitamin K is lost.
2. Ingestion of an antimetabolite. Sweet clover contains high levels of
dicumarol, which can decrease prothrombin levels. Although known to bepotentially toxic, sweet clover hay is still fed to animals in certain parts of
the world (e.g., the Northern Plains states of the U.S.). Under these conditions, vitamin K supplementation would be warranted, as it would when
animals accidentally consume a dicumarol-based rat poison (e.g., warfarin). Warfarin, a synthetic antimetabolite, is a highly effective rat poison. Rats ingest large amounts of this extremely palatable antimetaboliteand eventually die from internal hemorrhaging. Marks (1975) observedthat the most common cause of vitamin K deficiency in veterinary practiceis the accidental poisoning of domestic animals with warfarin. Aflatoxin(a mycotoxin) is a toxic substance produced by molds. Vitamin K supplements may be helpful in correcting the deficiency associated with aflatoxicosis.
255

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The level of supplemental vitamin K should be adequate to meet therequirements under the wide variety of stress conditions encountered inpractical poultry and swine production. Squibb (1964) obtained increased prothrombin times, indicating a higher vitamin K requirement inchicks during the early stages of Newcastle disease. Studies have shown
an interrelationship between the severity of coccidiosis and vitamin K requirement and indicated that as much as 8 mg of vitamin K per kilogramof diet was needed at times for maximum growth and feed efficiency(Scott et al., 1982). Field reports with swine have indicated that hemorrhaging in stressed animals occurs at birth in the navel and following
castration. Various reports have indicated that levels of 2 to as high as
16 g of vitamin K per ton of feed were needed because the lower levels
were not effective under certain farm conditions (Cunha, 1977). Scott etal. (1982) concluded that coccidiosis possibly produces a triple stress on
the vitamin K requirement by (1) reducing feed intake and thereby the
supply of vitamin K, (2) injuring the intestinal tract and reducing absorption of the vitamin, and (3) treatment with sulfaquinoxaline orother coccidiostats that cause an increased requirement for vitamin K.
In vitamin K-deficient calves, normal prothrombin time wasachieved within 6 to 8 days after intramuscular injection with phylloquinone at 0.22 to 1.1 mg/kg body weight. Vitamin K1 dosages of 1.1,
2.2, and 3.3. mg/kg administered intramuscularly were effective in lowering prothrombin times to approximately normal values within 24
hours (Alstad et al., 1985). For confirmed cases of vitamin K deficiency,
vitamin K1 was shown to be much more potent than vitamin K3 in cattle (Alstad et al., 1985).
Vitamin K1 is not available to the feed industry because it is too expensive for this purpose; instead, water-soluble menadione (vitamin K3)
salts are used to provide vitamin K activity in feeds. Because of instability, menadione is not used in feed as the pure vitamin, but is formulated
as an additional product with sodium bisulfite and its derivatives. Watersoluble derivatives of menadione, including menadione sodium bisulfite(MSB), menadione sodium bisulfite complex (MSBC), and menadione
dimethyl-pyrimidinol bisulfite (MPB), are the principal forms of vitamin
K included in commercial diets. The greatest menadione activity is in
MSB (50%), followed by MPB (45.4%), and MSBC (33%) (Schneider
and Hoppe, 1986). Sometimes MSB is coated with gelatin to increasestability, resulting in 25% menadione activity.
Stability of the naturally occurring sources of vitamin K is poor.
However, stability of the water-soluble menadione salts is excellent in
multivitamin premixes unless trace minerals are present (Frye, 1978).
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Basic pH conditions also accelerate the destruction of menadione salts;
thus soluble or slightly soluble basic mineral substances should not be
included in multivitamin premixes containing menadione. Coelho(1991) concluded that vitamin K in the form of MSB or MSBC is very
sensitive to moisture and trace minerals, sensitive to light and gastricpH, and moderately sensitive to reduction and acid pH. Choline chloride is particularly destructive to vitamin K, with an average monthly
loss of 34 to 38% for MSBC and MPB when stored in a vitamin premix
with choline (Coelho, 1991). Less water-soluble forms or coated K3
forms exhibit superior stability compared to uncoated MSB. At higher
temperatures, uncoated MSB preparations lost about 60% activity(Gropp and Mehringer, 1990). Heat, moisture, and trace minerals increase the rate of destruction of menadione salts in both pressure-pelleted and extruded feeds (Hoffmann-La Roche, 1981). For these reasons, greater quantities of vitamin K3 are recommended in premixes that
contain large quantities of choline chloride and certain trace elements,and especially when plain MSB is used or premixes are exported orstored for an extended period (Schneider and Hoppe, 1986).
Clinical use of vitamin K in human nutrition is largely limited to
two forms: a water-soluble form of menadione (menadiol sodium
diphosphate) and the more expensive phylloquinone. Some danger ofhyperbilirubinemia has been associated with menadione usage (Suttie,
1991), whereas vitamin K in the form of phylloquinone is biologicallymore available. Griminger and Brubacher (1966) found four times asmuch vitamin K in eggs of hens fed phylloquinone as in eggs of hens fed
an equivalent level of menadione.
Vitamin K supplementation has been found to have a definite value
in human therapy (1) as a preoperative and postoperative measure toprevent risk of bleeding; (2) when absorption is impaired, as in obstructive jaundice, because bile is necessary for the absorption of vitamin K;
and (3) in newborns with hemorrhagic disease. Surgical patients should
be given supplemental vitamin K for loss of blood from operations, therapy with antibiotics, and fasting procedures. Phylloquinone is rapidly
depleted by fasting; it may be difficult to prevent vitamin K deficiency
by dietary phylloquinone alone during long-term fasting after surgery. In
one study, plasma phylloquinone decreased rapidly from 1.16 to 0.36
nmol/L in 11 patients during postoperative fasting (Usui et al., 1990).
Low-dose maternal vitamin K1 supplementation does not sufficiently increase vitamin K in breast milk (Greer et al., 1991; Pietschnig
et al., 1993). Recommendations from various locations suggest oral supplementation of infants at birth with 1 mg (Birkbeck, 1988; Cornelissen257

CHAPTER FIVEet al., 1993; Groot and De-Groot, 1993). In babies at risk, intramuscular injection of 500 μg at birth, followed by 25 μg by mouth daily for 3
weeks, is recommended (Birkbeck, 1988). In a study involving 100 infants, the authors concluded that vitamin K is equally effective whether
administered orally or intramuscularly (Malik et al., 1992). Intracranial
hemorrhage due to vitamin K deficiency in breast-fed infants is dramatically decreased with vitamin K supplementation in newborns.
Nevertheless, breast-fed infants with cholestasis or fat malabsorption remain at high risk of vitamin K deficiency and hemorrhage (Bancroft andCohen, 1993).

TOXICITY
Toxic effects of the vitamin K family are manifested mainly as hematological and circulatory disorders. Not only is species variation encountered, but profound differences are observed in the ability of thevarious vitamin K compounds to evoke a toxic response (Barash, 1978).
The natural forms of vitamin K, phylloquinone and menaquinone, arenontoxic at very high dosage levels. The synthetic menadione compounds, however, have shown toxic effects when given to humans, rabbits, dogs, and mice in excessive amounts. The toxic dietary level of
menadione is at least 1,000 times the dietary requirement (NRC, 1987).
Menadione compounds can be safely used at low levels to prevent thedevelopment of deficiency but should not be used as a pharmacologicaltreatment for a hemorrhagic condition.
The parenteral LD50 of menadione or its derivatives is a few hundred milligrams per kilogram of body weight in some species. Dosagesof 2 to 8 mg/kg body weight were reported to be lethal in horses(Rebhun et al., 1984), resulting in renal colic, hematuria, azotemia, andelectrolyte abnormalities consistent with acute renal failure. At
necropsy, lesions of renal tubular nephrosis were found. Also, 200 mg of
menadione sodium bisulfite administered to horses intravenously resulted in depression, stiffness, colic, laminitis, and renal fibrosis and failure (Maxie et al., 1992).
In animal studies, oral ingestion of large amounts of vitamin K1 (25g/kg body weight) produced no fatalities, whereas menadione had an
LD50 (in mice) equal to 500 mg/kg of diet (Molitor and Robinson,1940). Anemia, hemoglobinuria, urobilinuria, and urobilinoguria wereobserved with oral doses of 25 to 50 mg/kg. These doses are approximately 125 times the recommended clinical dose (0.05 mg/kg) for humans. The anemia seen following excessive vitamin K3 administration
258

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appears to be reversible following withdrawal (Richards and Shapiro,1945).
Full-term infants appear to have a greater tolerance to vitamin K
than premature infants. It is concluded that all vitamin K analogs aresafe when administered in the proper dosage; however, the formulation
with the greatest margin of safety is vitamin K1.

■ REFERENCES
Almquist, H.J. (1971). In The Vitamins, (W.H. Sebrell Jr., and R.S. Harris,eds.),Vol. 3, 2nd Ed., p. 418. Academic Press, New York.
Almquist, H.J. (1975). Am. J. Clin. Nutr. 28, 656.
Almquist, H.J. (1978). In Handbook Series in Nutrition and Foods, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 2, p. 195. CRC Press, Boca
Raton, Florida.
Almquist, H.J., and Stokstad, L.R. (1935). Nature (London) 136, 31.
Alstad, A.D., Casper, H.H., and Johnson, L.J. (1985). J. Am. Vet. Med. Assoc.
187, 729.
Anonymous. (1985) Nutr. Rev. 43, 303.
Bach, A.V. (1994). Sensitivity of Various Indicies of Human Vitamin K
Deficiency, M.S. Thesis, University of Wisconsin, Madison.
Bancroft, J., and Cohen, M.B. (1993). J. Pediatr. 16, 78.
Barash, P.G. (1978). In Handbook Series in Nutrition and Foods, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 1, p. 97. CRC Press, Boca
Raton, Florida.
Barnes, R.H., Fiala, G., McGehee, B., and Brown, A. (1957). J. Nutr. 63, 489.
Bell, R.G. (1978). Fed. Proc. Fed. Am. Soc. Exp. Biol. 37, 2599.
Binkley, N.C., and Suttie, J.W. (1995). J. Nutr. 125, 1812.
Birkbeck, J.A. (1988). New Zealand Med. J., 101, 421.
Booth, S.L., and Mayer, J. (1997). Nutr. Rev. 55, 282.
Cancela, M.L., Williamson, M.K., and Price, P.A. (1993). Nutr. Res. 13, 87.
Canfield, L.M., Hopkinson, J.M., Lima, A.F., Silva, B., and Garza, C. (1991).
Am. J. Clin. Nutr. 3, 730.
Carlisle, T.L., Shah, D.V., Schlegel, R., and Suttie, J.W. (1975). Proc. Soc. Exp.
Biol. Med. 148, 140.
Carter, S.D., Cromwell, G.L., Combs, T.R., Colombo, G., and Fanti, P. (1996).
J. Anim. Sci. 74, 2719.
Coelho, M.B. (1991). Vitamin stability in premixes and feeds: A practical approach. BASF Technical Symposium, Bloomington, Minnesota.
Combs Jr., G.F. (1992). The Vitamins. Academic Press, San Diego, California.
Cornelissen, E., Kollee, L., Abreu, R., Motohara, K., Monnens, L., and DeAbreu, R. (1993). Acta Paediatr. 82, 656.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, Inc., NewYork.
Dam, H., Schonheyder, F., and Tage-Hansen, E. (1936). Biochem. J. 30, 1075.
De Boer-van den Berg, M.A.G., Verstijnen, C.P.H.J., and Vermeer, C. (1986). J.
259

CHAPTER FIVE

Invest. Dermatol. 87, 377.
DeHoogh, W. (1989). Bovine Pract. 24, 173.
Ferland, G. (1998). Nutr. Rev. 56, 223.
Ferland, G., and Sadowski, J.A. (1992a). J. Agric. Food Chem. 40, 1869.
Ferland, G., and Sadowski, J.A. (1992b). J. Agric. Food Chem. 40, 1874.
Fritschen, R.D., Peo Jr., E.R., Lucas, L.E., and Grace, O.D. (1970). J. Anim. Sci.
31, 199(Abstr.).
Frye, T.M. (1978). Proc. Roche Vitam. Nutr. Update Meet., Arkansas Nutr.
Conf., Little Rock, p. 54. Hoffmann-La Roche, Nutley, New Jersey.
Gallop, P.M., Lian, J.B., and Hauschka, P.V. (1980). N. Engl. J. Med. 302, 1460.
Goplen, B.P., and Bell, J.M. (1967). Can. J. Anim. Sci. 47, 91.
Greer, F.R., Marshall, S., Cherry, J., and Suttie, J.W. (1991). Pediatrics 88, 751.
Griminger, P. (1984a). Feedstuffs, 56(38), 26.
Griminger, P. (1984b). Feedstuffs, 56(39), 24.
Griminger, P., and Brubacher, G. (1966). Poult. Sci. 45, 512.
Griminger, P., and Donis, O. (1960). J. Nutr. 70, 361.
Groot, C.J., and De-Groot, C.J. (1993). Voeding 54, 10.
Gropp, J., and Mehringer, W. (1990). Zeitschrift-fur Ernahrungswissenschaft
29, 219.
Hadler, M.R., and Shadbolt, R.S. (1975). Nature (London) 253, 275.
Hall, D.D., Cromwell, G.L., and Stahly, T.S. (1991). J. Anim. Sci. 69, 646.
Haroon, Y., Shearer, M.J., Rahim, S., Gunn, W.G., McEnery, G., and Barkhan,
P. (1982). J. Nutr. 112, 1105.
Hazell, K., and Baloch, K.H. (1970) Gerontol. Clin. 12, 10.
Hill, R.B., Schendel, H.E., Rama Rao, P.B., and Johnson, B.C. (1964). J. Nutr.
84, 259.
Hoffmann-La Roche. (1981). Rationale for Roche Recommended Vitamin
Fortification—Dogs and Cats. RCD 5963/1280. Hoffmann-La Roche, Inc.,
Nutley, New Jersey.
Hollander, D. (1973). Am. J. Physiol. 225, 360.
Hollander, D., and Truscott, T.C. (1974). J. Lab. Clin. Med. 83, 648.
Hoppe, P.P. (1987). Praktische Tierarzt 68, 32.
Ingram, R.T., Park, Y.K., Clarke, B.L., and Fitzpatrick, L.A. (1994). J. Clin.
Invest. 93, 989.
Jolly, D.W., Craig, C., and Nelson, T.E. (1977). Am. J. Physiol. 232, H12.
Kerr, M.C. (1986). Vet. Rec. 119(17), 435.
Kindberg, C., Suttie, J.W., Uchida, K., Hirauchi, K., and Nakao, H. (1987). J.
Nutr. 117, 1032.
Konishi, T., Baba, S., and Sone, H. (1973). Chem. Pharm. Bull. 21, 220.
Lavelle, P.A., Lloyd, A.P., Gay, C.V., and Leach Jr., R.M. (1994). J. Nutr. 124,
371.
Lee, L., and Morre, D.M. (1992). Korean J. Nutr. 25, 492.
Loosli, J.K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.),Academic Press, San Diego, California.
Losito, R., Owen, C.A., and Flock, E.V. (1967). Biochemistry 6, 62.
Maddison, J.E., Watson, A.D.J., Eade, I.G., and Exner, T. (1990). J. Am. Vet.
Med. Assoc. 197, 1495.
Malik, S., Udani, R.H., Bichile, S.K., Agrawal, R.M., Bahrainwala, A.T., and260

V I TA M I N K

Tilaye, S. (1992). Indian Pediatr. 29, 857.
Manz, U., and Maurer, R. (1982). Int. J. Vitam. Nutr. 52, 248.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
Martius, C., and Alvino, C. (1964). Biochem Z. 340, 316.
Matschiner, J.T., Bell, R.G., Amelotti, J.M., and Knauer, T.E. (1970). Biochim.
Biophys. Acta 201, 309.
Mathers, J.C., Fernandez, F., Hill, M.J., McCarthy, P.T., Shearer, M.J., andOxley, A. (1990). Br. J. Nutr. 63, 639.
Maxie, G., Van Dreumel, T., McMaster, D., and Baird, J. (1992). Can. Vet. J.
33, 756.
McCollum, E.V. (1957). In A History of Nutrition, The Riverside Press,
Cambridge, Massachusetts.
McFarlane, W.D., Graham, W.R., and Richardson, F. (1931). Biochem. J. 25,
358.
Molitor, H., and Robinson, H. (1940). Proc. Soc. Exp. Biol. Med. 43, 125.
Mount, M.E., and Kass, P.H. (1989). Am. J. Vet. Res. 59, 1704.
Mundy, G.R., and Poser, J.W. (1983). Calcif. Tissue Int. 35, 164.
Nelsestuen, G.L., Zytkovicz, T.H., and Howard, J.B. (1974). J. Biol. Chem. 249,
6347.
Nelson, T.S., and Norris, L.C. (1961). J. Nutr. 73, 135.
Nestor Jr., K.E., and Conrad, H.R. (1990). J. Dairy Sci. 73, 3291.
Newsholme, S.J., Cullen, J.S.C., Nel, P.W., and Reyers, F. (1985). J. S. Afr. Vet.
Assoc. 56, 101.
NRC. Nutrient Requirements of Domestic Animals. National Academy of
Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of SciencesNational Research Council, Washington, D.C.
Olson, R.E., and Suttie, J.W. (1978). Vitam. Horm. 35, 59.
Pietschnig, B., Haschle, F., Vanura, H., Shearer, M., Veitl, V., Kellner, S., andSchuster, E. (1993). Eur. J. Clin. Nutr. 47, 209.
261

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Pineo, G.F., Gallus, A.S., and Hirsh, J. (1973). J. Can. Med. Assoc. 109, 880.
Price, P.A. (1993). J. Clin. Invest. 91, 1268.
RDA. (1989). Recommended Dietary Allowances, 10th Ed. National Academyof Sciences-National Research Council, Washington, D.C.
Rebhun, W.C., Tennant, B.C., Dill, S.G., and King, J.M. (1984). J. Am. Vet.
Med. Assoc. 184, 1237.
Richards, R.K., and Shapiro, S. (1945). J. Pharmacol. Exp.Ther. 84, 93.
Schendel, H.E., and Johnson, B.C. (1962). J. Nutr. 76, 124.
Schneider, J., and Hoppe, P.P. (1986). Bioavailability of Nutrients in Feed
Ingredients, p. 1. National Feed Ingredients Association (NFIA), Des
Moines, Iowa.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Shearer, M.J., and Barkhan, P. (1973). Biochim. Biophys. Acta 297, 300.
Shearer, M.J., Barkhan, P., and Webster, G.R. (1970). Br. J. Haematol. 18, 297.
Shearer, M.J., Bach, A., and Kohlmeier, M. (1996). J. Nutr. 126, 1181S.
Shiau, S., and Liu, J. (1994). J. Nutr. 124, 277.
Shino, M. (1988). Analyst, 113, 393.
Shirakawa, H., Komai, M., and Kimura, S. (1990). Inter. J. Vit. Nutr. Res. 60,
245.
Squibb, R.L. (1964). Poult. Sci. 43, 1443.
Stenflo, J., Fernlund, P., Egan, W., and Roepstorff, P. (1974). Proc. Natl. Acad.
Sci. 71, 2730.
Suttie, J.W. (1980). Fed. Proc. Fed. Am. Soc. Exp. Biol. 39, 2730.
Suttie, J.W. (1990). Clin. Cardiol. 13, 16.
Suttie, J.W. (1991). In Handbook of Vitamins, 2nd Ed. (L.J. Machlin, ed.), p.
145. Marcel Dekker, New York.
Suttie, J.W., and Jackson, C.M. (1977). Physiol. Rev. 57, 1.
Suttie, J.W., and Olson, R.E. (1990). In Nutrition Reviews. Present Knowledge
in Nutrition (R.E. Olson, ed.), p. 122. Nutrition Foundation, Washington,
D.C.
Suttie, J.W., Mummah-Schendel, L.L., Shah, D.V., Lyle, B.J., and Greger, J.L.
(1988). Am. J. Clin. Nutr. 47, 475.
Suzuki, S. (1979). J. Perinat. Med. 7, 229.
Suzuki, S., Maki, M., Shirakawa, K., and Terao, T. (1989). J. Perinat. Med. 17,
305.
Thierry, M.J., Hermodson, M.A., and Suttie, J.W. (1970). Am. J. Physiol. 219,
854.
Thijssen, H.H.W., Drittij-Reijnders, M.J., and Fischer, M.A. (1996). J. Nutr.
126, 543.
Tulchinsky, T.J., Patton, M.M., Randolph, L.A., Meyer, M.R., and Linden, J.V.
(1993). Am. J. Publ. Health 83, 1166.
Usui, Y., Tanimura, H., Nishimura,N., Kobayashi, N., Okanoue, T., and Ozawa,K. (1990). Am. J. Clin. Nutr. 51, 846.
Vanderschueren, D., Gevers, G., Raymaekers, G., Devos, P., and Dequeker, J.
(1990). Calcif. Tissue Int. 46, 179.
Vermeer, C. (1984). Mol. Cell. Biochem. 61, 17.
Vermeer, C. (1986). Haemostasis 16, 239.
262

V I TA M I N K

Wakeman, D.L., Ott, E.A., Crawford, B.H., and Cunha, T.J. (1975). Horse
Production in Florida, p. 75. Bull.No. 188, Florida Dept. of Agricultural
and Consumer Services, Gainesville, Florida.
Weiser, H., and Tagwerker, F. (1982). Optimization of the Vitamin K Bioassayof Vitamin K Forms for Feed Supplementation, No. 1794. Hoffmann-La
Roche, Basel, Switzerland.
Will, B.H., and Suttie, J.W. (1992). J. Nutr. 122, 953.
Will, B.H., Usui, Y., and Suttie, J.W. (1992). J. Nutr. 122, 2354.
Yonaga, T. (1990). Nutr. Res. 10, 761.

263

chapter six

THIAMIN

INTRODUCTION
Thiamin (also called thiamine, aneurin[e], and vitamin B1) was thefirst vitamin to be discovered. Under most circumstances, there is little
chance of thiamin deficiency for monogastric animals, including humans, when diets contain ample quantities of whole cereal grains or
starchy roots. However, many thiamin antagonists in the food supply,and sensitivity of the vitamin to processing (e.g., heat), can lead to deficiency. Thiamin deficiency in humans has been a problem mostly in
Asian countries, where highly milled (polished) rice is consumed, thus
eliminating the thiamin-rich bran fraction of the grain. For years, it was
accepted that ruminants did not require vitamin B supplementation because of adequate rumen microflora synthesis. Intensification of ruminant feeding, involving high-concentrate diets, and management systems
with increased levels of production have resulted in nervous disordersthat are responsive to thiamin supplementation.

HISTORY
Thiamin is considered to be the oldest vitamin, and the diseaseberiberi probably is the earliest documented deficiency disorder. Beriberi
was prevalent in many world regions (particularly Asia) where diets
were based on highly milled or polished rice. The early history of thiamin has been reviewed (Jansen, 1956; Williams, 1961; Sebrell and Harris, 1973; Loosli, 1991). Beriberi was recognized in China as early as
2697 B.C. Beriberi occurred sporadically until the middle of the nineteenth century, but after about 1870, with the introduction of the steampowered rice mills, the incidence of the disease rose steeply, chiefly in
265

6CHAPTER SIX

prisons and barracks. In prisons in Java, a sentence of longer than 3months was equivalent to capital punishment, because after that timenearly all prisoners developed beriberi and died. Beriberi was prevalent
in armies and navies, particularly during times of war. In 1895, 26% of
the entire Japanese army of 17,500 had beriberi. Infantile beriberi was
the leading cause of death in some countries, with infants comprising 70
to 80% of all deaths from beriberi in the Philippines from 1912 to 1930.
No cure was found for beriberi until K. Takaki studied sailors in theJapanese Navy in the early 1880s, when the incidence of beriberi was
32%. By replacing some of the polished rice with other foods, Takaki
was able to dramatically reduce the incidence of beriberi. He incorrectly
thought that added dietary protein was responsible for preventing
beriberi.
In the 1890s Eijkman, a Dutch investigator, produced paralysis inchickens fed boiled polished rice. He called the condition polyneuritis
and observed that clinical signs were similar to beriberi symptoms in humans. In the course of experiments, Eijkman noticed by chance thatboth polyneuritis and beriberi could be both prevented and cured if ricebran was consumed with the polished rice. He mistakenly believed thatthe polished-rice diet produced a toxin that was counteracted by feeding
rice polishings. The conclusion that beriberi was caused by a vitally important food constituent was formulated in 1901 by Grijns, who was
Eijkman’s successor.
Ten years later (1911), Funk, of the Lister Institute in London, obtained a potent anti-beriberi substance from rice bran (thiamin) that had
the character of an amine. Thus the term vitamin(e), for vital amine, wascoined. However, it was later found that many vitamins are not amines.
Around the same time (1910), Captain Edward B. Vedder, who wasin the U.S. Army in Manila in the Philippines, became convinced thatberiberi, which was very prevalent, was indeed caused by a nutritional
deficiency; he began in earnest to pursue the isolation of the active factor. In 1912 Vedder and Chamberlain presented a paper on preventing
and treating beriberi with rice polishings. This practice saved many fromdeath due to infantile beriberi and was influential in establishing the deficiency theory for beriberi.
In 1860-61 the Burk and Wills expedition crossed the Australiancontinent from south to north. These explorers were the first Europeans
to make this trip, but both died on the return trip as they consumed the
Nardoo fern, which contained a high level of thiaminase activity (Earland McCleary, 1994). In 1926 two Dutch chemists, Jansen and Donath,successors to Eijkman and Grijns in the Indonesian laboratory, suc266

THIAMIN

ceeded in crystallizing thiamin in pure form. In 1936 Williams and colleagues, after many years of work, determined the chemical structure ofthiamin and were able to synthesize the vitamin. Earlier in his career
Williams had worked closely with Vedder in the Philippines toward thecontrol of beriberi through better nutrition.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Thiamin consists of a molecule of pyrimidine and a molecule of thiazole linked by a methylene bridge (Fig. 6.1); it contains both nitrogenand sulfur atoms. Thiamin is isolated in pure form as the white thiaminhydrochloride. The vitamin has a characteristic sulfurous odor andslightly bitter taste. Thiamin is very soluble in water, sparingly so in alcohol, and insoluble in fat solvents. It is very sensitive to alkali, in which
the thiazole ring opens at room temperature when pH is above 7. In a
dry state, thiamin is stable at 100oC for several hours, but moisturegreatly accelerates destruction, and thus, it is much less stable to heat infresh than in dry foods. Under ordinary conditions, thiamin hydrochloride is more hygroscopic (takes up moisture) than the mononitrate form.
However, both products should be kept in sealed containers. Autoclaving destroys thiamin, an observation that played an important role in the
discovery that what was originally considered to be a single vitamin was
actually a member of the vitamin B complex (Maynard et al., 1979).
Substances with an anti-thiamin activity are fairly common in nature and include structurally similar antagonists as well as structure-altering antagonists and thiamin-degrading enzymes (thiaminases). The
synthetic compounds pyrithiamine, oxythiamine, and amprolium (coccidiostat) are structurally similar antagonists, and their mode of action
is competitive inhibition with biologically inactive compounds, thus interfering with thiamin at different points in metabolism. Synthetic thiamin antagonists are often used in studies of the pharmacodynamics ofthiamin (Barclay and Gibson, 1982). Pyrithiamine blocks chiefly the esterification of thiamin with phosphoric acid, resulting in inhibition of
the thiamin coenzyme cocarboxylase. Oxythiamin likewise displaces cocarboxylase from important metabolic reactions. Amprolium inhibits
the absorption of thiamin from the intestine and blocks the phosphorylation of the vitamin.
Heat-stable thiamin antagonists occur in a number of plants (e.g.,ferns and tea); these include polyphenols (e.g., caffeic acid and tannicacid), which oxidize the thiazole ring to yield the nonabsorbable thiamin
267

CHAPTER SIX

Fig. 6.1 Structure of thiamin hydrochloride.

disulfide. Tall fescue (Festuca arundinacea Schreb.) toxicosis resembles
diseases caused by enhanced rumen thiaminase activity (Lauriault et al.,1990). Thiamin supplementation has been found to reduce tall fescue(endophyte-infected) toxicosis (Lauriault et al., 1990; Dougherty et al.,1991).
Sulfur has been shown to be antagonistic to thiamin enzymes. The
sulfite ion has been shown to cleave thiamin from enzymes at the methylene bridge between the pyrimidine and thiazole rings and analyticallywill mimic thiaminase. Sulfate increases thiamin-destroying activity inthe rumen contents, and the destructive mechanism involves thermolabile factor(s); however, the ruminal synthesis of thiamin is not affectedby sulfate (Olkowski et al., 1993).
Thiaminase activity destroys thiamin activity by altering the structure of the vitamin. The disease Chastek paralysis in foxes and other carnivores fed certain types of raw fish results from a thiaminase that splits
the thiamin molecule into two components, thus inactivating it. Twotypes of thiaminase enzymes have been described (Brent and Bartley,1984). Thiaminase II simply cleaves the vitamin at the methylene bridge
between the thiazole and pyrimidine rings. Thiaminase I substitutes a nitrogen-containing ring for the thiazole ring. This leads to less thiamin,
but it also gives rise to thiamin analogs composed of the pyrimidine ringof the original thiamin, and another ring from the “co-substrate.” This
created thiamin analog inhibits one or more thiamin-requiring reactions
necessary for energy metabolism in the central nervous system (CNS)(Frye et al., 1991).
Since thiaminase is heat labile, the problem can be avoided in fish,for example, by cooking the fish at 83oC for at least 5 minutes. Manykinds of fish contain thiaminase, and thiamin deficiency has been reported in penguins, seals, and dolphins fed primarily fish diets in zoos
(Maynard et al., 1979). Thiaminase is found mainly in herrings, sprats,
268

THIAMIN

stints, and various carp species—a total of some 50 species, most ofwhich live in fresh water. Wild aquatic animals apparently do not develop thiamin deficiency even though they eat a diet primarily of fish,
because fish must undergo some putrefaction to release the enzyme(Evans, 1975). In vitro and in vivo experiments have shown that 1 kg offish can destroy up to 25 mg of thiamin. This degradation takes place
within the first 30 minutes after ingestion, while still in the stomach.
Certain microorganisms (bacteria and molds) also have been shown to
produce thiaminases. A disease in horses known as bracken fern poisoning results from antagonism to thiamin.

ANALYTICAL PROCEDURES
Thiamin activity can be analyzed by biological, microbiological, andchemical methods. Biological methods are based on curative ability forpolyneuritis in pigeons; bradycardia in rats; or growth in chicks, pigeons, and rats. Microbiological tests are fairly rapid, inexpensive, andvery sensitive, but some organisms lack specificity for thiamin comparedto the pyrimidine and thiazole moieties or the enzyme form of the vitamin. Chemical analytical analysis for thiamin is conducted by oxidative
condensation of thiamin to thiochrome, which shows a characteristic
blue fluorescence in ultraviolet light. The fluorometric thiochrome
method is the procedure most widely used to estimate thiamin in foodstuffs and feeds. A promising method using high-performance liquid
chromatography (HPLC) for routine determination of thiamin in foods
has been developed (Ollilainen et al., 1993).

METABOLISM
Digestion, Absorption, and Transport
Thiamin is readily digested and released from natural sources. A
precondition for normal absorption of thiamin is sufficient productionof stomach hydrochloric acid. Phosphoric acid esters of thiamin are split
in the intestine. Free thiamin is soluble in water and easily absorbed. Absorption occurs in the small intestine, particularly in the jejunum. In
most species, absorption is negligible in the distal portion of the small
intestine, and in the stomach and large intestines. Thus, thiamin synthesized by gut microflora in the cecum or large intestine is largely unavailable to the animal except by coprophagy. The horse, however, canabsorb thiamin from the cecum. Ruminants can also absorb free thiamin269

CHAPTER SIX

from the rumen, but the rumen wall is not permeable for bound thiaminor for thiamin contained in rumen microorganisms.
The mechanism of thiamin absorption is not yet fully understood,
but apparently both active transport and simple (passive) diffusion are
involved (S’Klan and Trostler, 1977; Gubler, 1991). At low concentrations there is an active sodium-dependent transport against the electrochemical potential, whereas at high concentrations it diffuses passively
through the intestinal wall. Absorption from the rumen is also believed
to be an active mechanism.
Absorbed thiamin is transported via the portal vein to the liver witha carrier plasma protein, thiamin-binding protein (Rose, 1990). Thisbinding protein is hormonally regulated (e.g., corticosteroid hormones)
and is associated with thiamin transport into and out of the cell. In
blood, 90% of total thiamin is in the cellular fraction (predominantly
erythrocytes), while the plasma contains largely free unesterified thiamin. Thiamin is efficiently transferred to the fetus. In human placentas
thiamin concentration was found to be 4.5 mmol/L in maternal plasma
versus 45.9 mmol/L in venous cord plasma, indicating a massive retention by the fetus (Zempleni et al., 1992).

Phosphorylation
Thiamin phosphorylation can take place in most tissues, but does soparticularly in the liver. Four-fifths of thiamin in animals is phosphorylated in liver under the action of adenosine triphosphate (ATP) to form
the metabolically active enzyme form thiamin pyrophosphate (TPP; orthiamin diphosphate or cocarboxylase). Of total body thiamin, about
80% is TPP, about 10% is thiamin triphosphate (TTP), and the remainder is thiamin monophosphate (TMP) and free thiamin. Extracellular
fluids, including serum, milk, and cerebrospinal fluid, contain only free(unesterified) thiamin and TMP, suggesting that these forms cross cell
membranes while TPP and TTP cannot.

Storage and Excretion
In animal tissues, thiamin occurs mostly as phosphate esters. Although thiamin is readily absorbed and transported to cells throughout
the body, it is not stored to any great extent. Thiamin content in individual organs varies considerably, with the vitamin preferentially retained in organs with high metabolic activity. During deficiency, thiaminis retained in greatest quantities in important organs such as the heart,brain, liver, and kidneys. The principal storage organs are the liver andkidney; however, approximately one-half of total thiamin is present in
270

THIAMIN

muscle (Tanphaichair, 1976). Thiamin is one of the most poorly stored
vitamins. Mammals can exhaust their body stores within 1 to 2 weeks(Ensminger et al., 1983). Since thiamin is not stored in large amounts, it
is relatively easy to develop severe deficiency in a short time, even in
adult animals.
Thiamin intakes in excess of current needs are rapidly excreted. Thismeans that the body needs a regular supply and that unneeded intakes
are wasted. The pig is somewhat of an exception, however—for some
unknown reason its tissues contain several times as much thiamin asother species studied. Thus it has a store that can meet body needs on athiamin-deficient diet for as long as 2 months (Heinemann et al., 1946).
Absorbed thiamin is excreted in both urine and feces, with small
quantities excreted in sweat. Urinary thiamin is mainly free thiamin and
TMP. However, when subjects consume low-thiamin diets, the main urinary excretion is pyrimidine and thiazole, and further degradation ofthese substances has been identified (Brown, 1990). Fecal thiamin may
originate from feed, synthesis by microorganisms, or endogenous synthesis (e.g., via bile or excretion through the mucosa of the large intestine). When thiamin is administered in large doses, urinary excretion
first increases, then reaches a saturation level, and with additional thiamin the fecal concentration increases considerably (Bräunlich andZintzen, 1976).

FUNCTIONS
Decarboxylation of α-Keto Acids and Transketolase
ReactionsThiamin in all cells functions principally as the coenzyme cocarboxylase or TPP. Thiamin monophosphate is completely inactive, andany coenzyme activity previously attributed to the triphosphate undoubtedly results from partial conversion to diphosphate by hydrolysisof the terminal phosphate ester bond by thiamin triphosphatase (Gubler,1991).
The citric acid cycle (Krebs cycle or tricarboxylic acid cycle) is responsible for production of energy in the body. In this cycle, breakdown
products of carbohydrates, fats, and proteins are brought together for
further breakdown and for synthesis. The vitamins riboflavin, niacin,
and pantothenic acid, as well as thiamin, play roles in the cycle. Thiaminis the coenzyme for all enzymatic decarboxylations of α-keto acids. Thus
it functions in the oxidative decarboxylation of pyruvate to acetate,
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CHAPTER SIX

which in turn is combined with coenzyme A for entrance into the tricarboxylic cycle.
In mammals, thiamin is essential in two oxidative decarboxylation
reactions in the citric acid cycle that take place in cell mitochondria andone reaction in the cytoplasm (Fig. 6.2). These are essential reactions forutilization of carbohydrates to provide energy. Decarboxylation in the
citric acid cycle removes carbon dioxide, and the substrate is converted
into the compound having the next lower number of carbon atoms. In
the mitochondria, reactions pyruvate dehydrogenase forms acetyl CoA
from pyruvate and α-ketoglutarate dehydrogenase to convert α-ketoglutarate to succinyl CoA as follows:
Pyruvate ♦ acetyl-CoA + CO2(1)

α-Ketoglutaric acid ♦ succinyl-CoA + CO2(2)

It has been shown that decarboxylation of the three branched-chain
α-ketoacids derived from the deamination of leucine, isoleucine, and valine are also oxidatively decarboxylated by a multienzyme complex
analogous to those for pyruvic and α-ketoglutaric acids, but more specific for these branched-chain α-ketoacids (Gubler, 1991).
TPP is a coenzyme in the transketolase reaction that is part of thedirection oxidative pathway (pentose phosphate cycle) of glucose metabolism that occurs not in mitochondria but in the cell cytoplasm inliver, brain, adrenal cortex, and kidney, but not skeletal muscle. Transketolase catalyzes transfer of C2 fragments; hence with ribulose 5-phosphate as donor and ribose 5-phosphate as acceptor, sedoheptulose 7phosphate and triose phosphate are formed. This is the only mechanism
known for synthesis of ribose, which is needed for nucleotide formationof RNA and DNA synthesis and results in formation of nicotinamide
adenine dinucleotide phosphate (NADP), which is essential for reducing
intermediates from carbohydrate metabolism to form fatty acids. It canalso supply intermediate sugars for glycolysis.

Other Functions
Thiamin has a vital role in nerve function, although the mechanismof action is unclear. It has been postulated that thiamin, probably as
TPP, plays an essential role in nerve transmission, apart from its enzymatic role in the Krebs cycle and the pentose pathway. When nerves arestimulated, the levels of TPP and TTP decrease, and TMP and free thiamin are released into the surrounding medium. This is believed to in272

THIAMIN

Fig. 6.2 Thiamin as thiamin pyrophosphate (TPP) in the metabolism of carbohydrates. (Modified from Bräunlich and Zintzen, 1976.)

volve sodium and potassium transport at the membrane (Barchi, 1976).
Fatty acids and cholesterol are the major constituents of cell membranes, and effects on their synthesis would affect membrane integrity
and function. Thiamin deficiency in cultured glial cells impairs theirability to synthesize fatty acids and cholesterol. The defect is related toreduced formation of key lipogenic enzymes. These changes could be273

CHAPTER SIX

the basis of degenerative changes seen in glial cells in early thiamin deficiency.
Stimulation of nerves by either electrical or chemical means has beenfound to result in the release of thiamin (as free thiamin and themonophosphate ester) that is associated with the dephosphorylation of
its higher phosphate esters (Combs, 1992). The antagonist pyrithiamine
can displace thiamin from nervous tissue and change the electrical activity of the tissue. Possible mechanisms of action of thiamin in nervous
tissue include the following (Muralt, 1962; Cooper et al., 1963): (1) Thiamin is involved in the synthesis of acetylcholine, which transmits neural impulses; (2) thiamin participates in the passive transport of sodiumof excitable membranes, which is important for the transmission of impulses at the membrane of ganglionic cells; and (3) the reduction in theactivity of transketolase in the pentose phosphate pathway that follows
thiamin deficiency reduces the synthesis of fatty acids and the metabolism of energy in the nervous system.
Thiamin has been shown to have a role in insulin biosynthesis. Isolated pancreatic islets from thiamin-deficient rats secreted less insulin
than those from controls (Rathanaswami and Sundaresan, 1991). In thiamin-deficient rats the total pancreatic protein content was not alteredwhen compared with control rats, whereas the pancreatic insulin content was decreased.

REQUIREMENTS
Many factors influence requirements for thiamin (Sebrell and Harris, 1973). Thiamin requirements in some species are difficult to establish because of vitamin synthesis by microflora in ruminants and most
likely for all species in the lower intestine. More important than dietary
effects on synthesis is the general conclusion that most of the synthesis
apparently occurs too far down the intestinal tract for absorption. Intestinal synthesis, therefore, would be of greatest benefit to animals thatpractice coprophagy. However, in horses, there is considerable synthesisof thiamin in the digestive tract, with an estimated 25% of the free thiamin in the cecum absorbed (Linerode, 1966). In humans and most livestock species other than ruminants and horses, it is doubtful whether theamount of thiamin produced by intestinal synthesis and absorbed islarge enough to make a significant contribution to body needs.
In ruminants, total feed thiamin and that available from ruminal
synthesis have to be considered together, and the total need is yet to be
defined. Animals with functional rumens are generally considered to274

THIAMIN

have no dietary thiamin requirements because of microbial synthesis.
However, thiamin deficiency can be produced in lambs, calves, andother young ruminants that do not have a functional rumen. Thiamindeficiency can be produced as late as the sixth week of life, the preruminant period. Mixed rations consisting of milk, sources of high energy,
and hay stimulate growth of the villi in the rumen, but milk alone retards development of the rumen mucosa and thus thiamin synthesis.
Self-synthesis of thiamin becomes significant in calves only from thesixth week of life on, and this synthesis is still relatively weak at weaning after 5 months (Zintzen, 1974).
The self-synthesis of thiamin is subject to dietary composition in therumen; synthesis is significantly enhanced by carbohydrates such as molasses, which are easily soluble, and by supplies of nitrogen, such as
urea. When monogastric animals receive only marginal amounts of thiamin, they will develop deficiency signs earlier if their feed is also low inprotein (Abe, 1969). In chickens, the amount of intestinal synthesis
would depend on carbohydrate type present in the diet, synthesis being
favored by cooked (dextrinized) starches versus glucose or sucrose (Scottet al., 1982). Some microorganisms in the rumen or intestine are detrimental to thiamin in that they may consume thiamin or produce thiaminases, thus making the ingested thiamin unavailable to the animal.
Concentration of thiamin in the gastrointestinal tract (especially in therumen) is more uniform than it is in the feed. The self-synthesis of thiamin is therefore high when the vitamin level in feed is low, and lesswhen feed is rich in thiamin. Therefore, there must be some regulating
mechanism that governs the concentrations of the vitamin in the rumen
and maintains this within certain limits (Zintzen, 1974).
Diet composition can dramatically influence thiamin requirements.
Inadequate dietary protein in the diets of lactating cows may decrease
thiamin synthesis in the rumen and the amount of thiamin entering theduodenum (Breves et al., 1984). Since thiamin is specifically involved incarbohydrate metabolism (see Functions), level of dietary carbohydrate
relative to other energy-supplying components influences thiamin requirement. The need for thiamin increases as consumption of carbohydrates increases. When thiamin is deficient, body reserves become depleted more rapidly when animals are being maintained on feed rich incarbohydrates than when they are receiving a diet rich in fat and protein. The “thiamin-sparing” effect of fats and protein has long been
known. The survival time of thiamin-deficient pigs was increased by increasing fat levels to 28% of the diet (Ellis and Madsen, 1944). This indicated that the requirement for thiamin was decreased as the dietary en275

CHAPTER SIX

ergy from carbohydrate was replaced with higher levels of fat. If thechief source of energy for muscular work is carbohydrate, then thiaminrequirements will increase. If fat or other noncarbohydrate materials
serve as the energy source, the work will have less impact on the thiaminrequirement. Thiamin status is influenced by folacin; rats deficient in folacin had decreased absorption of low doses of thiamin (NRC, 1995).
Thiamin requirements are obviously higher if feeds contain raw materials (e.g., fish) or additives with anti-thiamin action (see Chemical
Structure, Properties, and Antagonists). Spoiled and moldy feeds may
contain such antagonists or thiaminases. Chicks kept on feed infectedwith Fusarium moniliforme developed polyneuritis that could be cured
with thiamin injections (Fritz et al., 1973). High dietary intakes of sulfur (Gooneratne et al., 1989) as well as substances in tall fescue (Festuca
arundinacea Schreb.) (Edwin et al., 1968; Lauriault et al., 1990) wereantagonistic to thiamin, resulting in higher requirements.
Disease conditions also result in increased thiamin requirements.
When dietary thiamin is marginal, typical signs of thiamin deficiency are
more likely to develop in infected animals than in normal animals. Endoparasites such as strongylids and coccidia compete with the host for
thiamin contained in food. It has been shown experimentally that infection with these coccidia results in considerable reduction in thiaminblood levels. Thiamin levels were directly correlated to infection severity (McManus and Judith, 1972). Likewise, conditions such as diarrhea
and malabsorption increased the requirement.
Size, genetic factors, and metabolic status affect thiamin requirements. Thiamin requirement is proportional to size. Conditions that increase the metabolic rate (e.g., rapid growth, fever, hyperthyroidism,
pregnancy, and lactation) increase thiamin requirements. Light poultry
breeds (Leghorn) seem to have higher thiamin requirements than heavy
breeds (Thornton and Schutze, 1960), and leghorn hens deposit morethiamin in eggs than do heavy hens. Need for thiamin increases during
gestation, during lactation, and in hyperthyroidism; during lactation, therat needs five times the normal level. As an animal ages, its need for thiamin increases because efficiency of vitamin utilization likely diminishes.
Table 6.1 summarizes thiamin requirements for various livestock
species and humans; a more complete listing is given in the appendix(Table A1). The recommended dietary allowance of 0.5 mg/100 kcal for
thiamin is nearly four times the amount at which signs of deficiency
were observed in humans and is well within the range observed to beconsistent with good health (RDA, 1989). A minimum intake of 1

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THIAMIN

mg/day is recommended for individuals consuming 2,000 kcal/day. On
a calorie basis, infant requirements are similar to those of adults.
In humans, additional factors that influence thiamin requirements
include alcohol and drug consumption, smoking, and inborn errors of
metabolism. Alcoholism results in less dietary thiamin intake as well as
decreased absorption and utilization of the vitamin (Leevy, 1982). Drugs
that cause nausea and lack of appetite, induce diuresis, or increase intestinal motility will reduce the availability of thiamin (Gubler, 1991).
Strauss and Scheer (1939) reported a marked reduction in thiamin excretion in heavy smokers. Patients with thiamin-responsive inborn errorsof metabolism respond to pharmacological doses of thiamin (Scriver,1973).

■ Table 6.1

Thiamin Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Leghorn, 0–12 weeks
Leghorn, 12–18 weeks
Laying (100-g intake)
Broilers, 0–8 weeks
All classes
All classes
Adult
Growing-finishing, 3–5 kg
Growing-finishing, 5–120 kg
Adult
Growing, maintenance,
pregnant, and lactation
Working
Adult
Growing
Growing
Adult
GrowingCatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
Children
AdultsMicrobial synthesis
6.5 ppm milk replacer
Microbial synthesis
1.0 mg/kg
0.8 mg/kg
0.7 mg/kg
1.8 mg/kg
2.0 mg/kg
2.0 mg/kg
Microbial synthesis
1.5 mg/kg
1.0 mg/kg
1.0 mg/kg

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)

3.0 mg/kg
5.0 mg/kg
Microbial synthesis
1.2 mg/kg
2.0 mg/kg
5.0 mg/kg
0.75 mg/kg
1.0 mg/kg
10–15 mg/kg
1–10 mg/kg
4.0 mg/kg
5.0 mg/kg
0.3–0.4 mg/day
0.7–1.0 mg/day
1.1–1.6 mg/day

NRC (1989b)
NRC (1989b)
NRC (1981)
NRC (1982a)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

ChickenJapanese quail
Turkey
Sheep
Swine
Horse
Goat
Mink
Fox
Cat
Dog
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry
matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg/day.

277

CHAPTER SIX

NATURAL SOURCES
In plant products, most thiamin occurs as free thiamin, while in animal products, phosphorylated forms (mostly TPP) predominate. Cereal
grains and their by-products, soybean meal, and peanut meal are relatively rich sources of thiamin. Most foods are relatively low in thiamin.
It is essentially absent in oils and fats and some highly refined foods,
such as sugars, and the content in green vegetables, fruits, and sea foods
is relatively low. Lean pork, liver, kidney, and egg yolk are animal products rich in thiamin. The content in lean pork can be doubled by increasing the thiamin intake of the pig (Heinemann et al., 1946). Thiamincontent of typical feedstuffs is shown in Table 6.2.
Brewer’s yeast is the richest known natural source of thiamin. Since
the vitamin is present primarily in the germ and seed coats, by-products
containing the latter are richer than the whole kernel, while highly
milled flour is very deficient. Whole rice may contain 5 ppm of thiamin,with much lower and higher concentrations for polished rice (0.3 ppm)
and rice bran (23 ppm), respectively (Marks, 1975). Wheat germ ranks
next to yeast in thiamin concentration. The level of thiamin in grain rises
as the level of protein rises; it depends on species, strain, and use of nitrogenous fertilizers (Aitken and Hankin, 1970; Zintzen, 1974). The
content in hays decreases as plants mature and is lower in cured than in
fresh products. Thiamin concentration is correlated with leafiness andgreenness as well as protein content. In general, good-quality hay is a
substantial source, and in a dry climate there is practically no loss instorage. Reddy and Pushpamma (1986) studied the effects of 1 year’s
storage and insect infestation on thiamin content of feeds. Thiaminlosses were high in different varieties of sorghum and pigeonpea (40 to70%) and lower in rice and chickpea (10 to 40%), with insect infestation causing further loss.
The presence of molds in feeds can result in substantial nutrient loss,
including thiamin (Cook, 1990). The content of thiamin was reduced
from 43 to 50% for two cultivars of wheat infested with Aspergillus
flavus compared to the uncontaminated sound wheat (Kao and Robinson, 1973). Moldy feed analyses showed thiamin content of less than 0.1
mg/kg, whereas the same feed not contaminated with Fusarium moniliforme had a thiamin content of 5.33 mg/kg. The antagonistic factor
could be destroyed by treatment with steam (Fritz et al., 1973).
Thiamin is susceptible to destruction by several factors involved withhandling and processing of foods. Milk is not a rich source of thiamin,
278

THIAMIN■ Table 6.2

Thiamin Content in Foods and Feedstuffs

Energy-Protein Feed Sources (Plant)

mg/kg

Protein Sources (Animal)

mg/kg

Alfalfa meal
Barley grain, dried
Beans
Brewer’s grains, dried
Brewer’s yeast, dried
Coconut meal, driedCorn (maize), yellow grain
Corn (maize), germ meal
Corn (maize), dried gluten meal
Cottonseed meal, solvent extracted
Distiller’s dried solubles
Linseed meal, expeller extracted
Millet, dried
Oat grain
Peanut mealPeas, dried
Potatoes, sweet
Potatoes, white
Rice, branRice, grain
Rice, polished
Rye, dried
Sesame meal
Sorghum grain
Sugarbeet pulp, dried
Sugarcane molasses
Wheat bran
Wheat grain
Wheat standard middlings, dried
Whey, dried

3.9
5.7
6.0
0.8
95.2
0.8
3.5
10.9
2.1
6.4
6.8
5.1
4.5
5.2
12.0
9.0
0.9
1.0
23.0
5.0
0.3
4.4
10.0
3.9
0.4
1.2
8.0
5.5
12.0
8.0

Blood meal, dried
ChickenEggs, whole
Fish meal, with solubles
Liver and glandular meal
Meat and bone meal
Milk, cow’s
Skim milk, dried

0.2
0.4
3.4
2.0
2.6
0.1
0.4
3.5

Sources: Modified from Bräunlich and Zintzen (1976), Marks (1975), and Scott etal. (1982).

with processing resulting in the following losses: pasteurization, 9 to
20%; sterilization, 30 to 50%; spray-drying, 10%; roller-drying, 15%;and condensing (canning), 40% (Gubler, 1991). Since thiamin is watersoluble as well as unstable to heat, large losses in certain cooking operations result (McDowell, 1985). There was up to 40% loss after 8 months’
storage of frozen meat. Roasting beef and pork and boiling vegetables resulted in 40 to 50% loss. Cooking with microwave leads to much smaller
losses. In conventionally roasted pork and chicken, 48 to 96% of thiaminwas retained, whereas microwave-treated samples showed retention as
high as 88 to 96% (Uherová et al., 1993). Parboiled rice was found tocontain more thiamin than regular rice at both the same levels of milling,
because parboiling paddy rice results in inward diffusion of water-soluble vitamins to the endosperm (Grewal and Sangha, 1990).
279

CHAPTER SIX

DEFICIENCY
The classic diseases, beriberi in humans and polyneuritis in birds,
represent a late stage of thiamin deficiency resulting from peripheral
neuritis, perhaps caused by accumulation of intermediates of carbohydrate metabolism. Comparing signs of thiamin deficiency in various
species, one sees that disorders affecting the CNS are the same in all
species. This is explained by the fact that in all animals, the brain covers its energy requirement chiefly by the degradation of glucose and istherefore dependent on biochemical reactions in which thiamin plays a
key role.
In addition to neurological disorders, the other main group of disorders involves cardiovascular damage. Clinical signs associated withheart function are slowing of heartbeat (bradycardia), enlargement of
the heart, and edema. Less specific clinical signs include gastrointestinal
problems, muscle weakness, easy fatigue, hyperirritability, and lack ofappetite (anorexia). Of all nutrients, thiamin deficiency has the mostmarked effect on appetite. Animals consuming a low-thiamin diet soon
show severe anorexia, lose all interest in food, and will not resume eating unless given thiamin. If the deficiency is severe, thiamin must beforce-fed or injected to induce animals to resume eating. Table 6.3 lists
clinical signs (and symptoms) associated with thiamin deficiency in various species. Animals developing thiamin deficiency most readily arechickens and pigeons; pigs, rats, mice, and rabbits are less readily depleted, probably because of endogenous thiamin production.

Effects of Deficiency
Ruminants
Young animals that do not have a fully developed rumen can be thiamin deficient, as shown in experiments with calves and lambs (McDonald, 1982). Clinical signs include apparent weakness that is usually
first exhibited by poor leg coordination, especially of the forelimbs, andby inability to rise and stand. The head is frequently retracted (Fig. 6.3)
along the shoulder, and the heart may develop arrhythmia. Specific signs
are usually accompanied by anorexia and severe diarrhea, followed bydehydration and death (NRC, 1996). Signs in the calves can be either
acute or chronic. In one study, acutely affected calves were anorectic,
had severe diarrhea, and died within 24 hours. These signs appeared after 2 to 4 weeks on a low-thiamin diet (Johnson et al., 1948).
280

■ Table 6.3

Clinical Signs Associated with Thiamin Deficiency in Various Species
Cardiovascular Signs

Nervous Disorders

Human
Loss of appetite, tiredness/apathy, lack of concentration, irritable, anxiety states, depression

General Signs

ECG changes, falling blood pressure, arrhythmia edema, hypotension, tachycardia

Muscular weakness, calf cramps,
weakened reflexes, loss of feeling,neuritic symptoms, coma

Dyspnea

Hen
Loss of appetite, loss of weight,
weakness

Bradycardia (from 300 down to90–100/min), cyanosis, edema

PigeonLoss of weight

Opisthotonos, leg weakness,muscular weakness, extension
spasms, apathy, paresis

Sudden death, gonad damage,gut atony, diarrhea, hypothermia

Heart failure, tachycardia

Crop voiding (vomiting)

Mouse
Loss of appetite, loss of weight,
weakness

Leg weakness, opisthotonos,
ataxia, paresis

Pathological changes in the myocardium

Hyperexcitability, ataxia,
circumduction, hemorrhages in
the mesencephalon, opisthotonos

Pathological changes in liver,testicles, skeletal muscle,
diarrhea, hypothermia

Bradycardia (from 500 down to250–300/min), slackened pulse
and reduced respiratory rate

Ataxia and cramps, paralysis
circumduction, nodding of thehead

Diarrhea, skin atony,hypothermia—

Paralysis, cramps, opisthotonos

Coma

Myocardial damage

Ataxia, ventral flexion of thehead

Slow pulse

Paresis, cramps, opisthotonos

Rat
Loss of appetite, disturbancesof growth, general weakness
Rabbit—Cat
Loss of appetite, loss of weight
Dog
Loss of appetite, loss of weight,
weakness

Other Signs

Gastrointestinal disorders,
vomiting, hypothermia281

■ Table 6.3

Continued

282

Cardiovascular Signs

Nervous Disorders

Other Signs

Fox
Loss of appetite, loss of weight

General Signs

Pathological changes in the heart

Mink
Loss of appetite, loss of weight

Polyneuritis, ataxia, irritability,
cramps, spastic paresis, paralysis

Groaning, pathological changes
in the liver—

Polyneuritis, ataxia, cramps, convulsions, spastic paresis, paralysis,
opisthotonos

Diarrhea, poor quality of pelt

Slow pulse, cyanosis, heart failure, tachycardia, dilatation of theheart, edema, hemorrhages in thegastrointestinal wall and myocardium—

Vomiting, dyspnea, diarrhea,
skin atony, hypothermia, sudden
death

Cardiac arrhythmia, tachycardia

Hyperesthesia, polyneuritis, ataxia,
convulsions, cramps, opisthotonos,
spastic paresis, paralysis

Diarrhea, dyspnea, lachrymation, gnashing of the teeth,
sometimes disorders of vision

Heart failure, tachycardia

Nervousness, ataxia, apathy, muscle tremors, cramps, coma

Diarrhea, constipation, dyspnea

Pig
Loss of appetite, loss of weight,
weakness, premature births,
high mortality among the young
Young Ruminant
Loss of appetite, loss of weight,
weakness
Horse
Loss of appetite, loss of weight,
lusterless coat

Source: Modified from Bräunlich and Zintzen (1976).

THIAMIN

Fig. 6.3 Sheep with thiamin deficiency. Characteristics of the condition arehead bent backward (opisthotonos), cramp-like muscle contractions, disturbance of balance, and aggressiveness. (Courtesy of Michel Hidiroglou, Animal
Research Center, Ottawa, Ontario, Canada.)

Because of extensive ruminal thiamin synthesis, the general conclusion is that ruminants possessing a normally functioning rumen have no
dietary thiamin requirement. Despite the fact that rumen microbes synthesize thiamin, and feeds—particularly whole grains—contain thiamin,
deficiencies do develop in ruminants. In a number of world areas, a thiamin-responsive disease known as polioencephalomalacia (PEM) occurs
sporadically in cattle, sheep, and goats. Other names for PEM in different regions include cerebrocortical necrosis, cerebral necrosis, and forage poisoning.
The term PEM refers to a laminar softening or degeneration of brain
gray matter (Brent and Bartley, 1984). However, it is commonly used todescribe the CNS condition in ruminants. The condition affects mainly
calves and young cattle between 4 months and 2 years old, and lambs,
young sheep, and goats between 2 and 7 months old. The incidence of
PEM is reported to be between 1 and 20%, and mortality may reach
100%. Clinical signs in mild cases include dullness, blindness, muscle
283

CHAPTER SIX

tremors (especially of the head), and opisthotonos. The condition ischaracterized by circling, head pressing, and convulsions, and in severecases, the animal collapses within 12 to 72 hours after onset of the disease (Fig. 6.4). Because of the circling, the condition is sometimes called“circling disease.” The ears drop, and in the final stages, the limbs andhead are extended. General twitching of the musculature of the ears andeyelids, waving of the head and neck, and grinding of the teeth withgroaning sometimes occur. Without treatment, death usually comes
within a few days. The main lesions in these animals are necrotic areas
in both cerebral hemispheres.
PEM may run a short, acute course or may occur in a milder form
and run a more protracted course. The condition, particularly in its mild
form, probably is often not diagnosed and may occur more frequently
than is recognized. Clinical signs of CNS disorders associated with PEM
are more readily recognized than nonspecific signs such as scouring, reduced growth, and anorexia. However, these signs are exhibited at laterstages of thiamin deficiency (Rammell and Hill, 1986). Thornber et al.
(1979) reported that animals on thiamin-deficient diets may not show
clinical signs of a CNS disorder for 3 to 5 weeks or longer, although depressed blood thiamin levels and other clinical signs may be observed.
Without treatment, mortality is about 50% of animals with the mild
form and possibly up to 100% in animals with acute PEM. The incidence and death rates are higher in young animals from 2 to 5 monthsof age than for animals older than 1 year.
High-sulfur diets are associated with thiamin deficiency and PEM(Gould, 1998). Gould et al. (1991) reported that steers with the highest
rumen fluid sulfide concentrations coincided with the onset of clinicalsigns of PEM. Several cases of PEM occurred when gypsum had beenused as a feed intake limiter. It would appear that the sulfate ion of gypsum, during its conversion to sulfide, must pass through sulfite, which
may destroy thiamin. The sulfite ion apparently will cleave thiamin at
the methylene bridge, mimicking thiaminase. In lambs, PEM was induced by administration of a sulfide solution; neurological clinical signs
included stupor, visual impairment, and seizures (McAllister et al.,1992).
Feedlot cattle that received 0.72% sulfate had 50% less gains thancontrols, and 3 of 20 head developed PEM (Sadler et al., 1983). Supplementation of sheep fed diets high in sulfur (0.63%) had clinical signsof PEM, which were prevented with supplemental thiamin (243 ppm)
(Olkowski et al., 1992). However, thiamin did not totally prevent development of microscopic brain lesions, which are characteristic of284

THIAMIN

A

B

C

Fig. 6.4 (A and B). An animal with polioencephalomalacia, a disease of thiamin deficiency. Feedlot cattle suffering from this condition show dullness andsometimes blindness, with a series of nervous disorders such as circling, headpressing, and convulsions. Six to eight hours after thiamin injection, the same
animal was able to stand (C). With continued thiamin treatment, in 3 to 5days, the animal returned to almost normal, with slight brain damage. (Courtesy of B. Bock, University of Florida.)285

CHAPTER SIX

PEM. In sheep, high sulfur intake was shown to have a detrimental effect on in vitro polymorphonuclear leukocyte function. This can meanthat ruminants in areas where large quantities of sulfur are taken in withwater and feed have compromised immune function due to lower copper and thiamin status and hence are at risk of increased susceptibility
to infections (Olkowski et al., 1990). Field studies (Olkowski et al.,1991) found an inverse correlation between water sulfate content andblood thiamin in beef cattle. Cattle receiving water with less than 200
ppm of sulfate had higher blood thiamin than cattle receiving water withover 1,000 ppm of sulfate. Feedlot nutritionists have reported cases of
PEM when high percentages of wet-milled corn by-products were fed.
Sulfuric acid is used to steep corn in the wet-milling process; much of thesulfur then remains with the by-product.
Livestock grazing endophyte-infected (Acremonium coenophiatum)tall fescue develop tall fescue toxicosis. Clinical signs of fescue toxicity
include reduced feed intake and growth, elevated body temperature, andergot-like clinical signs with necrosis in the extremities. Smith et al.
(1986) grazed cattle from April to September on high-endophyte fescue.
The cattle exhibited typical signs of summer fescue toxicosis (reduced
performance, elevated body temperature, and rough hair coat) compared to cattle grazing endophyte-free fescue. Following grazing, thecattle were fed corn:corn silage diets with and without supplemental thiamin. Dietary supplementation with 0.5 g of thiamin during the feedlot
phase improved the daily weight gain by 0.19 kg/day.
Tall fescue toxicosis resembles conditions caused by enhanced rumen thiaminase activity (e.g., PEM) that may be treated with thiamin(Edwin et al., 1968; Lauriault et al., 1990). Responses to supplemental
thiamin were found to be greater when cattle grazing endophyte-infected tall fescue were exposed to heat stress (Lauriault et al., 1990). Results suggest that oral thiamin supplementation may alleviate tall fescue
toxicosis of beef cattle during warm weather. A contrary earlier report(Fontenot et al., 1988) found that feeding thiamin to cattle grazing moderately endophyte-infected fescue had no beneficial effect.
Zintzen (1974) concluded that PEM can be definitely established if
the following four situations exist:
1. Case history: Animals have been maintained on high-energy feeds
rich in carbohydrates, and other animals on the same farm have, from time
to time, died after exhibiting CNS disorders.
2. Biochemical evidence: Blood pyruvate has steeply increased, and theactivity of erythrocyte-transketolase has been reduced.
286

THIAMIN

3. Diagnosis and therapy: Animals thought to have PEM react
promptly to treatment with thiamin provided they are treated in the earlystages of the disease.
4. Pathological changes: Necropsy shows typical pathological anatomical changes (bilateral cortical necrosis) in the brain.

Seasonal trends have been associated with PEM, which may be due
to increased metabolic demands of gestation, lactation, and growth. Additionally, feeding high-concentrate diets may induce PEM. In goats in
Bikaner, India, PEM was prevalent throughout the year, with the highest incidence in late winter (Lonkar et al., 1993). It was suggested thatscarcity of grazing may have led the animals to consume plants containing substances with thiaminase activity.
Polioencephalomalacia generally occurs in feedlot cattle, frequently
about 3 weeks after a ration change. Research suggests that PEM is associated with lactic acid acidosis, with both conditions related to adaptation to grain diets. Oltjen et al. (1962) reported that thiamin in the rumen is decreased by a reduction in rumen pH; a low ruminant pH ischaracteristic of cattle fed high-concentrate diets. Although little information is available on the direct addition of thiamin to finishing cattle
diets, Brethour (1972) reported that in two trials, a combination of thiamin and sodium carbonate supplement increased feed intake by 5%
and daily gain by 8%. In a third trial, thiamin administered alone gave
an intermediate response to calves immediately after weaning.
PEM has caused significant economic losses in tropical countries,
not only in feedlots where high-grain diets are provided, but also wherehigh levels of molasses are fed. When molasses is provided (ad libitum)
together with rations containing little crude fiber, a disease referred to as“molasses toxicity” or “molasses drunkenness” occurs. Clinical signs of
this condition closely resemble PEM, and studies completed in Cuba
have suggested that thiamin treatment, together with additionalroughage, may be an effective cure (Losada et al., 1971).
A number of experiments have shown that PEM is caused by naturally occurring thiamin antagonists, in connection with reduced thiamin
synthesis or destruction of the vitamin (see Chemical Structure, Properties, and Antagonists). Several researchers have reported that most fieldcases of PEM result from progressive thiamin deficiency, likely a resultof gut and ruminal bacterial thiaminases (Frye et al., 1991). Clinical reports of PEM have shown that under high-concentrate feeding systemsof beef cattle and lambs, thiaminase may become active in the rumen
and cause thiamin deficiency in animals with functional rumens (Edwin
287

CHAPTER SIX

Fig. 6.5 A wasting disease (secadera) of cattle in the llanos of Colombia. The
disease is characterized by emaciation in spite of the availability of good-quality forage. Secadera has been reported as thiamin deficiency because it has
been alleviated with thiamin injections. However, the condition has also been
controlled with a highly fortified complete mineral supplement. (L.R. McDowell, University of Florida.)

and Lewis, 1971). These thiaminases may be produced by microorganisms (bacteria and fungi) in contaminated feeds (Davies et al., 1968).
Thiaminase I has been shown to increase in the feces when dairy cows
were switched from a low- to high-concentrate diet (Soita and Brent,1993). Amprolium’s mode of action as a coccidiostat is apparently
through inhibition of thiamin phosphorylation. Loew and Dunlop(1972) found that high levels of amprolium (considerably above the levels needed to prevent coccidiosis) could produce the physical signs andthe histological lesions of PEM.
Both Clostridium sporogenes and Bacillus thiaminolyticus havebeen isolated from the rumen of PEM cases. Both organisms produce
thiaminase type I. Thiaminases are found in certain plant species and are
produced by microorganisms believed to be responsible for PEM. Thishas been a special problem in Australia, where PEM occurs under pasture conditions, apparently being derived from some of the fern species.
From Colombia, Mullenax (1983) reported a wasting disease288

THIAMIN

known as secadera (drying up) as thiamin deficiency because the condition had been alleviated within thiamin injections. Mullenax suggested
that a fungus associated with native forages contains a thiaminase. On
the contrary, Miles and McDowell (1983) reported that the wasting disease secadera (Fig. 6.5) can be successfully controlled with a highly fortified complete mineral supplement. It is possible that supplementationof this wasting disease is controlled by either thiamin or minerals
through different mechanisms (McDowell, 1985).

Swine
Thiamin deficiency in swine reveals itself particularly in decreased
appetite and body weight, vomiting, slow pulse, subnormal body temperature, nervous signs, postmortem heart changes, and sudden deathbecause of heart failure. Animals consuming a low-thiamin diet soon
show severe anorexia, lose all interest in food, and will not resume eating unless given thiamin. If the deficiency is severe, thiamin must beforce-fed or injected to induce animals to resume eating. Heinemann etal. (1946) reported that pigs can utilize stored thiamin over a long period, as 56 days were required for the pigs in the study to lose their appetites after being placed on a thiamin-deficient diet. In another study,
death occurred 74 days after pigs were placed on a thiamin-free but otherwise adequate diet (Loew, 1978). For young pigs, severe thiamin deficiency resulted in death at the age of 3 to 4 weeks.
First signs of thiamin deficiency in pigs are reduced feed consumption and vomiting, with a sharp reduction in weight gains (Miller et al.,1955). Functional and structural cardiac changes are the main findings
in experimentally deficient swine; in contrast to clinical reports, nervous
system lesions were not detected. Electrocardiographically demonstrable
changes in heart tissue were seen in enlarged hearts obtained from pigs
on thiamin-deficient diets (Fig. 6.6). Microscopically, it is possible torecognize inflammations and necrotic changes in the myocardial fibers.
Thiamin-deficient animals have elevated plasma pyruvate concentrations (Miller et al., 1955), since with deficiency of the vitamin there
is an accumulation of intermediates of carbohydrate metabolism. The
red blood cell enzyme transketolase is lowered in thiamin-deficient pigs,
with this enzyme used as an indicator of thiamin status (see Deficiency,
Assessment of Status).

Poultry
Poultry are more susceptible to neuromuscular effects of thiamin deficiency than most mammals. In chickens and turkeys, there is loss of289

CHAPTER SIX

Fig. 6.6 Enlarged heart on right is due to thiamin deficiency. Heart on left isfrom a similar pig fed the same diet plus thiamin. (Courtesy of T.J. Cunha andWashington State University.)

appetite, emaciation, impairment of digestion, general weakness,
opisthotonos or stargazing, and frequent convulsions, with polyneuritis
as a late or terminal extreme clinical sign. Deficient birds can rapidly detect and discriminate against feeds that do not provide the vitamin(Hughes and Wood-Gush, 1971) and are high in carbohydrate content(Thornton and Shutze, 1960).
Early signs of thiamin deficiency are lethargy and head tremors.
Chicks fed very low thiamin (0.4 ppm) survived for only 7 to 10 days,
apparently only a few days after the supply of thiamin in the yolk sac
was exhausted (Gries and Scott, 1972). Some chicks developed nervous
disorders, apathy, and tremor as early as the third or fourth day of life.
These signs increased in severity up to ataxia, inability to stand, andhigh-grade opisthotonos or twisting of the neck. Severity of the spasms
increased when the chicks were frightened. Chicks that showed thesehigh-grade nervous disorders died within a few hours. Cardiac abnormalities have also been reported in acutely thiamin-deficient chicks(Sturkie et al., 1954). Paralysis of the crop, manifested as delayed emptying, accompanied the general neuropathy of experimental thiamin deficiency in chicks (Naidoo, 1956). Deficiency in pigeons resulted in crop290

THIAMIN

voiding (vomiting).
In mature chickens, polyneuritis is observed approximately 3
weeks after they are fed a thiamin-deficient diet (Scott et al., 1982). As
the deficiency progresses, paralysis of the muscles occurs, beginningwith the flexors of the toes and progressing upward, affecting the extensor muscles of the legs, wings, and neck. The chicken sits on its
flexed legs and draws back the head in a stargazing (opisthotonos) position (Fig. 6.7). Retraction of the head is due to paralysis of the anterior neck muscles. At this stage, the chicken soon loses the ability tostand or sit upright and falls to the floor, where it may lie with thehead still retracted.
Acutely deficient pigeons developed vomiting, emaciation, leg
weakness, and opisthotonos, the last of which appeared between 7 and
12 days after beginning the thiamin-free diet (Swank, 1940). Chronic
deficiency due to a diet partially inadequate in thiamin resulted in leg
weakness but no opisthotonos. Evidence of cardiac failure was also
noted. The lesions produced in thiamin-deficient pigeons were reported to be identical to those found in Wernicke’s encephalopathy(polioencephalitis) in humans (Lofland et al., 1963). Mycotoxininduced polyneuritis in pheasants was eliminated within hours after in-

Fig. 6.7 Polyneuritis in a thiamin-deficient chick. Muscle paralysis causes extended legs and retraction of the head. (Courtesy of M.L. Scott, Cornell University.)
291

CHAPTER SIX

traperitoneal injection of thiamin (Cook, 1990). In thiamin-deficientturkeys, onset of anorexia was rapid, and by the fourth day, deficientbirds had significantly lower feed consumption (Remus and Firman,
1990).
In chickens with thiamin deficiency, body temperature drops to as
low as 36oC, and respiratory rate progressively decreases (Scott et al.,1982). There is adrenal gland hypertrophy that apparently results in tissue edema, particularly in the skin. Atrophy of genital organs also occurs in chickens affected with chronic thiamin deficiency, being more
pronounced in the testes than in the ovaries. The heart shows a slight degree of atrophy.
The hen transfers thiamin to the egg in proportion to dietary content (Polin et al., 1963). Inadequate thiamin to the breeder flock will result in high mortality of embryos prior to hatching and of chicks that
hatch with polyneuritis (Polin et al., 1962; Charles et al., 1972).

HorsesUnder normal circumstances the natural diet plus synthesis by microorganisms in the gut probably meet the thiamin needs of horses(NRC, 1989b). Thiamin synthesis in the intestinal tract of horses was inferred by Carroll et al. (1949), who found the following concentrationsof thiamin (mg/kg dry matter) in the diet and intestinal contents: diet,
1.1; duodenum, 0.5; ileum, 2.2; cecum, 7.1; anterior large colon, 17.8;
and anterior small colon, 7.8. Linerode (1966) estimated that 25% of
the free thiamin in the cecum is absorbed by the horse. However, absorption of microbially synthesized thiamin in the intestine may notmeet total needs.
Horses fed experimental diets low in thiamin exhibited anorexia,
loss of weight, incoordination (especially in the hind legs), lower blood
thiamin, elevated blood pyruvic acid, and dilated and hypertrophiedheart (Carroll et al., 1949). Incoordination and other nervous signs were
alleviated by feeding thiamin, indicating that this species requires a dietary source of this vitamin. A lack of thiamin causes reproductive failure in both sexes.
Because of generally adequate thiamin dietary supplies and intestinal synthesis, deficiency results under practical conditions only whenthiaminase-containing plants are provided (see Chemical Structure,
Properties, and Antagonists). Ingestion of sufficient quantities of the thiaminase-containing bracken fern Pteridium aquilinum was shown toreadily cause thiamin deficiency in horses (Kingsbury, 1964). The signs
in affected horses included weight loss, incoordination, lethargy, cardiac
292

THIAMIN

irregularities, muscular twitching and tremors, and prostration; death
usually followed convulsions. Anorexia was not apparent until othersigns became severe. Treatment with thiamin was successful if given before terminal stages. The horsetail (Equisetum arvense) and star thistle(Centaurea solstitialis) may act in the same way as bracken fern.
Diniz et al. (1984) observed incoordination, staggering, and muscular tremors in 27 mules 2 months after introduction to bracken-infested
pasture. Eight died, and the rest recovered after removal from the pasture and injection with 100 mg of thiamin. Generalized congestion, pulmonary edema, and serosal and mucosal hemorrhages were noted at
necropsy.

Other Animal Species
DOGS AND CATS
The main clinical signs for thiamin deficiency in dogs and cats areanorexia and nervousness, leading to spasms and finally to death (NRC,
1985a; Davidson, 1992). Thiamin deficiency in cats and dogs has been
associated with feeding meat preserved with sulfur dioxide (Studdertand Labuc, 1991).
Of all the domestic animals, the cat is most often reported to be clinically thiamin deficient. This might well be expected as domesticatedcats often consume fish, and thiaminase may be present in many cat
foods (NRC, 1986). However, these are usually heat-treated, which
would be expected to inactivate thiaminase. Most cases have been the
result of feeding cats diets that contained a large proportion of raw fish
(Hoffmann-La Roche, 2000). Experimental studies with cats have produced signs of thiamin deficiency within 23 to 40 days of consuming diets composed solely of raw carp or raw salt-water herring (Smith andProutt, 1944). Subcutaneous administration of thiamin to affected cats
resulted in recovery in all cases.
In thiamin-deficient cats, anorexia and sometimes vomiting occur
within 2 weeks of ingestion of a thiamin-deficient diet, followed by thesudden development of neurological disorders including abnormal posture, ataxia, and seizures, culminating in progressive weakness anddeath. Affected cats often show ventroflexion of the head when suspended by the rear legs, or somersaulting when the cat leaps (HoffmannLa Roche, 1998). Affected kittens also have dilated pupils. Postural abnormalities likely develop and may include a spastic gait, curling up
when lifted, or a head tilt. Seizures or abnormal behavior, dilated pupils,
stupor, or opisthotonos (spasms causing head and heels to bend back293

CHAPTER SIXward) may also be observed (Shell, 1995).
Depending on the severity of the case, deficiency signs in cats can bealleviated by administration of thiamin; however, hemorrhages that occur in the periventricular gray matter of the brain because of thiamin deficiency can permanently affect the animal. Cats that recovered from experimentally induced thiamin deficiency were found to have significantly
greater difficulty learning or remembering maze tasks (Ralston Purina,1987).
Thiamin deficiency in dogs results from animals consuming diets inwhich marginal thiamin concentrations were destroyed in food processing, or thiaminases were sufficiently high in the diet. A group of sleddogs fed a diet of frozen, uncooked carp developed clinical signs of thiamin deficiency after 6 months. The addition of oatmeal, a dry dog
food, and 100 mg of thiamin daily to the affected dogs resulted in complete recovery within 2 months (Houston and Hulland, 1988).
Pathological changes in dogs due to thiamin deficiency predominately involve the nervous system and heart. The pattern of pathological changes depends on the period of induction; acute deficiency tends
to involve the brain and produce severe neurological signs, whereas
chronic deficiency produces pathological changes in the heart and peripheral nerves. Dogs sometimes show cardiac hypertrophy (enlargement), with slowing of the heart rate and signs of congestive heart failure, including labored breathing and edema.

FISH
Thiamin deficiency signs in fish are similar to those observed in
mammals and birds. The principal signs are anorexia, arrest of growth,and neurological disturbances. Fish swim restlessly, twist around withspasmodic jerks of the body, and often collide with the walls of the tank(Bräunlich and Zintzen, 1976). The slightest disturbances trigger fright
reactions that may result in sudden death. The surface of the body andthe fins are discolored and the liver is pale. Neurological disorders suchas hyperirritability due to thiamin deficiency have been reported insalmonids, channel catfish, Japanese eel, and Japanese parrot fish (NRC,
1993).
Chinook salmon reared on a thiamin-inadequate diet exhibited poor
appetite, muscle atrophy, instability, and loss of equilibrium, followedby convulsions and death. Freshwater eels developed deficiency after
consuming thiaminase-containing clams (Hashimoto et al., 1970). The
main cause of thiamin deficiency in commercial fish production is dietary provision of raw fish.
294

THIAMIN

FOXES AND MINK
Thiamin deficiency is also a problem in pelt animal farming and is
manifested as Chastek paralysis. The disease occurs in mink and foxes
and is induced by feeding these animals certain types of raw fish (seeChemical Structure, Properties, and Antagonists). Clinical signs of thiamin deficiency in mink are reported as anorexia, weight loss, and emaciation, followed later by nervous disorders such as involuntary movements, unsteady gait, muscular spasms, and attacks of cramp or generalparalysis. When affected animals are touched, they exhibit extension
spasms in the hind limbs. The terminal phase is characterized byopisthotonos and unconsciousness, and animals die in a characteristic
contorted position. Histological examination shows brain hemorrhages
and lesions of the neural pathways (Bräunlich and Zintzen, 1976). In
foxes, clinical signs include anorexia and abnormal gait, followed by severe ataxia, inability to stand, hyperesthesia, constant moaning, andconvulsions (Long and Shaw, 1943).

LABORATORY ANIMALSThiamin deficiency can be induced readily in rats and produces abnormalities of the central and peripheral nervous system and the heartand results in poor reproductive performance (NRC, 1995). Anorexia
and weight loss are prominent. Deficient rats have ataxia, impaired“righting” reflex, and drowsiness, which were reversed by injection ofthiamin hydrochloride. Chronic thiamin deprivation leads to selective
neuropathological damage in the brain.
Mice deficient in thiamin had violent convulsions—especially whenheld a few seconds by the tail—cartwheel or circular movements, brain
hemorrhages, decreased food intake, poor growth, early mortality, silver-streaked muscle lesions, and testicular degeneration (NRC, 1995).
Exposure of thiamin-deficient mice to ethanol resulted in brain damage
that was more severe than either treatment alone (Phillips, 1987).
Reduced food intake and weight losses, followed by CNS disorders,occurred in young, thiamin-deficient guinea pigs. An unsteady gait appeared, with some retraction of the head as the condition progressed.
Death was reported within 4 weeks (Reid and Bieri, 1967; NRC, 1995).
In hamsters, low thiamin intake was associated with chronically deficient animals with depressed growth and more rapid induction of tumors after applications of a chemical carcinogen (Salley et al., 1962).

295

CHAPTER SIXNONHUMAN PRIMATES
The reported clinical signs of thiamin deficiency in rhesus monkeys
include weight loss, apathy, weakness, loss of reflexes, paralysis, incoordination, convulsions, prostration, heart failure, and death (Blank et al.,
1975; NRC, 1978). Focal necrosis of myocardial fibers is a relatively
constant finding and has been associated with abnormalities in the electrocardiogram (Rinehart and Greenberg, 1949).

RABBITS
Rabbits fed a thiamin-free diet, along with a thiamin antagonist(neopyrithiamin), developed ataxia, flaccid paralysis, convulsions, andcoma, followed by death (Reid et al., 1963; NRC, 1977). The rabbit differed from nearly all other species in not developing anorexia. Apparently, intestinal synthesis and coprophagy prevented the usual signs fromdeveloping.

CROCODILES
Suspected thiamin deficiency was reported in farmed saltwater crocodiles (Crocodylus porosus) in northern Australia (Jubb, 1992). Disease
conditions were noted in 4 of 11 clutches of crocodile hatchlings. Sudden loss of righting reflex was the outstanding feature of the disease. Affected hatchlings would be found floating or lying on their sides orbacks, listless, with jaws open and unable to right themselves (Fig. 6.8).

HumansClassic beriberi has been known since the earliest recorded times in
the Far East, where it is endemic because of prevalent consumption ofpolished rice. Beriberi was the primary health problem in Indonesia,
Malaysia, Japan, and the Philippines as recently as the 1940s. In 1947
the Philippine mortality rate from beriberi was 132 per 100,000 population—second only to tuberculosis. Through rice enrichment with thiamin, this rate was reduced to 14 per 100,000 by 1960.
In beriberi, both cardiac and nervous functions are usually disturbed. The disease occurs in a wet form, characterized by edema (Fig.
6.9) and cardiovascular symptoms, and a dry form, characterized by peripheral neuritis, paralysis, and muscular atrophy. However, the forms
merge, making it hard to differentiate the two. The chronic form maylast for years; cardiac symptoms may appear suddenly (shoshin beriberi)and result in death within a short time. It is generally believed that themore serious the nervous lesions, the greater the muscular pain and296

Fig. 6.8 A 2-month-old saltwater crocodile hatchling with suspected thiamindeficiency, floating on its side in shallow water. The listless appearance andopen jaws are also characteristic of the disease. (Courtesy of T.F. Jubb, Department of Western Australia, Kununurra.)

A

B

Fig. 6.9 Thiamin deficiency in a man. (A) Swelling of the legs, with pitting inankle region, marks beginning of so-called wet beriberi. (B) The same patient 4
days after a single intravenous injection of 50 mg of thiamin. During this period, the patient’s excretion of fluid exceeded intake by 10.5 lb. His general
nutritional state improved as a result of increased appetite. (Courtesy of Alan
T. Forrester, Vitamin Manual, The Upjohn Company, Kalamazoo, Michigan.)297

CHAPTER SIX

weakness and the less likely the development of acute beriberi. The heartis saved from extreme insufficiency by forcing the patient into complete
rest at an early stage in the attack.
General symptoms of beriberi include anorexia, heart enlargement,tachycardia (versus bradycardia in animals), lassitude and muscle weakness, paresthesia, loss of knee and ankle reflex with subsequent foot andwrist drop, ataxia due to muscle weakness, and dyspnea on exertion(Tang et al., 1989; Gubler, 1991). The “squat test” illustrates the neurological damage, as beriberi patients are unable to rise from a squatting
position.
The signs and symptoms vary depending on age, individual, diet,
duration and severity of the deficiency, and abruptness of onset. Peripheral neuropathy (dry beriberi) is a symmetrical impairment, with loss of
sensory, motor, and reflex function affecting the distal segments of limbs
more severely than the proximal ones, and with less cardiac involvement. Disturbances of the higher nervous centers, nystagmus, and ophthalmoplegia are frequent. In advanced stages, patients exhibit generalmuscular atrophy, ataxia, mental confusion, and defective short-term
memory.
The sequence of symptoms is variable, but beriberi often begins withnumbness in feet, heaviness of legs, prickly sensations, and itching. Muscles become tender, and squeezing the calf of the leg causes pain. Cardiac symptoms develop at some stage and are characterized by palpitation, epigastric pain, coldness of extremities, and enlargement of theheart. Later symptoms and clinical signs include edema of ankles, puffiness of face, digestive disturbances, anorexia, nausea, and vomiting.
Infantile beriberi, common in the Orient, is an acute form of the disease. It is probably an important cause of the high infant mortality inSoutheast Asia. Nursing mothers, who provide milk deficient in thiamin,
may or may not show mild signs of deficiency. The condition usually occurs in the first few months of life and begins with anorexia, regurgitation, abdominal distention, and colicky pain. Oliguria is followed by
edema, and there is dyspnea, with a peculiar cry or grunt thought to becaused by edema of the vocal cords. Later, cardiovascular signs and congestive failure increase and nervous signs appear, with muscular twitching, coma, and death. Cardiac failure is often the cause of death. Each
phase may last only a few hours, and the whole course of the condition
only a day or two.
Beriberi has been largely eliminated in most countries where fortification of rice is practiced, but is still prevalent in those countries where
unfortified polished rice is widely used. More recent incidences of298

THIAMIN

beriberi or low thiamin status have been reported in Gambia (Tang etal., 1989), Indonesia (Djoenaidi et al., 1992), Lithuania (Piktelite et al.,1992), Poland (Pardo et al., 1991), and Turkey (Wetherilt et al., 1992).
In countries where large amounts of fish are eaten raw, human thiamindeficiency may occur. Elsewhere, poverty, alcoholism, food faddism, or
poorly prepared food may result in inadequate thiamin ingestion. A variety of malabsorptive disorders can jeopardize nutritional status withrespect to thiamin. Severe polyneuropathy developed in one patient after jejunoileal bypass surgery (Glad et al., 1978), and thiamin blood status was found more depleted in patients with Crohn’s disease (Kuroki etal., 1993).
Studies have suggested that some degree of thiamin deficiency iswidespread in the elderly (Marks, 1975). This may result mostly fromgenerally poor appetite and eating habits or low income. However, it has
been demonstrated that old rats require more thiamin per gram of foodeaten than young rats, and transport of thiamin across the intestine issignificantly lower in older than younger rats (Gubler, 1991).
Blass and Gibson (1978) have reported decreased levels of pyruvate
dehydrogenase activity in the brain of patients with Alzheimer’s disease.
Tucker et al. (1990) found a decrement in alpha-wave activity in theelectroencephalographic indices of older persons with low thiamin status, suggesting subtle neuropsychological impairment with mild thiamindeficiency.
The most frequently encountered type of thiamin deficiency in developed countries is associated with alcoholism (Wernicke’s syndrome),
resulting from inadequate intake of thiamin. Thiamin deficiency inchronic alcoholism has a triple origin: (1) inadequate intake resulting
from appetite loss, or decreased intake of a balanced diet for economic
and/or other reasons; (2) decreased absorption; and (3) decreased utilization. The signs of this syndrome range from mild confusion to coma
and include ophthalmoplegia, cerebellar ataxia, psychosis, confabulation and severely impaired retentive memory, and cognitive function.
Blass and Gibson (1977) showed that at least some patients with Wernicke’s syndrome have a genetically determined abnormal transketolase
that, in effect, requires a higher concentration of the coenzyme to saturate the apoenzyme.
Some Japanese studies (Zintzen, 1974) have indicated that the structure-altering antagonists of thiamin (see Chemical Structure, Properties,
and Antagonists) produced by bacteria and fungi are main causes ofberiberi, since they are involved in up to 70% of all cases. Bacillary andclostridial species that produce these antagonists have been isolated
299

CHAPTER SIX

from intestinal flora of Japanese people tested. They may be consumed
with contaminated food, such as moldy rice.

Assessment of Status
In thiamin deficiency determination, there is no question about the
specificity of such signs as polyneuritis and Chastek paralysis, but these
are terminal stages. The problem of identifying the early stages by any
specific sign is an unsolved one for thiamin and for several other of theB vitamins. When thiamin deficiency is acute, the specific effects of the
deficiency cannot be distinguished from some unspecific effects, such as
those of anorexia (Zintzen, 1974). In humans, thiamin deficiency can betentatively diagnosed on a clinical basis, particularly with careful inquiry into the patient’s dietary habits or other conditions that might predispose to deficiency, and confirmed by positive results of thiamin supplementation.
Some specific criteria that are suitable for the assessment of the thiamin status include (1) concentration of thiamin in blood and urine, (2)levels of products of intermediate metabolism that are dependent on the
enzyme function of thiamin, and (3) activities of enzymes in which thiamin is a coenzyme.
Thiamin concentrations are decreased in blood and urine when thisvitamin is deficient. Clinical signs of deficiency were noted when lessthan 7% of a 1-mg dose of thiamin was excreted in the urine in a doseretention test (Horwitt et al., 1948). Urinary excretion of thiamin reflects thiamin saturation or depletion because the vitamin is excreted
promptly when ingested in excess of needs. Both blood levels and urinary excretion of thiamin are only a reflection of the immediately preceding intake and other factors and may not be a reliable index of tissue
stores, distribution, or biochemical functioning and hence are of limited
value for interpretation of actual thiamin status.
Thiamin is needed for pyruvate metabolism, and with deficiency, abnormally high concentrations of pyruvic and lactic acids in the blood indicate thiamin inadequacy. For a calf on a deficient diet, urinary excretion of thiamin drops to very low levels in 20 to 25 days, and increased
pyruvate excretion follows. Blood pyruvate and lactate levels increase
suddenly to 400 and 500% of normal as the deficiency develops (NRC,
1989a). However, the measurements of these levels often cannot be used
to detect mild thiamin deficiency. Also these tests are not entirely specific, because toxicity by minerals such as arsenic and antimony will also
inhibit the utilization of pyruvate. Increased pyruvate can also result
from a number of pathological conditions, such as those arising from in300

THIAMIN

creased adrenal gland activity. Acute polyneuritis has been shown to develop before rise of blood pyruvate in pigeons.
So far, the best criterion for determining thiamin status is activity ofenzymes that depend on the vitamin as a coenzyme. These enzymes include pyruvate decarboxylase, α-ketoglutarate decarboxylase, andtransketolase. Transketolase activity measurement is the most convenient, feasible, specific, and sensitive of tests for thiamin deficiency. Blood(particularly the red cell) transketolase activity is a reliable index of theavailability of coenzyme TPP, and thus correlated well with the degreeof deficiency in both animals and humans (Brin, 1962; Takeuchi et al.,
1990; Lonkar et al., 1993; Rains et al., 1997). Transketolase is an excellent indicator in that it is useful in detecting marginal thiamin deficiency.
Diagnosis of thiamin deficiency in ruminants initially depended
upon recognition of the clinical signs in live animals, followed by confirmatory brain histopathology or clinical response to thiamin administration (Rammell and Hill, 1986). Moreover, sick animals react so
promptly to treatment with thiamin (sometimes within hours) that early
treatment with thiamin is even used for confirming the diagnosis of
PEM. The best transketolase assay for assessing thiamin deficiency isbased on the so-called TPP effect, which is the percent of increase intransketolase activity following addition of excess TPP to the sample.
Values of 120 to 250% have been reported for animals diagnosed with
PEM (Edwin et al., 1979). It has been suggested that fecal thiaminase I
is an indicator of thiaminase presence in the digestive tract, and high levels of this fecal enzyme may be correlated to PEM (Soita and Brent,1993).

SUPPLEMENTATION
Thiamin is found in most feedstuffs, but in widely differing concentrations. Good thiamin sources are cereals, milling by-products, oil extraction residues, and yeast (see Natural Sources). The thiamin content
of most common feeds should be three to four times greater than the required amounts for most species (Brent, 1985). For swine and poultry
consuming typical diets (e.g., corn-soybean meal), thiamin is one of thevitamins least likely to be deficient. Likewise, under normal feeding andmanagement conditions, and in the absence of antimetabolites, thiamindeficiency should theoretically not occur in either young or adult ruminants.
Drying and processing can lower the concentrations of available thi301

CHAPTER SIX

amin in feedstuffs because thiamin is heat labile. For example, it was reported that use of high-moisture barley treated with sulfur dioxide resulted in destruction of 61% of dietary thiamin (Gibson et al., 1987).
Treatment of feed ingredients with sulfur dioxide inactivates thiamin.
This process was used to produce deficient diets in early studies to determine the pig’s thiamin requirement (NRC, 1998).
Utilization of available thiamin in feedstuffs may be limited and may
be impaired by thiamin antagonists and processing methods (e.g., heat);
therefore, it is common practice to add supplemental thiamin to poultry
and pig feeds principally as low-cost insurance. Thiamin supplementation should normally be considered not for grazing ruminants but rather
for animals that might develop PEM as a result of consuming high-concentrate diets. Grazing ruminants would not normally receive supplemental thiamin unless evidence is provided that consumed pastures contain antagonists. An example would be thiamin incorporated into dietsfor livestock grazing potentially toxic tall fescue pastures during midsummer when toxicosis is likely to be severe. Thiamin may be supplied
to cattle at the daily rate of 1 g per head (Lauriault et al., 1990). Johnson and Krautmann (1989) reported that 500 mg of thiamin per headper day, for the first 30 days that cattle are in the feed yard, reduced theeffects of thermal stress.
Subclinical deficiencies of thiamin can result in reduced synthesis ofother B-vitamins since some rumen bacteria require thiamin to grow.
Mathison (1986) reported one feedlot trial in which a significant response to supplemental thiamin was observed. In this trial, transketolase
was numerically reduced in the controls. Thiamin did not increase gain
in two subsequent trials, although there appeared to be a reduction inbloat in one trial. A concurrent field survey, conducted in Alberta,
Canada, showed that 2.7% of 645 cattle sampled had marginal levels of
TPP. In acute PEM, 1 g/day of injected thiamin is indicated until the animals are eating, then 500 mg/day can be supplemented in the diet for 7
to 14 days (Mathison, 1986). A level of 4 to 6 mg/kg diet in high-grain
diets was suggested to help prevent subclinical deficiency.
Thiamin supplementation should be greatly modified if diets contain anti-thiamin substances, such as thiaminases from fish or moldy
feeds. As an example, in free-range farming, pigs may occasionally suffer from bracken poisoning, as the roots contain anti-thiamin substances. The animals can be saved by timely thiamin injections.
For mink, the thiamin requirement is estimated to be 1.2 mg/kg dry
feed. In Scandinavia, however, due to the risk of destruction of thiamin
when mink are fed on fish, much higher dosages (up to 6 mg/kg) are rec302

THIAMIN

ommended (Bräunlich and Zintzen, 1976). Other fish eaters, for example, seals, may also develop thiamin deficiency (Geraci, 1974). It has
been calculated and confirmed by determinations of blood transketolase
activity that a seal weighing 80 kg and consuming 4 to 6 kg of fish per
day has a daily thiamin requirement of 100 to 150 mg.
Intentionally added nonnutrient substances to diets are sometimesof concern, such as the coccidiostat amprolium, a thiamin antimetabolite. A mild thiamin deficiency from amprolium added to a standard
commercial hen feed caused a reduction in the feed intake and egg-laying performance and an increase in the mortality of embryos and chicks.
These phenomena could be prevented or effectively counteracted byhigh thiamin doses in the feed. At recommended levels, amprolium apparently does not interfere with the thiamin metabolism of the chicken(Scott et al., 1982). Sometimes other thiamin antagonists, such as thefree bisulfite present in certain menadione sulfite forms or other formsof sulfur, are added to the feed.
Animals with clinical signs of thiamin deficiency and/or other indicators of thiamin insufficiency (e.g., transketolase activity) should begiven thiamin at therapeutic doses. Since thiamin deficiency causes
anorexia, injection of the vitamin is preferred to oral doses when deficiency is severe. In calves weighing less than 50 kg, clinical signs wereprevented with 0.65 mg of thiamin-HCl per kilogram of liquid diet fed
at 10% of live weight (65 mg/kg live weight) (Johnson et al., 1948). Animals with PEM need to be rapidly provided with supplemental thiamin.
Levels of thiamin to be administered intravenously or intramuscularlyfor 3 days have been recommended for lambs and calves (100 to 400
mg/day) and for sheep and cattle (500 to 2,000 mg/day) (Zintzen, 1974).
For general maintenance following the treatment of mild cases (oras a prophylactic measure when a herd is at risk), 5 to 10 mg of thiaminshould be added to 1 kg of dry feed. Feeds should be enriched with thiamin in a concentration such that each animal will receive 100 to 500
mg daily. Likewise, roughage should be added to the daily ration at alevel of 1.5 kg/100 kg body weight. For therapeutic purposes, 6.6 to 11
mg/kg body weight repeated every 6 hours for 24 hours has been suggested for goats (Smith, 1979).
The administration of thiamin to animals with PEM generally produces rapid results, sometimes in a matter of hours. When recognitionof the disease has been delayed and irreversible necroses have alreadydeveloped in the brain, treatment with the vitamin may be useless. The
prospects of achieving satisfactory treatment of animals incapable ofstanding are very limited. Although treatment improves the condition of303

CHAPTER SIX

such animals, relapses and permanent damage are probable due to irreversible changes in the CNS. Without doubt, PEM is the most importantdisease arising from thiamin deficiency in ruminants. It is also noteworthy that thiamin can be used effectively as a support in the treatment of
the two metabolic disorders of rumen—acidosis and ketosis. Even
though treatment with thiamin can be therapeutically successful, it does
not follow that deficiency of thiamin contributes to the etiology of thesetwo diseases (Zintzen, 1974).
In horses, parenterally administered thiamin in high doses (1,000
mg intramuscularly) has a marked sedative effect that is particularly noticeable in nervous and excited horses (Bräunlich and Zintzen, 1976).
Intramuscular or intravenous injections of 1,000 to 2,000 mg of thiaminalso have a digitalis-like effect on the heart. Thiamin is also used indoses of 200 to 300 mg for the alleviation of muscle cramps (myoglobinuria, melanuric colic, spastic constipation).
Effects of thiamin supplements on exercising horses were exploredby Topliff et al. (1981), who proposed that 4 mg of thiamin per kilogram of air-dry diet may not be sufficient. It may be prudent to ensurethat the diets of performance horses contain 5 mg of thiamin per kilogram of diet dry matter (NRC, 1989b).
Thiamin sources available for addition to feed are the hydrochloride
and mononitrate forms. Because of its lower solubility in water, themononitrate form is preferred for addition to premixes; it has somewhat
better stability characteristics in dry products than the hydrochloride(Bauernfeind, 1969).
Stability of thiamin in feed premixes can be a problem. More than
50% of the thiamin was destroyed in premixes after 1 month at room
temperature (Verbeeck, 1975). When thiamin was observed in premixes
without minerals, no losses were encountered when kept at room temperature for 6 months. After 6 months only 27% of the activity of thiamin hydrochloride remained in a vitamin premix that also containedcholine and trace minerals (Coelho, 1991).
For humans in developed countries, a number of foods have been
fortified with thiamin (e.g., bread enrichment) for many years. Rice enrichment in the Far Eastern countries has dramatically reduced incidenceof beriberi. It has even been suggested that addition of thiamin to alcoholic beverages might help prevent the serious effects of thiamin deficiency in heavy consumers as well as in alcoholics. However, Crane andPrice (1983) conclude that the addition of thiamin to beer in the form ofalkyl disulfides is not a practical or effective method of enrichment andoffers no special advantage. In world regions where beriberi is still en304

THIAMIN

demic, a good case could be made for routine administration of thiamin
to all patients in whom heart failure is present without clear evidence ofthe cause (Djoenaidi et al., 1992).

TOXICITY
Thiamin in large amounts is not toxic, and usually the same is trueof parenteral doses. Excess thiamin is easily cleared by the kidneys. Dietary intakes of thiamin up to 1,000 times the requirement are apparently safe for most animal species (NRC, 1987). Intolerance to thiaminis relatively rare in humans; daily oral doses of 500 mg have been administered for as long as a month with no ill effects. However, there
were several reports of toxic reactions following repeated parenteral injections of large doses. These were anaphylactic reactions resulting fromsensitization to the vitamin. Parenteral doses of thiamin at 100 times therecommended intake produced headache, convulsions, weakness, paralysis, cardiac arrhythmia, and allergic reactions (Combs, 1992).
Lethal doses with intravenous injection were 125, 250, 300, and
350 mg/kg body weight for mice, rats, rabbits, and dogs, respectively(Gubler, 1991). Vasodilation, fall in blood pressure, bradycardia, respiratory arrhythmia, and depression resulted when animals were given thiamin in large doses intravenously.

■ REFERENCES
Abe, T. (1969). J. Vitaminol. 15, 339.
Aitken, F. C., and Hankin, R. G. (1970). Vitamins in Feeds for Livestock, Commonwealth Bureau of Animal Nutrition, Techn. Comm. No. 25, Central
Press, Aberdeen.
Barchi, R. L. (1976). Thiamine (C. J. Gubler, M. Fujiwara, and P. M. Dreyfus,eds.) p. 282. John Wiley and Sons, New York.
Barclay, L. L., and Gibson, G. E. (1982). J. Nutr. 112, 1899.
Bauernfeind, J. C. (1969). World Rev. Anim. Prod. 5(21), 20.
Blank, N. K., Vick, N. A., and Schulman, S. (1975). Acta Neuropathol. 31, 137.
Blass, J. P., and Gibson, G. E. (1977). N. Engl. J. Med. 297, 1367.
Blass, J. P., and Gibson, G. E. (1978). Advances in Neurology, Vol. 21. R. A. P.
Kark, R. N. Rosenberg, and L. J. Schut (Eds.). Raven Press, p. 181-198,
New York.
Bräunlich, K., and Zintzen, H. (1976). Vitamin B1 No. 1593. Hoffmann-La
Roche, Basel, Switzerland.
Brent, B. E. (1985). Feed Management 36(12), 8.
Brent, B. E., and Bartley, E. E. (1984). J. Anim. Sci. 59, 813.
Brethour, J. R. (1972). J. Anim. Sci. 35, 260(Abstr.).
305

CHAPTER SIX

Breves, G., Brandt, M., Hoeller, H., and Rohr, K. (1984). J. Agr. Sci. Camb. 96,
587.
Brin, M. (1962). Ann. N.Y. Acad. Sci. 98, 528.
Brown, M. L. (1990). In Present Knowledge in Nutrition (M. L. Brown, ed.), 6thEd., p. 142. International Life Sci. Inst., Washington, D.C.
Carroll, F. D., Gross, H., and Howell, C. E. (1949). J. Anim. Sci. 8, 290.
Charles, O. W., Roland, D. A., and Edwards Jr., H. M. (1972). Poult. Sci. 51,
419.
Coelho, M. B. (1991). Vitamin stability in premixes and feeds: A practical approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Combs Jr., G. F. (1992). The Vitamins. Academic Press, San Diego, California.
Cook, M. E. (1990). In Proc. 1990 Natl. Feed Ingred. Assoc. Nutr. Inst. Development in Vitamin Nutrition and Health Applications, Kansas City, Missouri. National Feed Ingredients Association (NFIA), Des Moines, Iowa.
Cooper, J. R., Roth, R. H., and Kini, M. M. (1963). Nature (London) 199, 609.
Crane, S., and Price, J. (1983). J. Nutr. Sci. Vitaminol. 29, 381.
Davidson, M. G. (1992). Vet. Rec. 130(5), 94.
Davies, E. T., Pill, A. H., and Austwick, P. K. A. (1968). Vet. Rec. 83, 681.
Diniz, J. M. F., Bashe, J. R., and deCamargo, N. J. (1984). Arquivo Brasileiro deMed. Vet. Zootech. 36, 512.
Djoenaidi, W., Notermans, S. L. H., and Dunda, G. (1992). Eur. J. Clin. Nutr.
46, 227.
Dougherty, L. T., Lauriault, L. M., Bradley, N. W., Gay, N., and Cornelius, P. L.
(1991). J. Anim. Sci. 69, 1008.
Earl, J.W., and McCleary, B.V. (1994). Nature 368, 683.
Edwin, E. E., and Lewis, G. (1971). J. Dairy Res. 38, 79.
Edwin, E. E., Lewis, G., and Allcroft, R. (1968). Vet. Rec. 83, 176.
Edwin, E. E., Markson, L. M., and Shreeve, J. (1979). Vet. Rec. 104, 4.
Ellis, N. R., and Madsen, L. L. (1944). J. Nutr. 27, 253.
Ensminger, A. H., Ensminger, M. E., Konlande, J. E., and Robson, J. R. K.
(1983). In Foods and Nutrition Encyclopedia, Vol. I, A-H, p. 1,208. Pegus
Press, California.
Evans, W. C. (1975). Vit. Horm. 33, 467.
Fontenot, J. P., Allen, V. G., and Brock, A. (1988). Grazing low and high endophyte infected fescue by cattle and effects of thiamin supplementation.
Anim.-Sci. Research Rep., Virginia Agricultural Experiment Station No.
8:108, Blacksburg, Virginia.
Fritz, J. C., Mislivec, P. B., Pla, G. W., Harrison, B. N., Weeks, C. E., and Dantzman, J. G. (1973). Poult. Sci. 52, 1523.
Frye, T. M., Williams, S. N., and Graham, W. (1991). Vet. Clin. North Am. Food
Anim. Pract. 7, 217.
Geraci, J. R. (1974). J. Am. Vet. Med. Assoc. 165, 801.
Gibson, D. M., Kennelly, J. J., and Aherne, F. X. (1987). Can. J. Anim. Sci. 67,
841.
Glad, B. W., Hodges, R. E., and Michas, C. A. (1978). Am. J. Med. 65, 69.
Gooneratne, S. R., Olkowski, A. A., and Christensen, D. A. (1989). Can. J. Vet.
Res. 53, 462.
Gould, D.H. (1998). J. Anim. Sci. 76, 309
306

THIAMIN

Gould, D. H., McAllister, M. M., Savage, J. C., and Hamar, D. W. (1991). Am.
J. Vet. Res. 52, 1164.
Grewal, P. K., and Sangha, J. K. (1990). J. Sci. Food Agric. 52, 387.
Gries, C. L., and Scott, M. L. (1972). J. Nutr. 102, 1269.
Gubler, C. J. (1991). In Handbook of Vitamins (L. J. Machlin, ed.), 2nd Ed., p.
233. Marcel Dekker, Inc., New York.
Hashimoto, Y., Arai, S., and Nose, T. (1970). Bull. Jap. Soc. Sci. Fish 36, 791.
Heinemann, W. W., Ensminger, M. E., Cunha, T. J., and McCulloch, E. C.
(1946). J. Nutr. 31, 107.
Hoffmann-La Roche. (2000). Vitamins for Dogs and Cats, Nutley, New Jersey
Horwitt, M. K., Liebert, E., Kreisler, O., and Wittman, P. (1948). Investigationsof Human Requirements for B-Complex Vitamins Bulletin No. 116. National Academy of Sciences, Washington, D.C.
Houston, D. M., and Hulland, T. J. (1988). Can. Vet. J. 29, 383.
Hughes, B. L., and Wood-Gush, D. G. M. (1971). Physiol. Behav. 6, 331.
Jansen, B. C. P. (1956). Nutr. Abstr. Rev. 26, 1.
Johnson, A. B., and Krautmann, B. A. (1989). In Proceedings of the 24th PacificNorthwest Animal Nutrition Conference, Boise, Idaho.
Johnson, B. C., Hamilton, T. S., Nevens, W. B., and Boley, L. E. (1948). J. Nutr.
35, 137.
Jubb, T. F. (1992). Vet. Rec. 131, 347.
Kao, C., and Robinson, R. J. (1973). J. Food Sci. 37, 261.
Kingsbury, J. M. (1964). Poisonous Plants of the United States and Canada. p.
105. Prentice-Hall, Englewood Cliffs, New Jersey.
Kuroki, F., Iida, M., Tominaga, M., Matsumoto, T., Hirakawa, K., Sugiyama, S.,
and Fujishima, M. (1993). Di. Dis. Sci. 38, 1614.
Lauriault, L. M., Dougherty, C. T., Bradley, N. W., and Cornelius, P. L. (1990).
J. Anim. Sci. 68, 1245.
Leevy, C. M. (1982). Ann. N.Y. Acad. Sci. 378, 316.
Linerode, P. A. (1966). Studies on the Synthesis and Absorption of B Complex
Vitamins in the Equine. Ph.D. Dissertation, Ohio State University, Wooster.
Loew, F. M. (1978). In Handbook Series in Nutrition and Food, Section E: Nutritional Disorders, (M. Rechcigl Jr., ed.) Vol. 2, p. 3. CRC Press, Boca Raton, Florida.
Loew, F. M., and Dunlop, R. H. (1972). Am J. Vet. Res. 32, 2195.
Lofland, H. B., Goodman, H. O., Clarkson, R. B., and Prichard, R. W. (1963).
J. Nutr. 79, 188.
Long, J. B., and Shaw, J. N. (1943). North Am. Vet. 24, 234.
Lonkar, P. S., Sharma, S. N., Yadav, J. S., and Prasad, M. C. (1993). Indian Vet.
J. 70, 873.
Loosli, J. K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.), p. 25.
Academic Press, San Diego, California.
Losada, H., Dixon, F., and Preston, T. R. (1971). Rev. Cubana Cienc. Agric. 5,369.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease.
Medical and Technical Publishing Co., Ltd., p. 73. Lancaster, England.
Mathison, G. W. (1986). In Proc., 21st Northwest Animal Nutrition Conference, p. 107. Vancouver, Canada.
307

CHAPTER SIX

Maynard, L. A., Loosli, J. K., Hintz, H. F., and Warner, R. G. (1979). Animal
Nutrition. McGraw-Hill Book Co., New York.
McAllister, M. M., Gould, M. M., and Hamar, D. W. (1992). J. Compar. Pathol.
106, 267.
McDonald, J. W. (1982). Aust. Vet. J. 58, 212.
McDowell, L. R. (1985). In Nutrition of Grazing Ruminants in Warm Climates(L. R. McDowell, ed.), p. 359. Academic Press, Orlando, Florida.
McManus, E. C., and Judith, F. R. (1972). Poult. Sci. 51, 1835.
Miles, W. H., and McDowell, L. R. (1983). World Anim. Rev. 46, 2.
Miller, E. R., Schmidt, D. A., Hoefer, J. A., and Luecke, R. W. (1955). J. Nutr.
56, 423.
Mullenax, C. (1983). Carta Ganadera 20(10), 7.
Muralt, A. (1962). Ann. N.Y. Acad. Sci. 98, 499.
Morris, H. P. (1947) Vitam. Horm. 5, 175.
Naidoo, D. (1956). Acta Psychiatr. Neurol. Scand. 31, 205.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Olkowski, A. A., Gooneratne, S. R., and Christensen, D. A. (1990). Res. Vet.
Sci. 48, 82.
Olkowski, A. A., Rousseaux, C. G., and Christensen, D. A. (1991). Can. J.
Anim. Sci. 71, 825.
Olkowski, A. A., Gooneratne, S. R., Rousseaux, C. G., and Christensen, D. A.
(1992). Res. Vet. Sci. 52, 78.
Olkowski, A. A., Laarvold, B., Patience, J. F., Francis, S. I., and Christensen, D.
A. (1993). Int. J. Vit. Nutr. Res. 63, 38.
Ollilainen, V., Vahteristo, L., Uusi-Rauva, A., Varo, P., Koivistoinen, P., and
Huttunen, J. (1993). J. Food Compos. Anal. 6, 152.
Oltjen, R. R., Sirny, R. J., and Tillman, A. D. (1962). J. Nutr. 77, 269.
Pardo, B., Sygnowska, E., Rywik, S., Kulesza, W., and Waskiewicz, A. (1991).
Appetite, Academic Press, p. 1. London, England.
Phillips, S. C. (1987). Acta Neuropathol. 73, 171.
Piktelite, O. S., Aleinik, S. I., Yakushina, L. M., Blazheevich, N. V., Isaeva, V. A.,
308

THIAMIN

Alekseeva, I. H., Glinka, E. Y., and Grishchenko, N. L. (1992). Voprosy-Pitaniyay 4, 32.
Polin, D., Wynosky, E. R., and Porter, C. C. (1962). Proc. Soc. Exp. Biol. Med.
110, 844.
Polin, D., Ott, W. H., Wynosky, E. R., and Porter, C. C. (1963). Poult. Sci. 42,
925.
Rains, T.M., Emmert, J.L., Baker, D.H., and Shay, N.F. (1997). J. Nutr. 127,
167.
Ralston Purina. (1987). Nutrition and Management of Dogs and Cats. Ralston
Purina Co., St. Louis, Missouri.
Rammell, C. G., and Hill, J. H. (1986). N Z Vet. J. 34, 202.
Rathanaswami, P., and Sundaresan, R. (1991). Biochem. Int. Marrickville 24(6),1057.
RDA. (1989). Recommended Dietary Allowances, 10th Ed., National Academyof Sciences-National Research Council, Washington, D.C.
Reddy, M. U., and Pushpamma, P. (1986). Nutr. Rep. Int. 34, 393.
Reid, J. M., Hove, E. L., Braucher, P. F., and Mickelsen, O. (1963). J. Nutr. 80,
381.
Reid, M. E., and Bieri, J. G. (1967). Proc. Soc. Exp. Biol. Med. 126, 11.
Remus, J. C., and Firman, J. D. (1990). J. Nutr. Biochem. 1, 636.
Rinehart, J. F., and Greenberg, L. D. (1949). Arch. Pathol. 48, 89.
Rose, R. (1990). In Proc. 1990 Natl. Feed Ingred. Assoc. Nutr. Inst. Developments in Vitamin Nutrition and Health Applications Kansas City, Missouri.
National Feed Ingredients Association (NIFA), Des Moines, Iowa.
Sadler, W. C., Mahoney, J. H., Puch, H. C., Williams, D. L., and Hodge, D. E.
(1983). Anim. Sci. Abstr. p. 467.
Salley, J. J., Eshleman, J. R., and Morgan, J. H. (1962). J. Dent. Res. 41, 1405.
Scott, N. L., Nesheim, M. C., and Young, R. J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Scriver, C. R. (1973). Metabolism, 22, 1319.
Sebrell, W. H., and Harris, R. S. (1973). The Vitamins: Chemistry, Physiology,
Pathology and Methods Vol. V., p. 98. Academic Press, New York.
Shell, L.G. (1995). Feline Pract. 23, 27.
S’Klan, D., and Trostler, N. (1977). J. Nutr. 107, 357.
Smith, D.C., and Proutt, L.M. (1944). Proc. Soc. Exp. Biol. Med. 56, 1.
Smith, M. C. (1979). J. Am. Vet. Med. Assoc. 174, 1328.
Smith, W. L., Gay, N., Boling, J. A., and Crowe, M. W. (1986). J. Anim. Sci.
63(Suppl. 1), 296.
Soita, H. W., and Brent, B. E. (1993). J. Anim. Sci. 71(Suppl. 1), 278.
Strauss, L. H., and Scheer, P. (1939). Int. Z. Vitam. Forsch 9, 39.
Studdert, V. P., and Labuc, R. H. (1991). Aust. Vet. J. 68, 54.
Sturkie, P. D., Singsen, E. P., Matterson, L. D., Kozeff, A., and Jungherr, E. L.
(1954). Am. J. Vet. Res. 15, 457.
Swank, R. L. (1940). J. Exp. Med. 71, 683.
Takeuchi, T., Jung, E. H., Nishino, K., and Itokawa, Y. (1990). Int. J. Vitam.
Nutr. Res. 60(2), 112.
Tang, C. M., Rolfe, M., Wells, J. C., and Cham, K. (1989). Lancet 2(8656), 206.
Tanphaichair, V. (1976). In Nutrition Reviews Present Knowledge in Nutrition
309

CHAPTER SIX(D. M. Hegsted, C. O. Chichester, W. J. Darby, K. W. McNutt, R. M.
Stalvey, and E. H. Stotz, eds.). The Nutrition Foundation Inc., New York.
Thornber, E. J., Dunlop, R. H., Gawthorne, J. M., and Huxtable, C. R. (1979).
Res. Vet. Sci. 26, 378.
Thornton, P. A., and Schutze, J. E. (1960). Poult. Sci. 39, 192.
Topliff, D. R., Potter, G. D., Kreider, J. L., and Creagor, C. R. (1981). In Proc.
7th Eq. Nutr. Physiol. Soc. Symp., p. 167. Warrenton, Virginia.
Tucker, D. M., Penland, J. G., Sandstead, H. H., Milne, D. B., Heck, D. G., andKlevay, L. M. (1990). J. Food Sci. 52, 93.
Uherová, R., Hozová, B., and Smirnov, V. (1993). Food Chem. 46, 293.
Verbeeck, J. (1975). Feedstuffs 47(36), 4.
Wetherilt, H., Ackurt, F., Brubacher, G., Okan, B., Aktas, S., and Turdu, S.
(1992). Int. J. Vitam. Nutr. Res. 62, 21.
Williams, R. R. (1961). Toward the Conquest of Beriberi. Harvard University
Press, Cambridge, Massachusetts.
Zempleni, J., Link, G., and Kubler, W. (1992). Int. J. Vitam. Nutr. Res. 62, 165.
Zintzen, H. (1974). Vitamin B (Thiamine) in the Nutrition of the Ruminant, No.
1460. Hoffmann-La Roche, Basel, Switzerland.

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INTRODUCTION
After isolation of thiamin (vitamin B1) as the “vitamin B” factor thatprevented beriberi and polyneuritis, riboflavin (vitamin B2) was the firstgrowth factor to be characterized from the remaining B-complex vitamins. Riboflavin in the form of flavin mononucleotide (FMN) and flavinadenine dinucleotide (FAD) functions as a coenzyme in diverse enzymatic reactions. The vitamin is required in the metabolism of all plants
and animals, and every plant and animal cell contains the vitamin.
Because of microbial ruminal synthesis, adult ruminants apparently
do not require dietary riboflavin. Young ruminants, prior to development of the rumen, and other species require dietary sources of riboflavin because of their very limited synthesis by intestinal flora. Riboflavin is one of the vitamins most likely to be deficient in typical swine
and poultry diets based on grains and plant protein supplements (e.g.,soybean meal). Likewise, human diets low in animal protein products(especially milk and egg products) and leafy vegetables are likely to be
deficient in riboflavin.

HISTORY
The historical aspects of riboflavin were reviewed by McCollum(1957), Wagner-Jauregg (1972), Sharman (1977), Scott et al. (1982),
and Loosli (1991). In 1915, it was known that a water-soluble factor or
factors promoted growth and prevented beriberi in rats. In 1920, it was
found that heating feedstuffs (e.g., yeast) destroyed the beriberi preventive effect more readily than the growth-promoting effect. Therefore, it
was determined that this water-soluble B vitamin contained two essen311

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tial factors, one of which was more stable to heat than the other. The
less stable factor was labeled vitamin F (B1 or thiamin) and the heat-stable factor vitamin G (B2 or riboflavin).
The biological importance of certain yellow pigments became apparent in 1932, when Warburg and Christain (Germany) isolated an oxidative enzyme (“old yellow enzyme”) from yeast that was yellow withgreen fluorescence. They split it into a protein and a nonprotein (pigment) fraction. This was the first identification of a prosthetic or activating group of an enzyme. Thus, riboflavin was found in a coenzyme
before it was discovered in free form. The yellow enzyme was found necessary for oxidation of glucose-6-phosphate.
In 1933, Kuhn (Germany) isolated a yellow pigment from egg whitethat showed green fluorescence and had oxidative properties. Kuhn suggested that this growth factor for rats be given the name flavin. Therefore, the terms ovoflavin from eggs, lactoflavin from milk, hepatoflavinfrom liver, and uroflavin from urine were used. Pure crystalline flavincompounds were found to contain ribose, and thus, the name riboflavinbecame popular.
In 1934–1935, riboflavin was synthesized by two groups—Kuhn’sin Germany and Karrer’s in Switzerland. At the same time, Györgi inGermany proved that the biological activity of the synthetic form was
the same as that of the natural vitamin. The synthesis of FMN was accomplished in 1936 by Kuhn and Rudy. In 1938, Warburg discovered
FAD as the coenzyme of D-amino acid oxidase. Other flavin-dependent
enzymes were discovered thereafter at a rapid rate.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Riboflavin consists of a dimethylisoalloxazine nucleus combinedwith the alcohol of ribose as a side chain. All flavins are isoalloxazines,
which are 10-substituted derivatives of alloxazine, the parent tricyclicring system, with nitrogens in positions 1, 3, 5, and 10. Riboflavin exists in three forms: as the free riboflavin and as the coenzyme derivatives
FMN (also called riboflavin 5-phosphate) and FAD (Fig. 7.1). The coenzyme derivatives are synthesized sequentially from riboflavin. In the firststep, catalyzed by flavokinase, riboflavin reacts with adenosine triphosphate (ATP) to form FMN; then FMN combines with a second moleculeof ATP to form FAD in a reaction catalyzed by the enzyme FAD pyrophosphorylase.
Riboflavin is an odorless, bitter orange-yellow compound that melts
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Fig. 7.1 Structures of riboflavin, flavin mononucleotide (FMN), and flavinadenine dinucleotide (FAD).

at about 280oC. Its empirical formula is C17H20N4O6, with an elemental analysis of carbon 54.25%, hydrogen 5.36%, and nitrogen 14.89%.
Riboflavin is only slightly soluble in water but readily soluble in dilutebasic or strong acidic solutions. It is quite stable to heat in neutral andacid, but not alkaline solutions; very little is lost in cooking. Aqueous solutions are unstable to visible and ultraviolet light, and instability is in313

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creased by heat and alkalinity. When dry, it is not affected appreciably
by light, but in solution, it is quickly destroyed.
Loss in milk during pasteurization and exposure to light is 10 to
20%. Much larger losses (50 to 70%) can occur if bottled milk is left
standing in bright sunlight for more than 2 hours. Poultry mashes left
exposed to direct sunlight for several days and frequently stirred are subject to some loss. In alkaline solution, light splits off ribityl residue,
rapidly forming lumiflavin (7,8,10-trimethylisoalloxazine). In acid or
neutral solution, light decomposition produces lumichrome (7,8-dimethylalloxazine).
Many flavins—which differ structurally and functionally from riboflavin, FMN, and FAD—have been discovered, with physiologicalroles often unknown. Antiriboflavin compounds may result from chemical changes in either the isoalloxazine nucleus or the ribityl side chain.
A number of synthetic homologs of riboflavin exist. The antagonist Dgalactoflavin, 7,8-dimethyl-10-(d-l´-dulcityl) isoalloxazine, has beenused experimentally in animals and humans to hasten the developmentof riboflavin deficiency (Cooperman and Lopez, 1991). The phenothiazines (e.g., chlorpromazine, used in the treatment of schizophrenia)
and the tricyclic antidepressant drugs (e.g., imipramine) are structural
analogues of riboflavin and inhibit flavokinase. In experimental animals,
administration of these drugs at doses equivalent to those used clinicallyresults in increased urinary excretion of riboflavin, with reduced tissue
concentrations of FMN and FAD, despite feeding diets providing moreriboflavin than is needed to meet requirements (Pinto et al., 1981).

ANALYTICAL PROCEDURES
Analytical methods for riboflavin determination in feeds and biological tissues employ fluorometric or microbiological assays. The firstassays measured the biological response in the rat and the chicken, but
riboflavin can be assayed more readily by chemical or microbiologicalmethods, so animal assays have given way to these. Growth and production of lactic acid by Lactobacillus casei are dependent on presenceof riboflavin in the medium. Turbidity from the growth of the bacteria
can be read in a colorimeter after 16 to 24 hours of incubation at 37oC.
Riboflavin rarely occurs free in nature, so natural products usuallymust be treated with acid or enzymes to liberate the riboflavin. If FMN
or FAD are to be measured, special nonhydrolytic extraction procedures
are required. Unique properties of riboflavin allowing for successful flu-314

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orometric analysis include the following (Scott et al., 1982): (1) Riboflavin gives off an intense greenish fluorescence, with intensity proportional to the concentration; (2) it is stable to heat and acid, allowing
release from proteins; and (3) it is stable to oxidizing agents (e.g., potassium permanganate) needed to destroy other fluorescing materials. Because riboflavin is readily destroyed by blue or violet light, operations
must be done in dim light or in amber or red glassware. Also, at a pHof greater than 7.0, dilute solutions may be destroyed by alkali even on
clean glassware.
A high-performance liquid chromatographic (HPLC) methodology
has been developed to determine riboflavin in plant (Fernando and Murphy, 1990) and animal (Li et al., 1993) sources that compares with fluorometric procedures. Using HPLC, it is possible to simultaneously determine the three principal forms of the vitamin, riboflavin, FMN, and
FAD in foods (Russell and Vanderslice, 1991).

METABOLISM
Digestion, Absorption, and Transport
Riboflavin is found in feeds as FAD, FMN, and free riboflavin. Riboflavin covalently bound to protein is released by proteolysis digestion.
Phosphorylated forms (FAD, FMN) of the riboflavin are hydrolyzed byphosphatases in the upper gastrointestinal tract to free the vitamin forabsorption. For the rat, the ribityl side chain and the NH group at position 3 of the isoalloxazine moiety are essential for riboflavin binding to
the small intestine brush border membrane (Casirola et al., 1994). Freeriboflavin enters mucosal cells of the small intestine after apparently being absorbed in all parts of the small intestine. In a comparison between
jejunal and ileal cells from guinea pigs, cells from both portions of thesmall intestine transported riboflavin almost equally (Hegazy and
Schwenk, 1983). Cells from deficient animals have a greater maximal
absorption uptake of riboflavin (Rose et al., 1986). At low concentrations, riboflavin absorption is an active Na+-dependent carrier-mediatedprocess (Hegazy and Schwenk, 1983). However, Said et al. (1993a,b)
suggest that the carrier-mediated system in the rabbit is Na+- and pH-independent in nature and transports the substrate rather by an electroneutral process. At high concentrations, riboflavin is absorbed bypassive diffusion, proportional to concentration (Middleton, 1990). In
mucosal cells, much of the riboflavin is phosphorylated to FMN by fla-315

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vokinase. Riboflavin enters the blood as both FMN and the free vitamin.
In tissues, most of the FMN is then converted to the other coenzyme
FAD by FAD-pyrophosphorylase.
Transport of flavin by blood plasma is known to involve both loose
association with albumin and tight associations with some globulins(McCormick, 1990). A genetically controlled riboflavin-binding protein
is present in serum and eggs. There is a hereditary recessive disorder inchickens—renal riboflavinuria—in which the riboflavin-binding protein
is absent (White, 1996). Eggs become riboflavin deficient, and embryos
generally do not survive beyond the fourteenth day of incubation(Clagett, 1971). Also, if riboflavin-binding protein is in excess, it can diminish riboflavin availability to the chicken embryo (Lee and White,1996). Presumably, the lack of the specific vitamin transport protein prevents adequate transfer of dietary riboflavin to the developing fetus, andriboflavin losses occur via maternal urine.
In addition to poultry, specific binding proteins have been found inserum from pregnant cows and rats, human fetal blood, and uterine secretions in the pig. Riboflavin-carrier proteins were induced by estrogens and synthesized in liver (Durgakumari and Adiga, 1986). Rivlin(1984) suggested there may be physiological mechanisms in pregnancy
that facilitate transfer of riboflavin from maternal stores to the fetus ina manner that is fundamentally similar to that in the laying hen. Riboflavin was efficiently transferred to the fetus. In human placenta, riboflavin concentration was 8.7 mmol/L in maternal plasma versus 40.6
mmol/L for venous cord plasma, indicating favorable retention by the
fetus (Zempleni et al., 1992).
Much of the control of riboflavin utilization is at the level of flavocoenzyme formation. Hormonal control affects riboflavin metabolism,
and specific binding proteins can be greatly increased by estrogen administration (Rivlin, 1984). Thyroid hormones enhance conversion of
riboflavin into its coenzyme derivatives and hypothyroidism decreases it.
The major site of thyroid hormone control appears to be flavokinase,and physiological doses of thyroxine increase hepatic activity of this enzyme (Rivlin, 1984). Adrenocorticotropic hormone and aldosteroneboth increase rate of formation of flavin coenzymes from riboflavin.

Tissue Distribution, Storage, and Excretion
Animals do not appear to have the ability to store appreciable
amounts of riboflavin; the liver, kidney, and heart have the richest concentrations. Liver, the major site of storage, contains about one-third of
the total body flavins. A significant amount of free riboflavin exists in
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retinal tissue, but its function is unclear. Free riboflavin constitutes lessthan 5% of the stored flavins, with 70 to 90% in the form of FAD. Intakes of riboflavin above current needs are rapidly excreted in urine, primarily as free riboflavin. The kidney excretes both riboflavin and FMN,
the latter being dephosphorylated in the bladder. Minor quantities of absorbed riboflavin are excreted in feces, bile, and sweat.

FUNCTIONSRiboflavin is required as part of many enzymes essential to utilization of carbohydrates, fat, and protein. More than 100 enzymes are
known to bind FAD or FMN in animal and microbial systems. Riboflavin and related natural flavins participate in numerous and diverse
reactions, perhaps more than for any other vitamin-coenzyme group.
FMN and FAD, which contain riboflavin, function as prosthetic groups
that combine with specific proteins to form active enzymes called flavoproteins. Most flavoproteins contain the FAD form, and a few contain
FMN. Riboflavin in these coenzyme forms acts as an intermediary in thetransfer of electrons in biological oxidation-reduction reactions. The enzymes that function aerobically are called oxidases, and those that function anaerobically are called dehydrogenases. The general function is inoxidation of substrate and generation of energy (ATP). By involvementin the hydrogen transport system, flavoproteins function by accepting
and passing on hydrogen, undergoing alternate oxidation and reduction.
Collectively, the flavoproteins show great versatility in accepting andtransferring one or two electrons with a range of potentials. Many flavoproteins contain a metal (e.g., iron, molybdenum, copper, zinc), and the
combination of flavin and metal ion is often involved in the adjustmentsof these enzymes in transfers between single- and double-electron
donors. Xanthine oxidase contains the metals molybdenum and iron. It
converts hypoxanthine to xanthine and the latter to uric acid. It also reacts with aldehydes to form acids, including the conversion of retinal (vitamin A aldehyde) to retinoic acid.
Flavoproteins may either accept H+ directly from the substrate (thematerial being oxidized) or catalyze the oxidation of some other enzyme
by accepting hydrogen from it, for example, from the niacin-containing
coenzymes, nicotinamide adenine dinucleotide (NAD), and nicotinamide adenine dinucleotide phosphate (NADP). About 40 flavoprotein
enzymes may be arbitrarily classified into three groups:
a. NADH2 dehydrogenases—enzymes whose substrate is a reduced
317

CHAPTER SEVEN

pyridine nucleotide, and the electron acceptor is a member of the cytochrome system or some other acceptor besides oxygen.
b. Oxidases (true)—enzymes that accept electrons from substrate andpass them directly to oxygen; they cannot reduce cytochromes. Oxygen is
reduced to H2O2.
c. Dehydrogenases—enzymes that accept electrons directly from substrate and can pass them to one of the cytochromes.

Substrate

NAD+

a

NADP+

flavoproteinbflavoprotein(other acceptor)
H2O2

CytochromesO2cflavoprotein

The main function of FMN and FAD is transferring hydrogen between the niacin-containing coenzymes, NAD and NADP, the iron porphyrin cytochromes, and directly from the substrate (Fig. 7.2). The sequence of electron acceptors in the early stages of the respiratory chain
indicates that coenzyme Q (ubiquinone) acts between flavoprotein andcytochrome b. Thus these enzymes are a part of the chain that carrieshydrogen from substrates (carbohydrates, amino acids, lipids, etc.) tomolecular oxygen, forming water. These flavoprotein pathways are themost important means of electron transport for both mitochondria andmicrosomes (Scott et al., 1982).
Approximately 40 flavoprotein enzymes participate in electrontransfer from metabolites and pyridine nucleotides to molecular oxygenand include the following:
1. Aerobic dehydrogenases (simple oxidases not containing metals)(a) D and L amino acid oxidases(b) Glycolic acid oxidase(c) Glucose oxidase(d) Warburg’s old yellow enzyme
2. Oxidases (containing copper, iron, molybdenum, or zinc)(a) Cuproflavoprotein in butyryl-CoA-dehydrogenase
318

R I B O F L AV I N

Fig. 7.2 Examples of hydrogen transfer involving flavoproteins. (Modified
from Scott et al., 1982.)(b) Xanthine oxidase (flavoprotein containing iron and molybdenum)(c) Molybdoflavoprotein in aldehyde oxidase(d) DPNH-cytochrome reductase (iron-containing flavoprotein)
3. Anaerobic dehydrogenases(a) Lipoyl dehydrogenase(b) Acyl-CoA dehydrogenases and electron-transferring flavoprotein319

CHAPTER SEVEN(c) Succinic dehydrogenase-fumaric reductase(d) Others
Choline dehydrogenase
α-Glycerophosphate dehydrogenase
L-Galactone-γ-lactone dehydrogenase
L-Lactate dehydrogenase
D-Lactate cytochrome reductase
Pyridine nucleotide-cytochrome c reductases

Riboflavin functions in flavoprotein-enzyme systems to help regulate cellular metabolism, and is also specifically involved in metabolism
of carbohydrates. Riboflavin is also an essential factor in amino acid metabolism as part of amino acid oxidases. These oxidize α-amino acids tocorresponding amino acids that decompose to give ammonia and α-ketoacid. There are distinct enzymes oxidizing D-amino acids (prosthetic
group FAD) and L-amino acids (prosthetic group FMN). An FAD-dependent hydroxylase is involved in the conversion of tryptophan toniacin.
Among the enzymes that require riboflavin is the FMN-dependentoxidase responsible for conversion of phosphorylated pyridoxine (vitamin B6) to a functional coenzyme. In view of the widespread deficiencyof riboflavin in Nigeria, and the involvement of riboflavin in vitamin B6
metabolism, it is speculated that riboflavin status may account for thehigh occurrence of vitamin B6 deficiency in young adults (Ajayi et al.,1989). Riboflavin deficiency also results in a decrease in the conversionof the vitamin B6 coenzyme pyridoxal phosphate to the main vitamin B6
urinary excretory product of 4-pyridoxic acid (Kodentsova et al., 1993).
Riboflavin deficiency had an effect on iron metabolism, with lessiron absorbed and an increased rate of iron loss due to an accelerated
rate of small intestinal epithelial turnover (Powers et al., 1991, 1993). In
Gambian men, riboflavin deficiency reduced iron utilization with hemoglobin levels increased after riboflavin supplementation (FairweatherTait et al., 1992).
Riboflavin plays a role in fat metabolism (Cooperman and Lopez,
1991), and a FAD flavoprotein is an important link in fatty acid oxidation. This includes the acyl-coenzyme A dehydrogenases, which are necessary for the stepwise degradation of fatty acids. An FMN flavoprotein
is required for synthesis of fatty acids from acetate. Thus, flavoproteins
are necessary for both degradation and synthesis of fatty acids.
The fluidity of red blood cell membranes isolated from riboflavindeficient rats was significantly lower than that of the controls. This de320

R I B O F L AV I N

creased fluidity was accompanied by an increase in the activity of themembrane-bound enzyme acetylcholinesterase. This study demonstrated
that a decrease in cells’ ability to cope with peroxidative damage as a result of riboflavin deficiency may lead to changes in the fluidity and function of membranes (Levin et al., 1990). There was increased plasma lipid
peroxidation in riboflavin-deficient, malaria-infected children (Das etal., 1990). It was proposed that riboflavin deficiency restricts regeneration of reduced glutathione, making the parasitized erythrocytes more
vulnerable to destructive lipid peroxidation and increasing plasma lipidhydroperoxides. Elevated riboflavin levels were reported to provide protection against oxidative damage caused by oxidized forms of hemeproteins (Christensen, 1993).
Although riboflavin is present mostly as flavoprotein enzymes FAD
and FMN, the retina contains free riboflavin in relatively large amounts.
The function it fulfills there is still not clear. In avascular tissues such as
the cornea, it is thought that oxidation takes place by means of a riboflavin-containing enzyme. In riboflavin deficiency, the body attempts
oxygenation by vascularization (Marks, 1975).

REQUIREMENTS
Riboflavin requirements vary with heredity, growth, environment,
age, activity, health, other dietary components, and synthesis by host.
Riboflavin requirements for selected animals and humans are presented
in Table 7.1. The majority of species have a requirement between 1 and
4 mg/kg of diet (dry basis). Where sufficient data are available, studies
indicate that riboflavin requirements decline with animal maturity andincrease for reproductive activity. Chicks receiving diets only partiallydeficient in riboflavin may recover spontaneously, indicating that requirement rapidly decreased with age (Scott et al., 1982). However, Deyhim et al. (1992) reported that 3.6 ppm dietary riboflavin for growing
broilers was satisfactory through 4 weeks, but that benefits were obtained by exceeding the 3.6 ppm recommendation through 8 weeks. Several studies have indicated that the riboflavin requirement for prevention of leg paralysis is higher than that for growth (Ruiz and Harms,
1988).
Based upon sow farrowing performance and concentration of erythrocyte glutathione reductase (an indicator of riboflavin status) concentration, Frank et al. (1984) estimated the available riboflavin requirement
for pregnancy to be about 6.5 mg daily. Using the same criteria, the suggested lactational requirement was about 16 mg daily (Frank et al., 1985,
321

CHAPTER SEVEN■ Table 7.1

Riboflavin Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Adult
Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100-g intake)
Broilers, 0–8 weeks
All classes
All classes
0–4 weeks
Adult
Growing-finishing, 3–20 kg
Growing-finishing, 20–120 kg
Adult
All classes
Growing
Growing
Growing
GrowingCatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
Children
AdultsMicrobial synthesis
6.5 ppm milk replacer
Microbial synthesis
Microbial synthesis
3.6 mg/kg
1.8 mg/kg
2.5 mg/kg
3.6 mg/kg
4.0 mg/kg
4.0 mg/kg
4.0 mg/kg
Microbial synthesis
3.0–4.0 mg/kg
2.0–2.5 mg/kg
3.75 mg/kg
2.0 mg/kg
1.5 mg/kg
1.25–4.0 mg/kg
1.0 mg/kg
2–4 mg/kg
9.0 mg/kg
7–25 mg/kg
2.7–15 mg/kg
3.0 mg/kg
7.0 mg/kg
0.4–0.5 mg/day
0.8–1.2 mg/day
1.3–1.8 mg/day

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1981)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1982a)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

Goat
ChickenDuck (Pekin)Japanese quail
Turkey
Sheep
Swine
Horse
Mink
Fox
Cat
Dog
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry
matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg/day.

1988). Massive riboflavin supplementation (10 to 160 mg/d) increased
the percentage of sows farrowed but did not increase litter size (Pettigrew
et al., 1996). Frank et al. (1988) suggested that first-litter gilts have ahigher requirement for riboflavin than the second litter sow based onneeds for both maternal growth and reproduction.
Increased dietary fat or protein increases requirements for riboflavinin rats and chickens. It was assumed that high urinary riboflavin excretion during periods of negative nitrogen balance for a number of species
was a reflection of impaired riboflavin utilization or retention. However,
Turkki and Holtzapple (1982) suggested, in studies with rats, that the effect of protein on riboflavin requirement is related to rate of growth andnot to protein intake per se.
Microbial biosynthesis of riboflavin has been shown to occur in thegastrointestinal tract of a number of animal species and thus affects re322

R I B O F L AV I N

quirements. However, utilization of this endogenously synthesized riboflavin varies from species to species. Within a single species, utilization depends on diet composition and incidence of coprophagy. Carbohydrates such as starch, cellulose, or lactose are absorbed slowly andtherefore exposed for a longer time to the intestinal bacteria, resulting in
an increased riboflavin synthesis. Dextrose, fat, or protein as chief dietary constituents decrease intestinal production, thereby increasing dietary riboflavin requirements. Young rats fed a riboflavin-free, purified
diet with sucrose as the only carbohydrate will cease to grow. However,when sucrose is replaced by starch, sorbitol, or lactose, growth is comparable to that of rats supplied with riboflavin (Haenel et al., 1959). For
humans, diets with increased carbohydrates and decreased fat are suggested to reduce riboflavin requirements (Boisvert et al., 1993). Antibiotics, such as tetracycline, penicillin, and streptomycin, reduce the requirements of several animal species for riboflavin. They may stimulate
microorganisms that synthesize riboflavin or inhibit microorganisms in
the gut that compete for riboflavin.
Because of microbial ruminal synthesis of riboflavin, ruminants
have no dietary requirements. Nevertheless, diet composition influences
total microbial synthesis. Data of Miller et al. (1983a) also suggested agreater riboflavin ruminal synthesis with an increased proportion of dietary concentrates. Buziassy and Tribe (1960) measured increased synthesis of riboflavin with diets containing more protein. These authors
also noted that ruminal synthesis was reduced with higher dietary intakes of the vitamin. Confirmation of net synthesis in the rumen is illustrated by McElroy and Goss (1940) in which the secretion of riboflavinin milk was equivalent to approximately 10 times the intake of the vitamin.
Temperature extremes apparently have an effect on riboflavin requirement. With pigs, riboflavin requirements were substantially higher
at a low environmental temperature (Seymour et al., 1968). At environmental temperatures below 11oC, feed required per unit of gain increased, and rate of gain decreased with decreasing temperature. On thecontrary, Onwudike and Adegbola (1984) reported riboflavin requirements to be higher for chickens in a tropical environment. A dietary levelof 4.1 ppm riboflavin was adequate for egg laying with 5.7 ppm forhatchability, compared to 2.5 ppm for the NRC (1994) requirement.
When requirement comparisons between animals residing at different
environmental temperatures are calculated, total feed intakes must be
considered.
Requirements of humans for riboflavin have been determined from323

CHAPTER SEVEN

experimental studies with adults and infants. Riboflavin allowance forhuman infants and adults ranges from 0.4 to 1.8 mg/day (RDA, 1989).
Levels are to be increased by 0.3 mg during pregnancy, and by 0.5 mg
during the first 6 months of lactation and possibly should be related toenergy expenditure (Roe et al., 1982). A level of 0.5 mg per 1,000 kcal
was estimated to be the minimum requirement for adults to maintain anormal urinary excretion. On a similar basis, the infant’s minimum dailyrequirement was estimated as 0.6 mg per 1,000 kcal.
A number of factors influence riboflavin requirements including
drugs (see Section III), disease, alcohol, heavy metals, and exercise(Cooperman and Lopez, 1991). Riboflavin absorption is decreased in
hyperthyroidism and increased in hypothyroidism. There is an increased
urinary excretion of riboflavin observed in diabetics (Cole et al., 1976).
Riboflavin in blood is significantly decreased in patients with Crohn’sdisease (Kuroki et al., 1993). Alcohol may antagonize the utilization of
FAD from foods (Combs, 1992). Divalent metals (e.g., copper, iron andzinc) form chelates with riboflavin and FMN, reducing absorption of thevitamin. Exercise has been shown to increase riboflavin requirements ofboth young and old individuals (Belko et al., 1985; Trebler-Winters etal., 1992).

NATURAL SOURCES
Riboflavin is synthesized by green plants, yeasts, fungi, and somebacteria. Rapidly growing, green, leafy vegetables and forages, particularly alfalfa, are a good source, with the vitamin richest in the leaves. Cereals and their by-products have a rather low content, in contrast to
their supply of thiamin. Oil meals are fair sources. While grains and protein meals contain some riboflavin, they should not be relied on as thesole source of the vitamin. Riboflavin concentrates obtained from wheyand distiller’s solubles are important commercial sources, particularly
for animal feeds. Although fruits and vegetables are moderately goodsources of this vitamin, they are not consumed in sufficient quantities tomeet daily requirements (Cooperman and Lopez, 1991). Riboflavin concentration in various foods and foodstuffs is shown in Table 7.2.
Riboflavin is more bioavailable from animal products than plantsources. Flavin complexes in plants are more stable to digestion and thus
less digestible than animal sources (Combs, 1992). Singh and Deodhar(1992) concluded that milk factor(s) enhanced intestinal uptake of riboflavin in rat intestine. Bioavailability of riboflavin was less for chicks
fed corn-soybean than purified amino acid diets; riboflavin bioavailabil324

R I B O F L AV I N■ Table 7.2

Riboflavin in Foods and Feedstuffs (ppm, dry basis)

Alfalfa hay, sun cured
Alfalfa leaves, sun cured
Barley, grain
Bean, navy (seed)
Blood meal
Brewer's grains
Buttermilk (cattle)
Chicken, broilers (whole)
Citrus pulp
Clover hay, ladino (sun cured)
Copra meal (coconut)
Corn, gluten meal
Corn, yellow grain
Cottonseed meal,
solvent extracted
Eggs
Fish meal, anchovy
Fish meal, menhaden
Fish, sardine

13.4
23.1
1.8
2.0
2.2
1.6
33.1
15.6
2.7
17.2
3.7
1.8
1.4
5.3
3.0
8.2
5.2
5.8

Linseed meal, solvent extracted
Liver, cattle
Milk, skim, cattle
Molasses, sugarcane
Oat, grain
Peanut meal, solvent extracted
Rice, branRice, grain
Rice, polished
Rye, grain
Sorghum, grain
Soybean meal, solvent extracted
Soybean seedSpleen, cattle
Timothy hay, sun cured
Wheat, bran
Wheat, grain
Yeast, brewer's
Yeast, torula

3.2
92.2
20.5
3.8
1.7
9.8
2.8
1.2
0.6
1.9
1.4
3.2
3.1
15.3
10.1
4.6
1.6
38.1
47.6

Source: NRC (1982b).

ity in the corn-soybean meal diet was 59.1% (Chung and Baker, 1990).
Fermentation significantly increased the proportions of riboflavin present in the free form, resulting in a greater bioavailability of the vitaminin curd than in milk (Raghvendar et al., 1993).
For human diets, milk, eggs, liver, heart, kidney, and muscle meat
are rich sources. Riboflavin content of milk, such as that of cows andgoats, is many times higher than content in their feed because of rumen
synthesis. Human milk contains about 0.5 mg riboflavin/L, while cow’s
milk is three times higher (e.g., 1.7 mg/L). For humans in the UnitedStates, it has been estimated that milk and milk products contribute almost one-half of dietary riboflavin in the diet, with meat, eggs, andlegumes contributing about 25%. Fruits, vegetables, and cereal grains
contribute about 10% each (Hunt, 1975).
Riboflavin is one of the more stable vitamins, but it can be easily destroyed by ultraviolet rays or sunlight. Appreciable amounts may be lost
upon exposure to light; up to one-half the riboflavin content is lost in
cooking eggs and pork chops in light and most of the vitamin in milk
stored in clear glass or plastic bottles. However, processes like pasteurization, evaporation, and condensation have little effect on riboflavincontent of milk. Sun-drying of fruits and vegetables is likely to lead to
substantial losses of vitamin activity. The practice of adding sodium bicarbonate to make vegetables appear fresher accelerates the photodegradation of riboflavin (Rivlin, 1984). Riboflavin in meat is rela325

CHAPTER SEVEN

tively stable during cooking, canning, and dehydration. There is no significant loss of riboflavin from microwave heating (Sigman-Grant et al.,1992). Milling of rice and wheat results in considerable loss of riboflavin, since most of the vitamin is in the germ and bran, which areremoved during this process. About one-half the riboflavin content islost when rice is milled, and whole wheat flour contains about twothirds more riboflavin than white flour (Cooperman and Lopez, 1991).
However, converted or parboiled rice, in which the whole brown rice issteamed, thereby driving the vitamins in the germ and aleurone layers
into the endosperm prior to milling, retains more riboflavin than whole
rice (Grewal and Sangha, 1990).

DEFICIENCY
Effects of Deficiency
Animals and humans are unable to synthesize riboflavin within tissues; therefore, requirements are met principally by dietary sources, withsome intestinal microbial synthesis. Since riboflavin plays many essential
roles in the release of food energy and the assimilation of nutrients, it isunderstandable why deficiency is reflected in a wide variety of signs thatvary among species. However, it is not possible to relate signs to specificbiochemical roles that have been established. A decreased growth rateand lowered feed efficiency are common signs in all species affected.
Typical clinical signs often involve the eyes, the skin, and the nervous
system. Riboflavin deficiency would not be expected in young nursing
animals, as milk is a rich source of the vitamin.

Ruminants
Riboflavin is not required in the diet of adult ruminants because ruminal microorganisms synthesize this vitamin in adequate amounts. Apparently no response to supplemental riboflavin has been reported in animals with a functional rumen. Confirmation of net synthesis in therumen can be obtained from work such as that of McElroy and Goss
(1940), in which secretion of riboflavin in milk was shown to be equivalent to approximately 10 times dietary intake. Miller et al. (1983b) reported that cattle on a concentrate-silage diet synthesize approximately
38 mg of riboflavin in the rumen. A dairy cow producing 21 kg milk per
day loses about 36 mg of riboflavin in milk alone, much more than theamount consumed in the diet. Beef cattle and other lactating ruminants
that have a much lower loss of riboflavin due to lowered milk produc326

R I B O F L AV I Ntion are even less likely to be deficient in the vitamin. Impaired riboflavin synthesis or destruction was reported in sheep with ruminal acidosis, reflected by plasma riboflavin decreasing from 9.5 to 0.09 mg/ml(Basoglu et al., 1993).
Riboflavin deficiencies have been demonstrated in young ruminants
whose rumen flora is not yet established. Failure to provide riboflavinresults in redness of the buccal mucosa, lesions in the corners of themouth, loss of hair, and excessive tear and saliva production (Radostits
and Bell, 1970). Less specific signs are anorexia, diarrhea, and reduced
growth.

Swine
Typical swine diets based largely on grains are often borderline to
deficient in riboflavin. Signs of riboflavin deficiency in the young growing pig include anorexia, slow growth (Fig. 7.3), rough hair coat, dermatitis, alopecia, abnormal stiffness, unsteady gait, scours, ulcerative
colitis, inflammation of anal mucosa, vomiting, cataracts, light sensitivity, and eye lens opacities (Cunha, 1977; NRC, 1998). Reduced feed intake was demonstrated in gilts given a lactation diet containing 1.3
mg/kg of riboflavin. These gilts consumed 30% less feed than gilts receiving 2.3 to 5.3 mg/kg of riboflavin (Frank et al., 1988). Typical clinical signs often involve the eye, the skin, and the nervous system. In severe riboflavin deficiency of pigs, researchers have observed increased
blood neutrophil granulocytes, decreased immune response, discolored
liver and kidney tissue, fatty liver, collapsed follicles, degenerating ova,
and degenerating myelin of the sciatic and brachial nerves (NRC, 1998).
In riboflavin-deficient swine, reproduction is impaired (Fig. 7.4).
Cunha (1977) summarized the clinical signs for gilts fed a riboflavin-deficient diet during reproduction and lactation as follows: (1) erratic or,at times, complete loss of appetite; (2) poor gains; (3) parturition 4 to
16 days premature; (4) one case of death of fetus in advanced stage withresorption in evidence; (5) all pigs either dead at birth or within 48
hours thereafter; (6) enlarged front legs in some pigs due to gelatinous
edema in the connective tissue and generalized edema in many others;and (7) two hairless litters. The longer the period on riboflavin-deficient
diets, the more severe the deficiency signs became. Christensen (1980)
likewise reported resorption of fetuses and premature farrowing for riboflavin-deficient sows.
Riboflavin deficiency has led to anestrus (Esch et al., 1981). Deficiency of riboflavin in postpubertal gilts has led to a cessation of ovarian cyclicity without overt signs of deficiency. Gilts fed a riboflavin-de327

A

B
Fig. 7.3 (A) Riboflavin deficiency in a pig that received no dietary riboflavin.
Note the rough hair coat, poor growth, and dermatitis. (B) Pig that received
adequate riboflavin. (Courtesy of R.W. Luecke, Michigan Agricultural Experiment Station, East Lansing, Michigan, and J. Nutr. 52 [1954], 409.)

328

A

B

C
Fig. 7.4 Riboflavin deficiency. (A) All of the pigs in this litter were born dead;some were in the process of resorption. A few had edema and enlargement offront legs as a result of gelatinous edema. (B) Pigs from a litter in which gelatinous edema was more pronounced. (C) Seven of the ten pigs farrowed wereborn dead, and the other three were dead within 48 hours. The sow received ariboflavin-deficient diet for a shorter period than the sows farrowing the othertwo litters. (Courtesy of T.J. Cunha and Washington State University.)329

CHAPTER SEVEN

ficient diet averaged progressively longer intervals between consecutive
estrus periods until becoming anestrus 63 days after the beginning of thestudy. Riboflavin supplementation to gilts on a corn-soybean diet resulted in improved conception rate, embryonic survival, litter size, andlive piglets at birth (Bazer and Zavy, 1988). Teratogenic effects in newborn pigs have been observed, including skeletal abnormalities, shortened bones, and fusions between ribs (Zintzen, 1975).

Poultry
Only a few of the feedstuffs fed to poultry contain enough riboflavinto meet the requirements of young growing poultry. The most critical requirements for riboflavin are those exhibited by the young chick and thebreeder hen. The characteristic sign of riboflavin deficiency in the chickis “curled-toe” paralysis. However, it does not develop when there is absolute deficiency, or when the deficiency is very marked, because thechicks die before it appears. Chicks are first noted to be walking on theirhocks with their toes curled inward. Deficient chicks do not move about
except when forced to do so, and their toes are curled inward (Fig. 7.5)
both when walking and when resting on their hocks (Scott et al., 1982).
Legs become paralyzed, but the birds may otherwise appear normal.
Changes in the sciatic nerve produces curled-toe paralysis in growing chicks. There is a marked enlargement of sciatic and brachial nerve
sheaths, with the sciatic nerve reaching a diameter four to six times normal size. Histological examinations of affected nerves showed definite
degenerative changes in myelin sheaths which, when severe, may pinch
the nerve, producing a permanent stimulus that causes curled-toe paralysis (Scott et al., 1982). Severe cases showed edema, nerve fiber separation, swelling in interstitial tissue, and leukocyte infiltration in thenerves. Ultrastructurally, the myelin sheaths of the affected nerves weretwisted, separated, or fractured (Gao et al., 1993). When the curled-toe
deformity is long standing, irreparable damage occurred in the sciatic
nerve, and administration of riboflavin no longer cured the condition.
Retarded growth, splay and hock-resting postures, and leg paralysis
rather than curled-toe paralysis have been reported in some studies as
the predominant signs of riboflavin deficiency in chicks (Ruiz andHarms, 1988; Chung and Baker, 1990).
Other signs of riboflavin deficiency are retardation of growth (Figs.
7.6 and 7.7), diarrhea after 8 to 10 days, and high mortality after about
3 weeks. When chicks are fed a diet deficient in riboflavin, their appetite
is fairly good, but they grow very slowly and become weak and emaciated. There is no apparent impairment of feather growth; on the con330

R I B O F L AV I N

Fig. 7.5 Curled-toe
paralysis in a riboflavin-deficient chick.
(Courtesy of M.L.
Scott, Cornell University.)

trary, main wing feathers often appear to be disproportionately long.
Signs of riboflavin deficiency in the poult and duckling differ fromthose in the chick. In the poult, dermatitis appears in about 8 days; thevent becomes encrusted, inflamed, and excoriated; growth is retarded orcompletely stopped by about the seventeenth day; and deaths begin tooccur about the twenty-first day. In the duckling, clinical signs include
diarrhea and cessation of growth.
In laying poultry, hatchability of incubated eggs is first reduced, andsubsequently, egg production is decreased, roughly in proportion to degree of deficiency. Embryonic mortality typically has two peaks, and often a third peak. These are, respectively, on the fourth and twentiethdays and on the fourteenth day of incubation. Embryos that fail to hatch
from eggs of hens receiving low-riboflavin diets are dwarfed and exhibit
pronounced micromelia; some embryos are edematous. The down fails
to emerge properly, resulting in a typical abnormality termed “clubbed”
down, which is most common in neck areas and around the vent. The
nervous systems of these embryos show degenerative changes much likethose described for riboflavin-deficient chicks. When dietary riboflavinprovided to breeder hens was decreased from 9.7 to 1.7 mg/kg, embryo
mortality increased to 83.3% and hatchability to 3.1%; decreasing riboflavin from 9.7 to 7.0, or 4.4 mg/kg had no effect on these variables(Flores-Garcia and Scholtyssek, 1992).
331

CHAPTER SEVEN

A

Fig. 7.6 Riboflavin deficiency in chicks. (A)The chick at left was
fed a corn-soybean
meal diet without supplemental riboflavin; it
exhibited the predominant type of paralysis
observed at the zerolevel of riboflavin supplementation. Bothchicks are female. (B)Same as in (A), but thechicks are male. (Courtesy of N. Ruiz and R.
Harms, University of
Florida.)B

Chicks fed a diet only marginally deficient in riboflavin often recover spontaneously. The condition is curable in the early stages, but in
its acute stage, it is irreversible (NRC, 1994). There is increasing evidence that vigor and livability of the baby chick are directly tied toamount of riboflavin in the hen’s diet (Anonymous, 1969).

Horses
It is generally felt that riboflavin synthesis in the cecum and colon
provides some of the horse’s requirement. When Carroll et al. (1949) fed
a diet containing 0.4 mg of riboflavin per kilogram of dry matter, riboflavin concentrations (mg/kg of ingesta dry matter) in the various intestinal sections were as follows: duodenum, 3.8; ileum, 1.1; cecum, 7.0;
anterior large colon, 9.2; and anterior small colon, 12.2. Horses fed lowriboflavin diets demonstrated anorexia, sporadic but severe weight loss,
general weakness, and poor growth (Pearson et al., 1944).

332

R I B O F L AV I N

ABFig. 7.7 Riboflavin deficiency in turkeys at 21
days of age. (A) The
turkey at left was fed acorn-soybean basal dietwithout supplemental
riboflavin. (B) Severe
leg paralysis and poorfeathering in a turkey
poult fed the riboflavindeficient diet. (Courtesyof N. Ruiz and R.
Harms, University of
Florida.)Other Animal Species
DOGS AND CATS
Riboflavin-deficient dogs exhibit low growth rates, anemia, andcorneal lesions (NRC, 1985a). The animals collapse, become comatose,and die. During the deficiency state, dermatitis develops on the chest,
abdomen, inner thigh, axilla, and scrotum. In the final stages of the deficiency disease, muscular weakness develops and progresses within a
few days to ataxia, followed by collapse, coma, and death (HoffmannLa Roche, 2000).
Cats deficient in the vitamin develop cataracts, fatty livers, testicular hypoplasia, and alopecia with epidermal atrophy (NRC, 1986). Riboflavin deficiency manifests itself in cats after 4 to 8 weeks withanorexia, weight loss, and death. Anorexia resulting in weight loss anddeath is the principal deficiency sign in cats (Gershoff et al., 1959).
333

CHAPTER SEVEN

Chronic riboflavin deficiency in cats results in hair loss extending to thechest and feet, cataracts, and alopecia with epidermal atrophy.

FISH
Signs of riboflavin deficiency in fish are species specific (NRC,
1993). The only common signs are anorexia and poor growth. The firstsign of riboflavin deficiency observed in salmonids (Halver, 1957; Steffens, 1970) appeared in the eyes and included photophobia, cataracts,
corneal vascularization, and hemorrhages. Lack of coordinated swimming and dark skin color were also reported for riboflavin-deficient chinook salmon (Halver, 1957) and rainbow trout (Steffens, 1970). Additional signs of riboflavin deficiency in trout include slow growth,anorexia, inefficient conversion of feed, and opaque lens and cornea(NRC, 1993).
Channel catfish fed riboflavin-deficient diets developed deficiency
signs including anorexia, poor growth, short-body dwarfism, andcataracts. Common carp exhibited nervousness, photophobia, and hemorrhages, with the initial deficiency signs of anorexia and nervousness.
Signs of deficiency in Japanese eels were poor growth, anorexia, hemorrhage in fins and abdomen, photophobia, and lethargy (NRC, 1993).

FOXES AND MINK
Riboflavin deficiency in foxes results in decreased growth rate, signsof muscular weakness, chronic spasms, coma, opacity of the cornea, anddecreased pigment production in the fur (Fig. 7.8). Foxes on a riboflavin-deficient diet show signs similar to those seen in riboflavin-deficient dogs. Mink on a riboflavin-deficient diet exhibit anorexia, weight
loss, extreme weakness, and poor breeding results (NRC, 1982a).

LABORATORY ANIMALSClassic signs of riboflavin deficiency in rats are dermatitis, alopecia,
weakness, and decreased growth. Corneal vascularization and ulceration, cataract formation, anemia, myelin degeneration of sciatic nerves
and spinal cord, fatty liver, congenital abnormalities, and metabolic abnormalities of hepatocytes may occur (Fig. 7.9) (NRC, 1995). Riboflavin deficiency in rats brings about an acceleration of the erythrocyte life cycle (Gaetani and D’Aquino, 1987). In mice, signs of riboflavindeficiency include poor performance, myelin degeneration in the spinal
cord, corneal vacularization with ulceration, and lowered resistance toSalmonella infection (NRC, 1995). Riboflavin-deficient guinea pigs exhibit poor growth; rough hair coat; pale feet, nose, and ears; corneal vas334

R I B O F L AV I N

Fig. 7.8 Riboflavin deficiency in foxes. After 7
weeks on a riboflavindeficient diet, the 12week-old blue fox atright showed depigmentation, shedding of fur,and dermatitis.The littermate at left was fed
a diet supplemented
with riboflavin. (Courtesy of H. Rimeslatten,
Agriculture College of
Norway, Vollebekk.)

cularization; skin atrophy; myelin degeneration of the spinal cord; andearly death (NRC, 1995).

NONHUMAN PRIMATES
In the rhesus monkey, signs of riboflavin deficiency include “freckled”
dermatitis; incoordination; faulty grasping reflex; impaired vision; scanty
hair coat; and hypochromic, normocytic anemia. Deficiency in cebusmonkeys results in weight loss, dermatitis, alopecia, ataxia, and sudden
death, but no anemia (NRC, 1978). If not supplemented with riboflavin,the monkeys may suddenly collapse and die. Deficiency in the baboon ischaracterized by apathy, dermatitis, anemia, gingivitis, diarrhea, and adrenal cortical hemorrhage; skin lesions develop after 8 to 12 weeks.

RABBITS
Olcese et al. (1948) found that rabbits grew normally when fed pantothenic acid- and riboflavin-deficient diets; furthermore, these rabbits
excreted amounts of these vitamins greatly in excess of dietary intakes.
335

A

B

C

Fig. 7.9 Riboflavin deficiency in the rat, exhibited by (A) generalized dermatitis, growth failure, and marked keratitis of the cornea. (B) After 1 month oftreatment with riboflavin, growth resumed, and ocular and skin lesions practically disappeared. (C) After 2 months of treatment, the rat showed no signs ofdeficiency. (Courtesy of Alan T. Forrester, Scope Manual on Nutrition, The
Upjohn Company, Kalamazoo, Michigan.)336

R I B O F L AV I N

HUMANS
Riboflavin deficiency signs have been observed in humans consuming nutritionally poor diets and under experimental conditions. As in animals, riboflavin deficiency is believed to retard growth in humans, although no growth experiments have been undertaken. Clinically,
riboflavin deficiency in humans is usually observed in conjunction with
deficiencies of other B vitamins. The reason for the multiplicity of vitamin deficits is that dietary deficiencies tend to be multiple because thefood sources are similar, and various metabolic functions involve a number of vitamin interrelationships.
Clinical features of riboflavin deficiency include seborrheic dermatitis around the nose and mouth; soreness and burning of the lips, mouth,
and tongue; photophobia; burning and itching of the eyes; superficial
vascularization of the cornea; cheilosis; angular stomatitis; glossitis; anemia; and neuropathy (Fig. 7.10) (Rivlin, 1984).
Clinical signs of glossitis begin with flattening, followed by disappearance of the tongue filiform papilla. The fungiform papillae become
enlarged, the tongue color changes to a beefy red, the tongue is sore, andloss of taste sensation develops (Cooperman and Lopez, 1991). Dermatitis due to riboflavin deficiency begins most often in the nasolabial
fold and is scaly and oily. Similar lesions may also appear around theeyes and on the ears. Dermatitis of the scrotum or vulva is frequently
present (Marks, 1975). Corneal vascularization due to riboflavin deficiency always occurs in the entire circumference of the cornea and isnearly always bilateral. Corneal vascularization may be a result of theoxygen requirement of the corneal epithelium. Cells of the cornea haveno hemoglobin supply but maintain integrity by intracellular oxidative
processes that depend on riboflavin activity. Tears are a rich source of riboflavin. Foy and Mbaya (1977) stated that in vitamin A deficiency thetear ducts are blocked with keratin, and vascularization of the cornea
follows from lack of tears. Thus, such patients should be given both vitamin A and riboflavin.
One of the first systemic experimental studies in humans resulted incheilosis in 10 of 18 women consuming a low-riboflavin diet during a
94- to 130-day period (Sebrell and Butler, 1938). Cheilosis began withpallor of the lips in the angles of the mouth, followed by maceration.
Within a few days, superficial transverse fissures appeared in the anglesof the mouth.
In more recent studies, volunteers were fed a purified diet deficient
only in riboflavin, along with the riboflavin antagonist galactoflavin337

CHAPTER SEVEN

A

B

Fig. 7.10 (A) Riboflavin deficiency manifested as fissures at angle of mouth.
(B) Complete eradication after treatment with riboflavin therapy. (Courtesy of
Alan T. Forrester, Scope Manual on Nutrition, The Upjohn Company, Kalamazoo, Michigan.)(Cooperman and Lopez, 1991). Additional clinical signs of riboflavin included anemia with reticulocytopenia after 3 to 9 weeks on the diet. This
anemia was normocytic and normochromic and included leukopenia
and thrombocytopenia, with morphological bone marrow changes. Peripheral neuropathy of the hands and feet occurred, characterized by hyperesthesia, coldness, and pain. There was also decreased perception totouch, pain, temperature, vibration, and position.
In developing countries, individuals consuming diets low in green
leafy vegetables and animal products (e.g., especially milk products) are
at high risk for riboflavin deficiency. Biochemical reflections of suboptimal intake and even minor signs of deficiency are common in many
parts of the world. Reports from the 1980s include infants in New
Guinea (Oppenheimer et al., 1983) and children and pregnant womenin Gambia (Bates et al., 1983, 1984) and Nigeria (Ajayi, 1984, 1985).
Incidences of riboflavin deficiency or low status in the 1990s were reported in Bangladesh (Hassan and Ahmad, 1992), China (Brun et al.,
1990; Shaw, 1993), Egypt (Calloway et al., 1993), Lithuania (Piktelite
et al., 1992), the Philippines (Kuizon et al., 1992), Thailand (Vudhivai
et al., 1990), and Turkey (Wetherilt et al., 1992). In China (Brun et al.,1990), more than two-thirds of the population surveyed were in themedium- or high-risk category for riboflavin deficiency, while 89.9% ofchildren surveyed in Turkey (Wetherilt et al., 1992) were at risk. Defi338

R I B O F L AV I N

ciency is encountered almost invariably in combination with deficits ofother water-soluble vitamins. In fact, classic deficiency symptoms suchas glossitis and dermatitis may have resulted from other complicating
deficiencies (McCormick, 1990).
Widespread riboflavin deficiency may result in vitamin B6 deficiencies. Severe riboflavin deficiency can affect the conversion of vitamin B6
to its coenzyme (Anonymous, 1981b). The similar skin lesions seen in riboflavin and vitamin B6 deficiencies reflect impaired maturation of collagen, which has been attributed to the need for pyridoxal 5´-phosphate(enzyme form of vitamin B6), the formation of which from the 5´-phosphates of pyridoxine and pyridoxamine requires riboflavin operating as
FMN with this oxidase (Prasad et al., 1983). Dual deficiencies of riboflavin and vitamin B6 have been reported in Nigeria, Thailand, andTurkey; as an example, 89.9% of Turkish children were deficient in riboflavin and 83.4% deficient in vitamin B6 (Wetherilt et al., 1992).
Overt clinical signs of riboflavin deficiency are rarely seen among inhabitants of developed countries. However, the so-called subclinicalstage of the deficiency, characterized by a change in biochemical indices,
is common. For Italy, Mobarhan et al. (1982) reported marginal deficiency in 18% of people over 65 years of age and 12% of children. The
deficiency resulted in a shortened life span of erythrocytes. Riboflavindeficiency with biochemical changes, but not necessarily with clinicalsigns, was observed in women taking oral contraceptive agents, diabetics, children and adolescents from low socioeconomic backgrounds,
children with chronic heart disease, and the aged (Cooperman and
Lopez, 1991). In addition to major clinical signs of deficiency, retarded
intellectual development has been observed in children, and even minor
degrees of riboflavin deficiency in adults produced marked deteriorationof personality (Sterner and Price, 1973).

Assessment of Status
Several methods have been used to assess nutritional status of riboflavin. These include clinical signs, blood and urine levels of the vitamin, and measurement of enzymatic coenzyme activity. The most certain
way of identifying a riboflavin deficiency seems to be to cause a remission of the clinical signs in question by feeding riboflavin alone. Measuring riboflavin excretion in the urine is of some value, with the limitation that daily output of riboflavin is usually an estimate of dietaryintake. Riboflavin red blood cell content appears to be a better measureon the basis of human studies. For the laying hen, riboflavin concentration in egg albumen is an excellent status indicator of the vitamin339

CHAPTER SEVEN(Squires and Naber, 1993). The estimated minimum critical egg albumen
riboflavin concentrations needed to support maximum reproductivefunction are between 1.9 and 2.9 ppm.
The most rapid and dramatic biochemical method of determining riboflavin status is loss of erythrocyte glutathione reductase (EGR) activity, an FAD-containing enzyme. The ratio of EGR activity in erythrocytes with and without added FAD is considered the activity coefficient.
Experimental studies—including those with livestock, laboratory animals, humans, and even fish—have shown that EGR activity is a very
sensitive and specific indicator of riboflavin deficiency (Frank et al.,
1988; Boisvert et al., 1993; Pettigrew et al., 1996).

SUPPLEMENTATION
Riboflavin is one of the vitamins most likely to be deficient for both
nonruminant animals and humans. Before their rumens are developed,young ruminants (up to 2 months of age), if early weaned or dependenton milk replacer, have a dietary need for riboflavin. Cereal grains,though poor sources of riboflavin, are important for people in many developing countries where cereals constitute the major dietary component. Often, individuals in developing countries do not have the economic resources to consume the major dietary sources of riboflavin ofmeat, milk, and dairy products.
Supplementation and education programs for humans need to be
implemented in developing countries to ensure adequate riboflavin intake. Growing children and pregnant-lactating mothers are groups at
risk of deficiency. In India, children who received 5 mg of riboflavindaily for 1 year had elevated EGR levels, improved hand steadiness, andreduced incidence of glossitis (Prasad et al., 1990). Additional risk
groups for whom supplemental riboflavin is required are people who
exercise, alcoholics, diabetics, drug users, patients with chronic heartdisease, patients on diuretics or undergoing hemodialysis, preterm infants, and people with other disease conditions (see Requirements).
Swine and poultry diets based on grains and plant protein sources
are likewise often borderline deficient in riboflavin. Only a few feedstuffs fed to poultry and swine contain enough riboflavin to meet the requirements of growth and reproduction. The trend toward confinement
feeding and use of less pasture and/or alfalfa in swine diets has increased
need for riboflavin supplementation; both are excellent sources of the vitamin (Cunha, 1977).
Swine and poultry in confinement become more dependent on ade340

R I B O F L AV I N

quate vitamin (including riboflavin) and trace mineral supplementation,
because least-cost feed formulation (e.g., using principally corn and soybean meal) limits the number of riboflavin-rich feed ingredients (milkfermentation and fish by-products and dehydrated alfalfa). The greater
the variety of feed ingredients, the lower the chance of vitamin and traceelement deficiencies for animals and humans alike.
Riboflavin is commercially available as a crystalline compound produced by chemical synthesis or fermentation. Most of the commerciallyavailable riboflavin is made by bacterial synthesis, a cheap and convenient way to produce crystalline riboflavin. It is available to the feed,
food, and pharmaceutical industries as a high-potency U.S. Pharmacopeia (USP) or feed-grade powder, spray-dried powders, and dry dilutions. The water-soluble, riboflavin 5-phosphate salt is available for liquid oral and parenteral pharmaceuticals. High-potency USP or
feed-grade powders are electrostatic, hygroscopic, and dusty; thus they
do not flow freely, and they show poor distribution in feeds. In contrast,
dilution of riboflavin as a spray-dried powder or dry dilution product,
or including it in a premix, reduces its electrostacity and hygroscopicity
for better flowability and distribution in feeds (Adams, 1978). Diluted
riboflavin is added to feeds to increase its mixing properties.
Riboflavin is remarkably stable during heat processing (see Chemical Structure, Properties, and Antagonists; Natural Resources). However, considerable loss may occur if foods are exposed to light duringcooking, and some losses occur in feed administered to animals out ofdoors. Only the portion of the feed exposed to light would be destroyed;
therefore, this may be of little significance as only the top of concentrate
mixtures in automatic feeders would be affected. In dry form, riboflavinis extremely resistant to oxidation even when heated in air for long periods. While it has been shown that field-cured alfalfa hay exposed tomoisture can lose a significant amount of its riboflavin content in a relatively short time, under common circumstances riboflavin has goodstability when added to mixed feeds (Anonymous, 1969).
Riboflavin is stable in multivitamin premixes (Frye, 1978). Up to
26% of riboflavin in pet food is lost during extrusion (Anonymous,
1981a). Storage losses of riboflavin in pelleted feeds are slight, but after
pellets are in water for 20 minutes for fish feeding, about 40% of riboflavin may be lost (Goldblatt et al., 1979). One report demonstrated
a 98% retention of riboflavin after 6 months in a vitamin premix; however, retention was only 59% when the premix contained choline andtrace minerals (Coelho, 1991).

341

CHAPTER SEVENTOXICITY
A large body of evidence has accumulated indicating that treatment
with riboflavin in excess of nutritional requirements has very little toxicity for experimental animals or for humans (Rivlin, 1978). Most datafrom rats suggest that dietary levels between 10 and 20 times the requirement (possibly 100 times) can be tolerated safely (NRC, 1987).
When massive amounts of riboflavin are administered orally, only asmall fraction of the dose is absorbed; the remainder is excreted in thefeces. Toxicity is lacking probably because the transport system necessary for the absorption of riboflavin across the gastrointestinal mucosa
becomes saturated, limiting riboflavin absorption (Christensen, 1973).
Also, capacity of the tissues to store riboflavin and its coenzyme derivatives appears to be limited when excessive amounts are administered.
No case of riboflavin toxicity in humans has been reported, and oral
doses per kilogram of body weight of 340 mg for mice, 10 g for rats, and
2 g for dogs produced no toxic effects (Cooperman and Lopez, 1991).
Riboflavin is somewhat more toxic when administered parenterally.
Only when doses of 600 mg/kg body weight were administered intraperitoneally to rats were ill effects evident (Unna and Greslin, 1942).
Then anuria developed, and at autopsy, crystals were apparent in thecollecting tubules and renal pelvis. Rainbow trout, like other animals,
were insensitive to excess dietary riboflavin, as levels of 600 mg/kg ofdiet produced no undesirable effect on growth (Hughes, 1984).

■ REFERENCES
Adams, C. R. (1978). Proc. Roche Vitamin Nutrition Update Meeting, Arkansas
Nutrition Conference p. 54. RCD 5483/1078. Hoffmann-La Roche, Nutley,
New Jersey.
Ajayi, O. A. (1984). Hum. Nutr. Clin. Nutr. 38, 149.
Ajayi, O. A. (1985). Hum. Nutr. Clin. Nutr. 39, 383.
Ajayi, O. A., Maja, S. O., and Onabolu, Y. O. (1989). Nutr. Res. 9, 1339.
Anonymous (1969). Riboflavin No. 1170. Hoffmann-La Roche, Basel, Switzerland.
Anonymous (1981a). Rationale for Roche Recommended Vitamin Fortification
RCD 5963/1280. Hoffmann-La Roche, Nutley, New Jersey.
Anonymous (1981b). Nutr. Rev. 39, 331.
Basoglu, A., Turgut, K., Eksen, M., Tras, B., Maden, M., Ok, M., Bas, A. L., andKececi, T. (1993). Vet. Fakultesi. Dergisi-Selcuk. Universitesi. 9, 31.
Bates, C. J., Flewitt, A., Prentice, A. M., Lamb, W. H., and Whitehead, R. G.
342

R I B O F L AV I N(1983). Hum. Nutr. Clin. Nutr. 37, 427.
Bates, C. J., Prentice, A. M., and Watkinson, M. (1984). Hum. Nutr. Clin. Nutr.
38, 363.
Bazer, F. W., and Zavy, M. T. (1988). J. Anim. Sci. 66(Suppl. 1), 324(Abstr.).
Belko, A. Z., Meredith, M. P., and Kalkwarf, H. J. (1985). Am. J. Clin. Nutr. 41,
270.
Boisvert, W. A., Mendoza, I., Castaneda, C., Portocarrero, L. de, Solomons, N.
W., Gershoff, S. N., and Russell, R. M. (1993) J. Nutr. 123, 915.
Brun, T. A., Chen, J., Campbell, T. C., Boreham, J., Feng, Z., Parpia, B., Shen,
T. F., and Li, M. (1990). Eur. J. Clin. Nutr. 44, 195.
Buziassy, C., and Tribe, D. E. (1960). Aust. J. Agric. Res. 11, 989.
Calloway, D. H., Murphy, S. P., Beaton, G. H., and Lein, D. (1993). Am. J. Clin.
Nutr. 58, 376.
Carroll, F. D., Goss, H., and Howell, C. E. (1949). J. Anim. Sci. 8, 290.
Casirola, D., Kasai, S., Gastaldi, G., Ferrari, G., and Matsui, K. (1994). J. Nutr.
Sci. Vitaminol. 40, 289.
Christensen, H. N. (1993). Nutr. Rev. 51, 149.
Christensen, K. (1980). Livestock Prod. Sci. 7, 569.
Christensen, S. (1973). Acta Pharmacol. Toxicol. 32, 1.
Chung, T. K., and Baker, D. H. (1990). Poult. Sci. 69, 1357.
Clagett, C. O. (1971). Fed. Proc. Am. Soc. Expt. Biol. 30, 127.
Coelho, M. B. (1991). Vitamin Stability in Premixes and Feeds: A Practical Approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Cole, H. S., Lopez, R., and Cooperman, J. M. (1976). Acta Diabet. Latina 13,
25.
Combs Jr., G. F. (1992). The Vitamins. Academic Press, San Diego, California.
Cooperman, J. M., and Lopez, R. (1991). In Handbook of Vitamins (L.J. Machlin, ed.) 2nd Ed., p. 299. Marcel Dekker, Inc., New York.
Cunha, T. J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Das, B. S., Thurnham, D. I., Patnaik, J. K., Das, D. B., Satpathy, R., and Bose,
T. K. (1990). Am. J. Clin. Nutr. 51, 859.
Deyhim, F., Belay, T., and Teeter, R. G. (1992). Nutr. Res. 12, 1123.
Durgakumari, B., and Adiga, P. R. (1986). Mol. Cell. Endocrinol. 44, 285.
Esch, M. W., Easter, R. A., and Bahr, J. M. (1981). Biol. Reprod. 25, 659.
Fairweather-Tait, S. J., Powers, H. J., Minski, M. J., Whitehead, J., and Downes,
R. (1992). Ann. Nutr. Metab. 36, 34.
Fernando, S. M., and Murphy, P. A. (1990). J. Agr. Food Chem. 38, 163.
Flores-Garcia, W., and Scholtyssek, S. (1992). Proc. 19th World’s Poultry Congress, Vol. 1, p. 622. Beekbergen, Netherlands.
Foy, H., and Mbaya, V. (1977). Prog. Food Nutr. Sci. 2, 357.
Frank, G. R., Bahr, J. M., and Easter, R. A. (1984). J. Anim. Sci. 59, 1567.
Frank, G. R., Bahr, J. M., and Easter, R. A. (1985). J. Anim. Sci. 61(Suppl. 1),
315.
Frank, G. R., Bahr, J. M., and Easter, R. A. (1988). J. Anim. Sci. 66, 47.
Frye, T. M. (1978). Proc. Roche Vitamin Nutrition Update Meeting, Arkansas
Nutrition Conference, p. 70. RCD 5483/1078. Hoffmann-La Roche, Nutley, New Jersey.
Gaetani, S., and D’Aquino, M. (1987). Nutr. Rept. Intern. 35, 93.
343

CHAPTER SEVEN

Gao, Q. Y., Zhu, X. P., Cao, X. D., and Ma, X. X. (1993). Acta Vet. Zootech.
Sinica 24, 469.
Gershoff, S.N., Andrus, S.B., and Hegsted, D.M. (1959). J. Nutr. 68, 75.
Goldblatt, M. J., Conklin, D. E., and Brown, W. D. (1979). Proc. World Symp.
Finfish Nutr. Fishfeed Technol. 2, 117.
Grewal, P. K., and Sangha, J. K. (1990). J. Sci. Food and Agr. 52, 387.
Haenel, H., Ruttloff, H., and Ackermann, H. (1959). Biochem. Ztschr. 331,
209.
Halver, J. E. (1957). J. Nutr. 62, 225.
Hassan, N., and Ahmad, K. U. (1992). Ecol. Food Nutr. 28, 131.
Hegazy, E., and Schwenk, M. (1983). J. Nutr. 113, 1702.
Hoffmann-La Roche. (2000). In Vitamins for Dogs and Cats. Nutley, New Jersey.
Hughes, S. G. (1984). J. Nutr. 114, 1660.
Hunt, S. M. (1975). In Riboflavin (R.S. Rivlin, ed.), p. 199, Plenum Press, NewYork.
Kodentsova, V. M., Yakushina, L. M., Vrzhesinskaya, O. A., Beketova, N. A.,
and Spirichev, V. B. (1993). Voprosy-Pitaniya 5, 32.
Kuizon, M. D., Natera, M. G., Alberto, S. P., Perlas, L. A., Desnacide, J. A.,
Avena, E. M., Tajaon, R. T., and Macapinlac, M. P. (1992). Eur. J. Clin.
Nutr. 46, 257.
Kuroki, F., Iida, M., Tominaga, M., Matsumoto, T., Hirakawa, K., Sugiyama, S.,
and Fujishima, M. (1993). Di. Dis. Sci. 38, 1614.
Lee, C.M., and White, H.B. (1996). J. Nutr. 126, 523.
Levin, G., Cogan, U., Levy, Y., and Mokady, S. (1990). J. Nutr. 120, 857.
Li, L. P., Chen, W. L., and Cui, J. T. (1993). Chinese J. Chrom. 11, 49.
Loosli, J. K. (1991). In Handbook of Animal Science, (P.A. Putnam, ed.), p. 25.
Academic Press, San Diego, California.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease.
Medical and Technical Publishing Co., Ltd., p. 73. Lancaster, England.
McCollum, E. V. (1957). In A History of Nutrition p. 291. The Riverside Press,
Cambridge, Massachusetts.
McCormick, D. B. (1990). In Nutrition Reviews, Present Knowledge in Nutrition (M.L. Brown, ed.), 6th Ed., p. 146. International Life Sci. Inst., Washington, D.C.
McElroy, L. W., and Goss, H. (1940). J. Nutr. 20, 527.
Middleton, H. M. (1990). J. Nutr. 120, 588.
Miller, B. L., Goodrich, R. D., and Meiske, J. C. (1983a). J. Anim. Sci. 57(Suppl.
1), 454(Abstr.).
Miller, B. L., Plegge, S. D., Goodrich, R. D., and Meiske, J. C. (1983b). Minnesota Beef Report B-299, p. 1. University of Minnesota, St. Paul.
Mobarhan, S., Maiani, G., Zanacchi, E., Ferrini, A. M., Scaccini, C., Sette, S.,
and Ferro-Luzz, A. (1982). Clin. Nutr. 360, 71.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
344

R I B O F L AV I N(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Olcese, O., Pearson, P. B., and Schweigert, B. S. (1948). J. Nutr. 35, 577.
Onwudike, O. C., and Adegbola, A. A. (1984). Trop. Agr. (Trinidad) 61, 205.
Oppenheimer, S. J., Bull, R., and Thurnham, D. J. (1983). P N G Med. J. 26, 17.
Pearson, P. B., Sheybani, M. K., and Schmidt, H. (1944). J. Anim. Sci. 3, 166.
Pettigrew, J. E., El-Kandelgy, S. M., Johnston, L. J., and Shurson, G. C. (1996).
J. Anim. Sci. 74, 2226.
Piktelite, O. S., Aleinik, S. I., Yakushina, L. M., and Blazheevich, N. V. (1992).
Voprosy-Pitaniya 4, 32.
Pinto, J., Huang, Y. P., and Rivlin, R. S. (1981). J. Clin. Invest. 67, 1500.
Powers, H. J., Weaver, L. T., Austin, S., Wright, A. J. A., and Fairweather-Tait,
S. J. (1991). Br. J. Nutr. 65, 487.
Powers, H. J., Weaver, L. T., Austin, S., and Beresford, J. K. (1993). Br. J. Nutr.
69, 553.
Prasad, P., Lakshmi, A. V., and Bamji, M. S. (1983). Biochem. Med. 30, 333.
Prasad, P., Bamji, M. S., Lakshmi, A. V., and Satyanarayana, K. (1990). Nutr.
Res. 10, 275.
Radostits, O. M., and Bell, J. M. (1970). Can. J. Anim. Sci. 50, 405.
Raghvendar, S., Deodhar, A. D., and Singh, R. (1993). Indian J. Dairy Sci. 46,
525.
RDA. (1989). Recommended Dietary Allowances, 10th Ed., National Academyof Sciences-National Research Council, Washington, D.C.
Rivlin, R. S. (1978). In Handbook Series in Nutrition and Food, Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 1, p. 25. CRC Press, Boca Raton, Florida.
Rivlin, R. S. (1984). In Nutrition Reviews, Present Knowledge in Nutrition (R.E.
Olson, ed.), p. 285. The Nutrition Foundation, Washington, D.C.
Roe, D. A., Bogusz, S., Sheu, J., and McCormick, D. B. (1982). Am. J. Clin.
Nutr. 35, 495.
Rose, R. C., McCormick, D. B., Li, T. K., Lumeng, L., Haddad, J. G., and Spector, R. (1986). Fed. Proc. Am. Soc. Expt. Biol. 45, 30.
Ruiz, N., and Harms, R. H. (1988). Poult. Sci. 67, 794.
Russell, L. F., and Vanderslice, J. T. (1991). Food Chem. 43(2), 151.
Said, H. M., Hollander, D., and Mohammadkhani, R. (1993a). Biochim. Bio345

CHAPTER SEVEN

phys. Acta 1148, 263.
Said, H. M., Mohammadkhani, R., and McCloud, E. (1993b). Proc. Soc. Exp.
Biol. Med. 202, 428.
Scott, N. L., Nesheim, M. C., and Young, R. J. (1982). Nutrition of the Chicken.
Scott, Ithaca, New York.
Sebrell, W. H., and Butler, R. E. (1938). Public Health Rep. 53, 2282.
Seymour, E. W., Speer, V. C., and Hays, V. W. (1968). J. Anim. Sci. 27, 389.
Sharman, I. (1977). Endeavour 1, 97.
Shaw, N. S. (1993). Nutr. Res. 13, 147.
Sigman-Grant, M., Bush, G., and Anantheswaran, R. (1992). Pediatrics 90, 412.
Singh, R., and Deodhar, A. D. (1992). Ann. Nutr. Metab. 36, 279.
Squires, M. W., and Naber, E. C. (1993). Poult. Sci. 72, 483.
Steffens, W. (1970). Int. Rev. Ges. Hydrobiol. 59, 255.
Sterner, R. T., and Price, W. R. (1973). Am. J. Clin. Nutr. 26, 150.
Trebler-Winters, L. R., Yoon, J. S., Kalkwarf, H. J., Davies, J. C., Berkowitz, M.
G., Haas, J., and Roe, D. A. (1992). Am. J. Clin. Nutr. 56, 526.
Turkki, P. R., and Holtzapple, P. G. (1982). J. Nutr. 112, 1940.
Unna, K., and Greslin, J. G. (1942). J. Pharmacol. Exp. Ther. 76, 75.
Vudhivai, N., Pongpaew, P., Prayurahong, B., Kwanbunjan, K., Migasena, P.,
Chitwattanakorn, M., Hempfling, A., and Schelp, F. P. (1990). Int. J. Vitam.
Res. 60, 75.
Wagner-Jauregg, T. (1972). In The Vitamins (W.H. Sebrell and R. S. Harris,eds.), p. 3. Academic Press, New York.
Wetherilt, H., Ackurt, F., Brubacher, G., Okan, B., Aktas, S., and Turdu, S.
(1992). Int. J. Vitam. Nutr. Res. 62, 21.
White, H.B. (1996). J. Nutr. 126, 1303S.
Zempleni, J., Link, G., and Kubler, W. (1992). Int. J. Vitam. Nutr. Res. 62, 165.
Zintzen, H. (1975). A Guide to the Nutritional Management of Breeding Sows
and Pigs. No. 1465. Hoffmann-La Roche, Basel, Switzerland.

346

chapter eightNIACININTRODUCTION
After isolation of thiamin and riboflavin from the vitamin B complex, niacin was the third B vitamin to be established and is sometimes
referred to as vitamin B3. Niacin exerts its major physiological effects
through its role in the enzyme system for cell respiration. Niacin deficiency results in the disease pellagra in humans and blacktongue in dogs,
and in 1937, scientists discovered that this vitamin would cure these
conditions. Soon after, niacin was found to be a dietary essential for
pigs, poultry, and other nonruminant animals.
Although the need for niacin for simple-stomached animals has been
established, exact requirements are difficult to determine since the vitamin can be synthesized from the amino acid tryptophan. In many
species, a single deficiency of only niacin does not exist; rather the deficiency condition is an inadequate intake of both niacin and tryptophan.
The vitamin can also be considered a drug when administered at pharmacological levels.

HISTORY
Niacin has been known to organic chemists since 1867, long before
its importance as an essential nutrient was recognized. The history of
niacin and the significance of its deficiency have been reported (Harris,
1919; McCollum, 1957; Darby et al., 1975; Hankes, 1984; Anonymous,
1987; Loosli, 1991). As early as 1911 to 1913, Funk had isolated it fromyeast and rice polishings in the course of an attempt to identify the water-soluble anti-beriberi vitamin. But interest in niacin was lost when it
was found ineffective in curing pigeons of beriberi. Although Funk
347

8

CHAPTER EIGHT

found that niacin did not cure beriberi, cures were more rapid when it
was administered in conjunction with the concentrates containing theanti-beriberi vitamin (thiamin). Warburg and coworkers first demonstrated a biochemical function for nicotinic acid when they isolated itfrom an enzyme in 1935 and showed that it functioned as part of a hydrogen transport system.
Niacin deficiency in humans has been equated with pellagra, thedisease considered a condition of the corn (maize)-eating population.
Corn was brought to Spain by Columbus from America. Pellagra was
not recognized until 1735 when Gaspar Casal, physician to Philip V of
Spain, observed it in northern Spain. The local people had called it mal
de la rosa, and Casal associated the disease with poverty and spoiled
corn. The popularity of corn, and thus pellagra, spread eastward from
Spain to southern France, Italy, Romania, Russia, and Egypt. Although
the disease was first reported in Spain, most of the important early studies of the disease came from Italy. Pellagra was so prevalent in northern
Italy that the first pellagra hospital was established at Legano in 1784.
Babcock, of Columbia, South Carolina, who established the identityof U.S. pellagra as the same disease in Italy, studied the case records of
the South Carolina State Hospital and concluded that the disease occurred there as early as 1828. Most cases occurred in low-income
groups, whose diet was limited to cheap foodstuffs. Diets characteristically associated with the disease were referred to as the three Ms—meal(corn), meat (back fat), and molasses—plus poverty.
In Spartanburg, South Carolina, a Public Health Service Hospitalfor the care and study of pellagra was in operation from 1917 to 1919.
However, the hospital was closed in 1919 because there were no casesof pellagra, emphasizing the relationship between poverty and pellagra.
Pellagra virtually disappeared from the southern United States as a result of the great increase in the price of cotton during and at the end ofWorld War I. Prosperity made a wider choice of food available to all.
Pellagra rapidly reappeared when the price of cotton fell and poverty
again forced greater reliance on corn-based diets.
The word pellagra means rough skin, which would be dermatitis.
Other descriptive names for the condition were mal del sol (illness of thesun) and “corn bread fever.” After the turn of the century in the UnitedStates, particularly the South, it was common for 20,000 deaths to occur annually from pellagra. It was estimated that for every death due topellagra, there were at least 35 cases of the disease. Even as late as 1941,
which was 5 years after the cause of pellagra was known, 2,000 deaths
due to the disease were reported. Clinical signs and mortality from pel348

NIACIN

lagra could be referred to as the four Ds: dermatitis (of areas exposed tothe sun), diarrhea, dementia (mental problems), and death. Several mental institutions in the United States, Europe, and Egypt were primarily
devoted to care of pellagrins (Darby et al., 1975).
In 1914, Voegtlin reported the earliest study which proved that human pellagra is unquestionably caused by a dietary deficiency. He tested
the effects on pellagrins receiving two contrasting diets; those receiving
eggs, meat, and milk were cured, while those remaining on corn-based
diets still had the disease. Also in 1914, Goldberger, a bacteriologist in
the United States Public Health Service, was assigned the task of identifying the cause of pellagra. In his studies he observed that the disease
was associated with poor diet and poverty and that well-fed persons did
not contract the disease. In orphanages, prisons, and mental institutions
in South Carolina, Georgia, and Mississippi the therapeutic value ofgood diets was demonstrated. Goldberger, his wife, and 14 volunteers
constituted a “filth squad” who ingested and were injected with various
biological materials and/or excreta from pellagrins, thus demonstrating
the noninfectious nature of pellagra. At this time researchers and physicians did not want to believe that pellagra resulted from poor nutrition,
but rather from an infection based on the popularity of the “germ theory” of disease. An important step toward the isolation of the preventive factor for pellagra was discovery of a suitable laboratory animal for
testing its potency in various concentrated preparations. It was foundthat a pellagra-like disease (blacktongue) could be produced in dogs.
After it had been established that pellagra was a disease of nutritional deficiency, the next step was to discover the missing nutrient. Additional dietary protein was shown to be beneficial, so it was concluded
that pellagra was due to protein deficiency. This view and, later, that it
was more specifically due to a deficiency of the amino acid tryptophan,
was held for some time. Following discovery that a crude extract of liver
was effective in curing pellagra, and therefore was a source of the preventive factor, Elvehjem and colleagues (1937) isolated nicotinamide
from liver as the factor that would cure blacktongue in dogs. Reports ofthe dramatic therapeutic effects of niacin in human pellagra quickly followed from several clinics.
In 1945, Krehl and coworkers found that tryptophan was as active
as niacin in treatment of pellagra. Positive proof that tryptophan wasconverted to nicotinic acid was published by Heidelberger, who showedthat the L-[14C]tryptophan was converted to 14C-labeled nicotinic acidin the rat. The conversion of tryptophan to niacin explained why foods
rich in animal protein (e.g., milk) prevented and cured pellagra.
349

CHAPTER EIGHTCHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Chemically, niacin (C6H5O2N) is one of the simplest vitamins. The
two forms of niacin—nicotinic acid and nicotinamide—correspond to 3pyridine carboxylic acid and its amide, respectively (Fig. 8.1). The term
niacin is used as a generic descriptor of pyridine 3-carboxylic acid andderivatives exhibiting the same qualitative biological activity of nicotinamide. There are antivitamins or antagonists for niacin. These compounds have the basic pyridine structure, with two of the important antagonists of nicotinic acid being 3-acetyl pyridine and pyridine sulfonicacid. Nicotinic acid and nicotinamide (niacinamide) possess the same vitamin activity; the free acid is converted to the amide in the body.
Nicotinamide functions as a component of two coenzymes: nicotinamide adenine dinucleotide (NAD) and nicotinamide adenine dinucleotide phosphate (NADP) (Fig. 8.1).
Both nicotinic acid and nicotinamide are white, odorless, crystalline
solids soluble in water and alcohol. They are very resistant to heat, air,light, and alkaline conditions and thus are stable in foods. Niacin is alsostable in the presence of the usual oxidizing agents. However, it will undergo decarboxylation at a high temperature when in an alkaline
medium. Nicotinic acid readily forms salts with metals such as aluminum, calcium, copper, and sodium. When in acid solution, niacin
readily forms quaternary ammonium compounds, such as nicotinic acidhydrochloride, which is soluble in water. When in a basic solution, nicotinic acid readily forms carboxylic acid salts.

ANALYTICAL PROCEDURES
The most sensitive method for the determination of niacin andclosely related compounds is microbiological. Lactobacillus plantarum
responds to both forms of the vitamin, whereas Leuconostoc mesenteroides measures only nicotinic acid. Niacin must be freed from boundforms before assay. Since niacin is very stable to strong acids, it can bereleased by acid hydrolysis. Chemical methods of analysis are less sensitive than microbiological procedures and generally require more extensive extraction methods. The cyanogen bromide method of analysis isbased on the reaction of pyridine derivatives with cyanogen bromide,
forming a colored compound that can be used for quantitative measurement of the vitamin. The active coenzyme forms of niacin, the pyridine
350

NIACIN

Fig. 8.1 Chemical structures of nicotinic acid, nicotinamide, nicotinamide adenine dinucleotide (NAD), and nicotinamide adenine dinucleotide phosphate(NADP).

nucleotides NAD and NADP, can be determined by an enzyme-cycling
colorimetric procedure (Nisselbaum and Green, 1969) or high-performance liquid chromatography (HPLC) (Stocchi et al., 1987). Bioassay
procedures for niacin present two major difficulties in that (1) tryptophan in the diet is converted to niacin in the tissues, and (2) niacin is synthesized by intestinal bacteria to varying degrees. Chicks, puppies, andweanling rats have been used for biological niacin assay.

METABOLISM
Absorption, Transport, and Storage
Nicotinic acid and its amide are readily and very efficiently absorbed

351

CHAPTER EIGHT

by diffusion at either physiological or pharmacological doses. At low
concentrations absorption occurs as an Na+-dependent facilitated diffusion, but at higher concentrations passive diffusion predominates. By
employing the gastrointestinal tube technique, niacin was shown to beequally well absorbed from both the stomach and upper small intestinein humans (Bechgaard and Jespersen, 1977). In a steady-state situation,
approximately 85% of a dose of 3 g of niacin per day was recovered fromthe urine of humans, and absorption was considered almost complete. In
cattle studies, nicotinamide was more rapidly absorbed from the rumen
than nicotinic acid (Erickson et al., 1991). This may have been due to differences in the dissociation constants of the compounds, with significanceof ruminal fermentation and metabolism unknown.
Niacin in foods occurs mostly in its coenzyme forms, which are hydrolyzed during digestion, yielding nicotinamide, which seems to be absorbed without further hydrolysis in the gastrointestinal tract. The intestinal mucosa is rich in niacin conversion enzymes such as NAD
glycohydrolase (Henderson and Gross, 1979). Nicotinamide is released
from NAD in the liver and intestines by glycohydrolases for transport totissues that synthesize NAD as needed. In the gut, mucosa nicotinic acidis converted to nicotinamide (Stein et al., 1994). Nicotinamide is the primary circulating form of the vitamin and is converted into its coenzyme
forms in the tissues.
The tissue content of niacin and its analogs, NAD and NADP, isvariable, dependent on the diet and a number of other factors, such as
strain, sex, age, and treatment of animals (Hankes, 1984). Althoughniacin coenzymes are widely distributed in the body, no true storage occurs. Liver contains the greatest niacin concentration, but the amountstored is not great.

Excretion
Urine is the primary pathway of excretion of absorbed niacin and its
metabolites. At high dosages, the half-life of both nicotinic acid andnicotinamide is determined mainly by rate of excretion of the unchanged
compound in urine and not by metabolic change. When low dosages areused, both compounds are excreted principally as metabolites rather
than as unchanged compounds.
The principal excretory product in humans, dogs, rats, and pigs isthe methylated metabolite N′-methylnicotinamide or one of two oxidation products of this compound, 4-pyridone or 6-pyridone of N′-methylnicotinamide. On the other hand, in herbivores niacin does not seem to352

NIACIN

be metabolized by methylation, but large amounts are excreted unchanged. In the chicken, however, nicotinic acid is conjugated with ornithine as either α- or δ-nicotinyl ornithine or dinicotinyl ornithine. The
excretion of these metabolites is measured in studies of niacin requirements and metabolism. Such studies are complicated by the fact that thekinds and relative amounts of these products vary with the species andlevel of niacin intake.

Tryptophan-Niacin Conversion
The amino acid tryptophan is a precursor for the synthesis of niacinin the body, and there is considerable evidence that synthesis can takeplace in the intestine. Investigations indicate that urinary excretion of
the metabolite N′-methylnicotinamide is much larger when tryptophan
is administered by stomach tube than when injected parenterally. Thereis also evidence that synthesis occurs in the developing chick embryo.
The extent to which the metabolic requirement for niacin can be met
from tryptophan will depend, first, on the amount of tryptophan in thediet and its bioavailability and, second, on the efficiency of the conversion of tryptophan to niacin. The pathway of tryptophan conversion tonicotinic acid mononucleotide in the body is shown in Fig. 8.2. Protein,energy, vitamin B6, and riboflavin nutritional status and hormones affectone or more steps in the conversion sequence shown in Fig. 8.2, andhence can influence the yield of niacin from tryptophan. In addition,copper, iron, and magnesium all have roles in the transformation. Ironis required by two enzymes for the conversion of tryptophan to niacinwith a deficiency reducing tryptophan utilization (Oduho et al., 1994).
It is suggested that pellagra is not simply a disease of niacin deficiency,
but more appropriately a disease of tryptophan metabolism (Anonymous, 1987).
At low levels of tryptophan intake, the efficiency of conversion ishigh. It decreases when niacin and tryptophan levels in the diet are increased. When animals have received inadequate levels of tryptophan,increasing amounts of dietary tryptophan are used first to restore nitrogen balance, next to restore blood pyridine nucleotides, and then to be
excreted as niacin metabolites. Under starvation or energy restriction,
efficiency of conversion increases, while pregnant women can convert
tryptophan to niacin more efficiently than other adults (Narasinga Rao
and Gopalan, 1984). A threefold increase in conversion efficiency wasreported in pregnant women during the third trimester (Wertz et al.,1958). This increase presumably was due to the stimulation by estrogen353

CHAPTER EIGHT

Fig. 8.2 Conversion of tryptophan to nicotinic acid mononucleotide plus someside reactions.

of tryptophan oxygenase, a suggested rate-limiting enzyme in the pathway (Rose and Braidman, 1971). Another factor that may affect thisconversion ratio is amino acid imbalance due to excess leucine in the diet(see Deficiency, Humans).
Animal species differ widely in ability to synthesize niacin fromtryptophan. According to a variety of experiments, approximately 60
mg of tryptophan is estimated to be equivalent to 1 mg of niacin in humans, while the rat is more efficient, with a conversion rate of 35 to 50
mg of tryptophan required. Conversion efficiency of tryptophan to354

NIACIN

niacin in the chick is estimated to be 45:1 (Baker et al., 1973; Chen andAustic, 1989) and relatively efficient, while in the turkey it is inefficient,with conversions ranging from 103 to 119:1 (Ruiz and Harms, 1990c).
Conversion efficiency is probably due to inherent differences in liver levels of picolinic acid carboxylase, the enzyme that diverts one of the intermediates (2-amino-3-acroleyl fumaric acid) toward the glutaryl-CoA
pathway instead of allowing this compound to condense to quinolinicacid, the immediate precursor of nicotinic acid. Picolinic acid carboxylase in livers of various species (Table 8.1) has a very close inverse relationship to experimentally determined niacin requirements. The cat has
been shown to have 30 to 50 times as much picolinic acid carboxylase
activity as the rat. Thus, the cat has so much of this enzyme that it cannot convert any of its dietary tryptophan to niacin. The cat, therefore,
has an absolute requirement for niacin itself. Conversely, the rat diverts
very little of its dietary tryptophan to carbon dioxide and water and thus
is very efficient in converting tryptophan to niacin. The duck has a very
high niacin requirement (approximately two times that of chickens),
with considerably higher levels of picolinic acid carboxylase activity(Scott et al., 1982).

FUNCTIONS
The major function of niacin is in the coenzyme forms of nicotinamide, NAD and NADP. Enzymes containing NAD and NADP are important links in a series of reactions associated with carbohydrate, protein, and lipid metabolism. They are especially important in themetabolic reactions that furnish energy to the animal. The coenzymes
act as an intermediate in most of the H+ transfers in metabolism, including more than 200 reactions in the metabolism of carbohydrates,
fatty acids, and amino acids. These reactions are of paramount importance for normal tissue integrity, particularly of the skin, gastrointestinal
tract, and nervous system.
The NAD- and NADP-containing enzymes play an important role inbiological oxidation-reduction systems by virtue of their capacity toserve as hydrogen-transfer agents. Hydrogen is effectively transferredfrom the oxidizable substrate to oxygen through a series of graded enzymatic hydrogen transfers. Nicotinamide-containing enzyme systems
constitute one such group of hydrogen-transfer agents. In electrontransport, oxidation of reduced NAD or NADP is carried out by a second hydrogen-carrying system, the riboflavin coenzymes (see Chapter 7,355

CHAPTER EIGHT■ Table 8.1

Liver Picolinic Acid Carboxylase Activity Among Species

SpeciesCat
LizardDuck
Frog
TurkeyCow
Pig
PigeonChicken (high niacin-requiring strain)
Rabbit
Mouse
Guinea pig
Chicken (strain with low niacin requirement)
Human
Hamster
Toad
Rat

Units/g liver (wet basis)
50,50029,64017,33013,720
9,230
8,300
7,120
6,950
5,380
4,270
4,200
3,940
3,200
3,180
3,140
3,050
1,570

Sources: Modified from Ikeda et al. (1965), DiLorenzo (1972), and Scottet al. (1982).

Fig. 7.2). The nicotinamide moiety of the coenzyme operates in thesesystems by reversibly alternating between an oxidized quaternary pyridinium ion and a reduced tertiary amine. The transfer of hydrogen is reversible and stereospecific.
Some of these enzymes show a strict specificity for either NAD orNADP, while others utilize these coenzymes equally well. Important
metabolic reactions catalyzed by NAD and NADP are summarized below:
1. Carbohydrate metabolism—(a) glycolysis (anaerobic oxidation of
glucose) and (b) the Krebs cycle.
2. Lipid metabolism—(a) glycerol synthesis and breakdown, (b) fatty
acid oxidation and synthesis, and (c) steroid synthesis.
3. Protein metabolism—(a) degradation and synthesis of amino acids
and (b) oxidation of carbon chains via the Krebs cycle.
4. Photosynthesis.
5. Rhodopsin synthesis (see Chapter 2, Fig. 2.6).

NAD is specific for hydrogenases involved in passing electrons on tooxygen via the electron-transport system in the tricarboxylic acid cycle.
Here NAD serves as the electron acceptor in three of the four dehydrogenation steps. As a hydrogen acceptor, NADH is formed which in turn
356

NIACIN

functions as a hydrogen donor to the respiratory chain for ATP production; NAD as well participates as a codehydrogenase, with enzymes involved in the oxidation of fuel molecules such as glyceraldehyde 3-phosphate, lactate, alcohol, 3-hydroxybutyrate, pyruvate, andα-ketoglutarate dehydrogenases. NADP also participates in dehydrogenation reactions, particularly in the hexose monophosphate shunt of
glucose metabolism. Reduced NADP has an important role in the synthesis of fats and steroids and is contained in the alcohol-dehydrogenasesystem, the lactic acid-dehydrogenase system, and other systems.
Both NAD and NADP are involved in degradation and synthesis ofamino acids. NAD has a role, as the source of ADP-ribose for the ADPribosylation of proteins and poly(ADP-ribose). ADP-ribosyltransferases
are enzymes of the cytosol, plasma membrane, and nuclear envelope,which catalyze the transfer of ADP-ribose onto arginine, lysine, or asparagine residues in acceptor proteins to form N-glycosides.
Niacin-dependent poly(ADP-ribose) is involved in the post-translational modification of nuclear proteins. The poly ADP-ribosylated proteins seem to function in DNA repair, DNA replication, and cell differentiation (Carson et al., 1987). In the event of DNA strand breaks,
ADP-ribose subunits, cleaved from NAD+, are covalently linked toamino acid residues, and the polymer is subsequently elongated andbranched. Poly(ADP-ribose) is synthesized in response to DNA strand
breaks. Rat data have shown that even mild niacin deficiency decreases
liver poly(ADP-ribose) concentrations and that poly(ADP-ribose) levels
are also altered by food restriction (Rawling et al., 1994). Zhang et al.
(1993) suggested that severe niacin deficiency may increase the susceptibility of DNA to oxidative damage, likely due to lower availability ofNAD. Turnover rates of protein in Japanese quail have been related toniacin deficiency, and a high turnover rate due to the deficiency was primarily attributed to enhanced degradation rate of proteins rather thanenhanced synthesis rate of proteins (Park et al., 1991).
Niacin was found to be part of the glucose tolerance factor, anorganochromium complex isolated from yeast that potentiates the insulin
response (Mertz, 1975). The role of the niacin moiety in the glucose tolerance factor is unknown because free niacin has no similar effect.

REQUIREMENTS
Niacin requirements for various animals and humans are presented
in Table 8.2. Niacin requirements vary widely, generally because (1)niacin is synthesized from tryptophan, and (2) much of dietary niacin is357

CHAPTER EIGHT■ Table 8.2

Niacin Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100-g intake)
Broilers, all classes
All classes
All classes
All classes
Adult
Growing-finishing, 3–5 kg
Growing-finishing, 5–120 kg
Adult
Adult
Adult
Growing
Growing
GrowingCatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
Children
AdultsMicrobial synthesis
2.6 ppm milk replacer
Microbial synthesis
27.0 mg/kg
11.0 mg/kg
10.0 mg/kg
25–30 mg/kg
55 mg/kg
20–40 mg/kg
40–60 mg/kg
Microbial synthesis
20 mg/kg
7–15 mg/kg
10.0 mg/kg
Microbial synthesis
Microbial synthesis
10 mg/kg
40 mg/kg
450 μg/kg body wt.
14 mg/kg
150–200 mg/kg
1–10 mg/kg
15 mg/kg
15 mg/kg
5–6 mg/day
9–13 mg/day
17–20 mg/day

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989)
NRC (1981b)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

ChickenDuck (Pekin)Japanese quail
Turkey
Sheep
Swine
Horse
Goat
Fox
Cat
Dog
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg/day.

in a bound form unavailable to humans and animals.
Some species, such as the cat, mink, and most fish, apparently lackthe ability to convert tryptophan to niacin. For species that have the capacity to synthesize niacin from tryptophan, it is impossible to set theniacin requirement unless the tryptophan level is specified and it is
known that the diet is adequate in vitamin B6, since this vitamin isneeded in synthesis of niacin from tryptophan. However, the vitamin B6
level is adequate in most practical livestock diets. Riboflavin is required
for synthesis of niacin from tryptophan as well as for conversion of vitamin B6 to the functional coenzyme (pyrodoxal phosphate), which isneeded for niacin synthesis (see Metabolism; Chapter 7, Functions).
Likewise, many other nutrients and hormones affect conversion of
niacin from tryptophan (see Metabolism).
Cunha (1982) listed a number of factors that influence niacin requirements for livestock, as follows:
358

NIACIN

1. Genetic differences, which can influence niacin needs, such as selection for meatier, faster-growing animals and increased production levels.
2. Ability to synthesize niacin from tryptophan (see Metabolism).
3. Increased stress and subclinical disease level on the farm because of
closer and more frequent contact between animals in confinement.
4. Trend toward more intensified operations that may lessen opportunity for coprophagy and will reduce access to pasture.
5. Newer methods of handling and processing feeds that may affectniacin and tryptophan level and availability.
6. Various nutrient interrelationships, including amino acid imbalances.
7. Trend toward earlier weaning, which increases the need for higher vitamin levels in milk-substitute diets (prestarter and starter feeds).
8. Molds and antimetabolites in feeds that can increase certain nutrient
needs.

The type of carbohydrate consumed may influence niacin requirements. For maximal growth, tilapia fingerlings required 26 ppm niacin
when fed a glucose diet compared to 121 ppm for those fish fed a dextrin (from corn) diet (Shiau and Suen, 1992).
Niacin requirements of ruminants are unknown; the vitamin is normally synthesized in adequate quantities in the rumen. Niacin supplementation has been shown to be beneficial in beef cattle receiving highconcentrate diets and in high-producing dairy cows. In a review,
Olentine (1984) summarized the factors affecting ruminant niacin requirements as follows:
1. Protein balance—excess of leucine, arginine, and glycine increases therequirement.
2. High tryptophan content of feeds—as tryptophan content increases,
niacin requirements decrease.
3. Energy content—high-energy rations require more niacin per unit offeed.
4. Antibiotics—depending on the product, niacin requirements are increased.
5. Dietary rancidity—if fat is rancid, niacin requirements are increased.
6. Gastrointestinal synthesis—niacin is synthesized in the gastric and intestinal regions.
7. Availability of niacin and tryptophan in feedstuffs—cereal grains
and other feedstuffs have varying degrees of niacin and tryptophan availability.
359

CHAPTER EIGHT

Human niacin requirements are based on niacin intake of pellagrins
and on human depletion and repletion studies. Most food tables for humans list niacin as niacin equivalents (NEs), which represent the sum of
milligrams of niacin (nicotinic acid plus nicotinamide) plus 1/60 mg oftryptophan (conversion rate of tryptophan to niacin). Niacin equivalents
range from 13 to 19 mg for adults or 6.6 mg/1,000 kcal (RDA, 1989).
This amount provides for differences in various diets consumed in termsof the bioavailability of niacin and the contribution of tryptophan. The
Food and Agriculture Organization/World Health Organization Expert
Group recommended an intake of 6.6 mg NE per 1,000 kcal for all age
groups (Narasinga Rao and Gopalan, 1984). The RDA (1989) provided
an increase of 2 and 5 mg NE for pregnancy and lactation, respectively.
Exercise (Haralambie, 1976), diabetes mellitus (Vague et al., 1987),
and genetic disorders such as Hartnup’s disease (Halvorsen andHalvorsen, 1963) can increase niacin requirements in humans. In Hartnup’s disease there is impaired intestinal absorption of tryptophan(Halvorsen and Halvorsen, 1963), but in diabetes there are elevated concentrations of picolinic acid carboxylase, resulting in less efficient conversion of niacin from tryptophan (Ikeda et al., 1965).

NATURAL SOURCES
Niacin is widely distributed in foods of both plant and animal origin (Table 8.3). Large quantities of niacin are found in brewer’s yeast
and meats. Animal and fish by-products, distiller’s grains and yeast, various distillation and fermentation solubles, and certain oil meals aregood sources. Leafy materials, especially pasture grass, are fair sources.
Milk, dairy products, fruits, and eggs are poor sources. Niacin is present in uncooked foods mainly as the pyridine nucleotides NAD andNADP, but some hydrolysis of these nucleotides to free forms may occur during food preparation.
Most species can use the essential amino acid tryptophan and synthesize niacin from it (see Metabolism and Requirements sections). Because tryptophan can give rise to body niacin, both niacin and tryptophan content should be considered in expressing niacin values of foods.
However, since there is a preferential use of tryptophan for protein synthesis before any becomes available for conversion to niacin (Kodicek etal., 1974), it seems unlikely, given the low tryptophan content in many
feedstuffs, that tryptophan conversion greatly contributes to the niacin
supply.
360

NIACIN■ Table 8.3

Niacin in Foods and Feedstuffs (mg/kg, dry basis)

Alfalfa hay, sun cured
Alfalfa leaves, sun cured
Barley, grain
Bean, navy (seed)
Blood meal
Brewer's grains
Buttermilk (cattle)
Chicken, broilers (whole)
Citrus pulp
Clover hay, ladino (sun cured)
Copra meal (coconut)
Corn, gluten meal
Corn, yellow grain
Cottonseed meal, solvent extracted
Fish meal, anchovy
Fish meal, menhaden
Fish, sardine
Linseed meal, solvent extracted
Liver, cattle
Milk, skim, cattle

42
53
94
28
34
47
9
230
23
11
28
55
28
48
89
60
81
37
269
12

Molasses, sugarcane
Oat, grain
Pea seeds
Peanut meal, solvent extracted
PotatoRice, branRice, grain
Rice, polished
Rye, grain
Sorghum, grain
Soybean meal, solvent extracted
Soybean seedSpleen, cattle
Timothy hay, sun cured
Wheat, bran
Wheat, grain
Whey
Yeast, brewer's
Yeast, torula

49
16
36
188
37
330
39
17
21
43
31
24
25
29
268
64
11
482
525

Source: NRC (1982b).

Niacin is often present in feeds in a bound form that is not available.
The niacin in cereal grains and their by-products is in a bound, complex
form that is virtually unavailable, at least to simple-stomached animals(Luce et al., 1967). It seems that much of this niacin is also unavailable
to rumen microorganisms. Any dietary niacin escaping degradation in
the rumen will also likely be unavailable for absorption in the lower gut.
Initial evidence of bound forms of niacin stemmed from observed
discrepancies between niacin values in cereals determined colorimetrically before and after hydrolysis with dilute NaOH. Microbiological assays similarly indicated that about 20% more niacin was obtained in dilute alkaline extracts of cereals than in aqueous or acid extracts.
Two types of bound niacin were initially described: (1) a peptide
with molecular weight of 12,000 to 13,000, the so-called niacinogens,and (2) a carbohydrate complex with a molecular weight of approximately 2,370 (Darby et al., 1975). The name niacytin has been used todesignate this latter material from wheat bran. A number of boundforms of niacin are apparently present in cereals. Ghosh et al. (1963),
using a microbiological assay, reported that 85 to 90% of the total
nicotinic acid in cereals is in a bound form. Mason et al. (1973) reported that bound niacin is linked to the macromolecules, of which
about 60% were polysaccharides and 40% peptides or glycopeptides.
Lack of niacin availability may be due to a blocking effect of these mol361

CHAPTER EIGHT

ecules, which prevent access of digestive enzymes to the niacin-macromolecule bond, and therefore free niacin would not be available for absorption.
About 40% of the total niacin in oilseeds is in bound form, while
only a small proportion of the niacin in pulses, yeast, crustacea, fish, animal tissue, or milk is bound. Using a rat assay procedure, Carter andCarpenter (1982) showed that in eight samples of mature cooked cereals (corn, wheat, rice, and milo), only about 35% of the total niacin wasavailable. In the calculation of the niacin content of formulated diets,
probably all niacin from cereal grain sources should be ignored or atleast given a value no greater than one-third of the total niacin.
Some bound forms of niacin are biologically available, but theniacin in corn is particularly unavailable and is implicated in the etiology of pellagra in societies that consume large quantities of the grain.
Pellagra occurs in Mexico less frequently than one might expect; there
the custom is to treat the maize with limewater before making tortillas,and in this way the nicotinic acid is liberated. Boiling releases niacin insweet corn, a traditional way of treating corn used by the Hopi Indians(Kodicek et al., 1974). In immature seeds, niacin occurs as part of biologically available coenzymes necessary for seed metabolism. Binding of
niacin to carbohydrate by ester linkages may cause it to be retained in
the mature seed until it is utilized; the vitamin availability for man andanimal is thus impaired (Wall and Carpenter, 1988). In rat growth assays for available niacin, corn harvested immaturely (“milk stage”) gave
values from 74 to 88 μg/g, whereas corn harvested at maturity gave assay values of 16 to 18 μg/g (Carpenter et al., 1988).
Beverages including tea and coffee also can contribute significantly
to niacin intake. Roasted coffee contains more niacin than does raw coffee because the trigonelline present in raw coffee is converted to nicotinic acid when the coffee is roasted. In several Central and South American countries, absence of pellagra among those whose staple is maize isattributed to widespread consumption of coffee.

DEFICIENCY
Effects of Deficiency
Niacin deficiency is characterized by severe metabolic disorders inthe skin and digestive organs. The first signs to appear are loss of appetite, retarded growth, weakness, digestive disorders, and diarrhea. The

362

NIACIN

deficiency is found in both human and animal populations that are
overly dependent on foods (particularly corn) low in available niacin
and its precursor, tryptophan.

Ruminants
Young ruminants prior to the development of the rumen would be
expected to suffer B-vitamin deficiencies, including niacin, when diets
contain insufficient quantities of these vitamins. However, early studies
with lambs and calves failed to produce deficiency when niacin-free diets were fed. Ability to produce niacin deficiency is dependent on use ofa low-tryptophan milk diet. From these studies, it may be concluded thata dietary requirement of niacin for young ruminants does not exist as
long as the level of tryptophan is maintained near 0.2% of the diet. In
calves, a diet free of niacin and low in tryptophan produced deficiency
signs of sudden anorexia, severe diarrhea, inability to stand, and dehydration, followed by sudden death. Supplementation with 2.6 mg of
nicotinic acid per liter of milk offered ad libitum twice daily prevented
the deficiency (Hopper and Johnson, 1955).
Synthesis of niacin in the rumen of cattle has been demonstrated andmay be under metabolic control; that is, more niacin is synthesized whensmall amounts are provided in the diet and vice versa (Porter, 1961).
Adult ruminants synthesize adequate niacin by microorganisms and/or
conversion from tryptophan. However, the synthesizing ability appears
to be low, since production responses from niacin supplementation canbe demonstrated in both sheep and cattle. Niacin supplementation is especially beneficial to stressed animals, such as beef cattle being adapted
to high-grain diets or lactating cows that have just calved.

BEEF CATTLE
The addition of niacin to beef cattle diets has been shown to significantly improve average daily gain and feed efficiency. Feeder calves responded positively in rate and efficiency of gain in the 29-day adaptation study by gaining an additional 8.3 kg with 70 ppm added niacin(Byers, 1979). Byers (1979, 1981) summarized 14 beef cattle studies
demonstrating improved gains and feed efficiency by 9.7 and 10.9%, respectively. Growth from all trials was especially beneficial during theadaptation of cattle to feedlot diets (i.e., during the first 40 days).
Niacin appears to be effective in enhancing acclimation and adaptation to urea-supplemented diets. However, niacin stimulation of microbial protein production occurs regardless of dietary nitrogen source, but

363

CHAPTER EIGHTis greatest when natural protein rather than nonprotein nitrogen (urea)
is fed (Brent and Bartley, 1984). This may be partly due to the synthesisof niacin from tryptophan. Minimal amounts of niacin supplied via microbial synthesis and diet become adequate in the later stages of thefeedlot period, whereas earlier they are insufficient to meet demands.
The large response exhibited during the feedlot adaptation phase indicates that supplemental niacin assists in overcoming shipping stress andallows for a more rapid adjustment to feedlot conditions and changing
diets. Summaries of beef cattle studies over the total feeding period (73to 176 days) indicate that 50 or 100 ppm of niacin is more effective thanhigher levels of 150, 250, or 500 ppm, with respect to gain.

DAIRY COWS
Niacin has been assumed to be synthesized in adequate amounts in
the rumen to meet the needs of the dairy cow; however, evidence suggests that this may not be true for the high-producing dairy cow in earlylactation. The effect of niacin on milk production and composition has
been found to be variable, but it appears that niacin (5 to 6 g per cow
per day) may have a beneficial effect on milk production early in the lactation period and when given to ketotic cows.
One report indicated that about 50% of dairy cows in high-production herds go through borderline ketosis during early lactation(Emery et al., 1964). The concentration of serum β-hydroxybutyrate
was reduced from 1.24 to 0.74 mmol/L after 5 days, when 10 g of niacin
was given to cows with ketosis (Flachowsky et al., 1988). Fronk and
Schultz (1979) indicated that treating ketotic dairy cows with 12 g of
nicotinic acid daily had a beneficial effect on the reversal of both subclinical and clinical ketosis. Other studies indicate that 6 g of niacin may
be sufficient (Hoffmann-La Roche, 1994).
Other workers have reported that supplemental niacin increased microbial protein synthesis. Enhanced production of microbial protein
might explain increased milk production, weight gain, and feed efficiency observed when urea-containing diets were supplemented with
250 to 500 mg/kg niacin (Cunha, 1982). Daily niacin supplementationof 3 to 6 g in early lactation dairy cows resulted in slight increases inmilk production. In a study involving six dairy herds, Jaster et al. (1983)
found that milk production of niacin-supplemented cows peaked earlier,and milk production of high-producing cows in first lactation was
greater when they received supplemental niacin. In these herds there wasonly a slight increase in milk fat percentage. Klippel et al. (1993) reported that supplemental niacin decreased milk short- and medium364

NIACIN

chain fatty acids and increased monounsaturated fatty acids.
In one study, increased dry matter intake of 0.8 kg per cow was reported with added niacin in diets based on soybean meal. An increase in
1 kg of dry matter can support 2.5 kg of higher milk yield or minimize
weight loss, improving energy status (Hutjens, 1990). Part of this greater
dry matter intake response could be related to less subclinical ketosis.
Horner et al. (1986) reported that feeding whole cottonseed andmost other dietary fat sources to dairy cows resulted in a reduction ofmilk protein percentage and protein yield. Diets supplemented with
0.03% niacin (6 g niacin/20.45 kg dry matter) increased milk protein
percentage compared to diets with 15% whole cottonseed. The authors
concluded that milk protein depression with whole cottonseed was alleviated by niacin because of stimulation of mammary casein synthesis.
Another study showed no beneficial effect of niacin on milk casein synthesis in cows fed whole cottonseed, which probably was due to theirlate stage of lactation (Lanham et al., 1992). Feeding cows niacin has
been shown to correct fat-induced milk protein depression (Driver et al.,
1990; Cervantes et al., 1996).

LAMBS
Mizwicki et al. (1975) observed that 500 ppm of supplemental
niacin improved feed efficiency of lambs fed a high-concentrate diet containing urea. Supplementation studies (100 ppm niacin) with growing
and finishing lambs revealed increased performance, with evidence thatniacin stimulates rumen microbial protein synthesis (Shields and Perry,1982). These studies indicated that supplemental niacin (1) increased nitrogen utilization, (2) improved the percentage of absorbed nitrogen retained, (3) reduced urinary nitrogen excretion, and (4) reduced the percentage of nitrogen found as urea nitrogen. All these positive changes
point toward improved protein metabolism. Dittrich et al. (1993) indicated that supplemental niacin for lambs increased digestibility of crudefiber and improved wool production.

Swine
One would expect niacin to be deficient in typical swine diets, particularly when corn is fed, because corn is low in available niacin andtryptophan. Wide variation has been observed in the severity of clinicalsigns of niacin deficiency in pigs with similar breeding and environmental backgrounds. Occasionally, animals appear to thrive with no niacin,
and other animals appear to vary in their requirement (Cunha, 1977).
Young pigs require a dietary source of niacin when fed diets deficient in365

CHAPTER EIGHT

tryptophan (Luecke et al., 1948). However, niacin deficiency is lesslikely in pigs that weigh more than 27 kg (Luecke et al., 1948), and,more recently, Ivers and Veum (1993) found no benefit from niacin supplementation in 5-week-old pigs (7.5 kg) fed low-protein corn-soybean
meal diets. During reproduction and lactation, it was not possible toproduce niacin deficiency in sows fed a purified diet with either 18 or
26.1% casein (Ensminger et al., 1951). Ivers et al. (1993) reported nobenefit from supplemental niacin in sow and litter performance whensows were fed corn-soybean meal-oat diets containing 12.8% crude protein. Evidently, the diets used in these studies contained enough tryptophan and/or synthesis of niacin by bacteria in the intestinal tract to supply niacin needs, or the experiment was not long enough for niacin
deficiency to develop.
As mentioned earlier, niacin deficiency is characterized by severe
metabolic disorders in the skin and digestive organs. Signs include poor
appetite, decreased rate of gain (Fig. 8.3), stomatitis, normocytic ane-A

Fig. 8.3 Niacin deficiency. (A) Pig that hasreceived adequate
niacin; (B) pig that has
not received adequate
niacin. The differencein growth and condition is due to the addition of niacin in a dietcontaining 80% groundyellow corn. (Courtesyof D.E. Becker, Illinois
Agriculture Experiment
Station.)366

B

NIACINmia, and achlorhydria, followed by diarrhea, occasional vomiting, andan exfoliate type of dermatitis and hair loss (Cunha, 1977). Degenerative changes in the nervous system are reported to occur in the ganglion
cells in the posterior root, with extensive chromatolysis in the dorsal
root (Wintrobe et al., 1945).
Niacin-deficient pigs have inflammatory lesions of the gastrointestinal tract. Ulcerative necrotic lesions of the large intestine swarm withfusiform bacteria and spirochetes (Fig. 8.4). Diarrhea with foul-smelling
feces particularly involves the large intestine, which thickens, is very red,and appears weak and “rotten.” Enteric conditions may be due to niacin
deficiency, bacterial infection, or both. Niacin-deficient pigs respondreadily to niacin therapy, but there is no benefit for infectious enteritis.
However, adequate dietary niacin probably allows the pig to maintainits resistance to bacterial invasion.

Poultry
There is good evidence that poultry—even chick and turkey embryos—are able to synthesize niacin, but that rate of synthesis may be
too slow for optimal growth. Broilers from 3 to 7 weeks of age are reported not to require supplemental niacin while fed a corn-soybean meal
diet, but the vitamin is required from 1 to 21 days of age (Ruiz andHarms, 1990a,b). It has been claimed that before a marked deficiency ofniacin can occur in the chicken, there must first be a deficiency of tryptophan. Chicks at hatch have considerable tryptophan contained in theprotein of the yolk; thus niacin deficiency will not readily occur unlessthe feed is low for both the amino acid and the vitamin (NRC, 1994).
Experiments using diets containing a limited amount of tryptophan
have shown that the chick requires niacin and that deficiency causes en-

Fig. 8.4 Intestine from
niacin-deficient pig
shows thickened andhemorrhagic mucous
membrane and denuded
areas. (Courtesy ofR.W. Luecke, Michigan

367

CHAPTER EIGHT

largement of the tibiotarsal joint, bowing of the legs, poor feathering,and dermatitis on the feet and head (Scott et al., 1982). The main clinical signs of niacin deficiency in young chicks is enlargement of the hockjoint and bowing of the legs, similar to signs of perosis (Figs. 8.5 and
8.6). The main difference between this condition and the perosis of manganese or choline deficiency is that in niacin deficiency the Achilles tendon rarely slips from its condyles.
Niacin deficiency in the chick is characterized by appetite loss andgrowth failure. The deficiency results in blacktongue, a condition characterized by inflammation of the tongue and mouth cavity. Beginningat about 2 weeks of age, the entire mouth cavity, as well as the esophagus, becomes distinctly inflamed; growth is retarded, and feed consumption is reduced. Weight loss occurs, and both egg production andhatchability are reduced in niacin-deficient laying hens. Shell quality isimproved with niacin supplementation (Leeson et al., 1991). Death loss
can be affected. Jackson (1992) reported that 30 ppm of dietary niacin
significantly decreased the mortality rate of layers compared to 10
ppm.
Turkey poults, pheasant chicks, ducklings, and goslings all expressed perosis as the primary niacin deficiency sign (NRC, 1994). Signs
in turkeys and ducks, while similar to those of chickens, are much more
severe. Compared to the chick, the turkey poult, duckling, pheasantchick, and gosling have higher requirements for niacin. This higher requirement is related to the less efficient conversion of tryptophan toniacin by these species (see Metabolism). Ducks receiving low-niacin diets show severely bowed legs and ultimately become too crippled andweak to walk. Niacin deficiency in the turkey is also characterized by severe bowing of the legs and enlargement of the hock joint. Goslings fed
purified rations develop perosis and hock deformities that are preventable with administration of nicotinic acid (Briggs et al., 1953).

HorsesNo requirements for niacin have been established for the horse, anddeficiency has not been reported (NRC, 1989b). Synthesis of niacin bymicroflora in the lower digestive tract, as well as tissue synthesis fromtryptophan, should likely preclude niacin deficiency. Microbial synthesisin the horse intestine was shown by Carroll et al. (1949), who fed a dietcontaining 3 mg of nicotinic acid per kilogram of dry matter and found
the following nicotinic acid concentrations (ppm dry matter) in ingesta:
duodenum, 55; ileum, 58; cecum, 121; anterior large colon, 96; and anterior small colon, 119.
368

NIACIN

Fig. 8.5 Leg disorders
in niacin-deficientbroiler chicks. The birdon the left, with bowed
legs, was fed a cornsoybean meal diet without supplemental
niacin. (Courtesy of N.
Ruiz and R. Harms,
University of Florida.)

Other Animal Species
DOGS AND CATS
The similarity of niacin deficiency signs in dogs and humans is important because the dog became the laboratory animal used to identify
the vitamin deficiency. Signs of deficiency in the dog are collectively
known as blacktongue (canine pellagra). The condition occurs in dogs
fed a diet primarily of flaked corn (maize) with little material of animal
origin. Clinical signs include anorexia, apathy, retarded growth, weight
loss, and drooling of thick ropy saliva (Fig. 8.7). There is severe cheilosis, glossitis, and gingivitis. Necrotic patches and ulcers may be seen on
the oral mucosa, and there is a foul odor. The inflammatory changes
may extend to the esophagus and eventually to the large intestine. Thereis bloody diarrhea, inflammation, and hemorrhagic necrosis of the duodenum and jejunum, with shortening and clubbing of villi and inflammation and degeneration of the mucosa of the large intestine. Uncorrected deficiencies lead to dehydration, emaciation, and death (NRC,
1985a). Because of burning or itching, animals scratch or bite at theskin, producing a traumatic dermatitis. Death generally occurs within
10 days of the onset of clinical signs.
Virtually no niacin is synthesized from tryptophan in the cat (seeMetabolism). Niacin-deficient cats exhibit weight loss, anorexia, weakness, and apathy (NRC, 1986). There is drooling of thick saliva with afoul odor. The oral cavity is characterized by ulceration of the upper
palate, and the tongue is fiery red, with ulceration and congestion along
the anterior border. The fur may be unkempt, and diarrhea occurs. The
deficiency can be associated with respiratory disease, which contributes
to an early death.

369

CHAPTER EIGHTA

B

Fig. 8.6 Niacin deficiency
in turkey poults. (A and B)
The birds on the left side,which were fed a corn-soybean meal without supplemental niacin, showed perosis-like signs. (C)Comparison of the legs ofthe poults in B. (Courtesyof N. Ruiz and R. Harms,
University of Florida.)

C

FISH
In fish species studied, tryptophan is not an adequate precursor toniacin, and the vitamin deficiency develops rapidly in many fish fed a
niacin-deficient diet. Signs of niacin deficiency in trout and salmon include anorexia, reduced growth, poor feed conversion, photosensitivity
or sunburn, intestinal lesions, muscular weakness and spasms, and increased mortality (NRC, 1993). Channel catfish and common carp deficient in niacin show skin and fin lesions, hemorrhages, deformed jaws,
370

NIACIN

Fig. 8.7 Niacin-deficient dog with blacktongue exhibits drooling of thick, ropy
saliva. (Courtesy of V. Ramadus Murthy, National Institute of Nutrition, Jamasi-Osmania, Hyderabad, India.)anemia, exophthalmia, and high mortality rate (NRC, 1993). Niacin-deficient tilapia grew poorly, and skin, fin, and mouth lesions and hemorrhages developed; the snout was deformed and edema occurred (Shiau
and Suen, 1992). Japanese eels exhibited reduced growth, ataxia, skinlesions, and dark coloration.

FOXES AND MINK
The mink is similar to the cat concerning niacin metabolism, withvery inefficient conversion of tryptophan to niacin. Young mink fed a
deficient diet displayed nonspecific signs, including weight loss, weak
voice, general weakness, and bloody diarrhea (NRC, 1982a). Foxes fed
a niacin-deficient diet exhibited anorexia, weight loss, and typical blacktongue, characterized by severe inflammation of the gums and fiery redness of the lips, tongue, and gums (NRC, 1982a).

LABORATORY ANIMALS
Laboratory rodents often do not show characteristic signs of niacin
deficiency but exhibit nonspecific signs of appetite loss and growth re371

CHAPTER EIGHT

tardation (NRC, 1995). It is difficult to produce niacin deficiency in rats
because compared to other species, rats are very efficient in converting
tryptophan to niacin; 33 to 40 mg of tryptophan yield 1 mg of niacin.
Rats fed niacin- and tryptophan-deficient diets develop behavioral
changes, convulsions, diarrhea, rough hair coat, and alopecia. Niacindeficient guinea pigs exhibit poor growth; reduced appetite; pale feet,nose, and ears; drooling, anemia, and a tendency toward diarrhea.
Niacin deficiency has not been reported in the mouse, gerbil, hamster, orvole (NRC, 1995).

NONHUMAN PRIMATES
Niacin deficiency in the rhesus monkey is characterized by weight
loss, apathy, anemia, skin pigmentation, and bloody diarrhea (NRC,
1978). Inefficient conversion of tryptophan to niacin has been suggested.

RABBITS
Substantial synthesis of niacin occurs in the intestinal tract of rabbits fed niacin-deficient diets. Niacin deficiency in rabbits has resulted inpronounced loss of appetite, followed by emaciation and diarrhea(NRC, 1977).

HumansTraditionally, niacin deficiency in humans has been equated withpellagra. Symptoms of deficiency can be considered under three headings: (1) skin changes; (2) lesions of the mucous membranes of themouth, tongue, stomach, and intestinal tract; and (3) changes of nervous origin.
The earliest symptom of pellagra is inflammation and soreness ofthe mouth, followed by bilateral symmetrical erythema on all parts ofthe body exposed to sunlight. Therefore, common sites are the extensor
surfaces of the extremities, face, and neck (Fig. 8.8). The lesions are also
found at sites of constant irritation, such as under the breast, scrotum,
axilla, and perineum. A characteristic feature of these photosensitive lesions is their clear-cut demarcation from the adjoining, unaffected parts.
The appearance of the dermatosis varies with the severity of the disease
and from patient to patient.
Gastrointestinal symptoms often include glossitis and stomatitis,
and the tongue has a characteristic swollen and beefy-red appearance.
There is also anorexia, abdominal discomfort, and diarrhea. A burning

372

A

B

C

Fig. 8.8 Niacin deficiency: typical dermal lesions associated with pellagra. Dermatitis is severe when sunlight reacts with exposed skin. (A) Dermal lesions
around neck have been referred to as Casal’s necklace. (B) Affected hands andfeet of a pellagrin. (C) Affected hands of a child. (Courtesy of V. Ramadus
Murthy, National Institute of Nutrition, Jamasi-Osmania, Hyderabad, India.)373

CHAPTER EIGHT

or raw sensation of the mouth, stomach, and rectum has been described.
Angular stomatitis, cheilosis, atrophy of the lingual papillae, and glossitis are commonly seen. However, it is not known whether these are manifestations of niacin deficiency or are due to the associated deficiency ofriboflavin and other members of the vitamin B-complex group.
Early mental symptoms include lassitude, lack of ambition, apprehension, depression, morbid fears, and memory loss, and these may besucceeded by disorientation, confusion, hysteria, and sometimes maniacal outbursts. More general symptoms, such as headache, irritability, inability to concentrate, and apathy, may also occur. Insomnia is a very
common complaint. Various psychoses, such as manic-depressive syndromes, severe paranoia, and hallucinations, may necessitate confinement in a psychiatric institution. A considerable percentage of the patients in such institutions have been reported to be pellagrins, whichbears testimony to the serious nature of the mental changes that occur(Gopalan and Rao, 1978).
As with deficiencies of the other B vitamins, pellagra often accompanies poverty, chronic alcoholism, dietary peculiarities, fever, hyperthyroidism, pregnancy, and the stress of injury or surgical procedures. Its
incidence is much greater during the spring than any other season. In theadvanced stages, pellagra can be diagnosed by the classic “three D’s”—
dermatitis, diarrhea, and dementia (death being the “fourth D”). Recognition of the disease in an early stage must depend on reliable medical
and dietary history and thorough physical examination.
Pellagra has been endemic in certain areas of Italy, Spain, Romania,
Egypt, and the southeastern United States. However, it has almost disappeared from industrialized countries except for its occurrence in somealcoholics. Hereditary disorders of schizophrenia and Hartnup disease
involve impaired niacin function and are treated with high doses of thevitamin. Prolonged treatment with isoniazid also may lead to niacin deficiency by competition of the drug with pyridoxal phosphate, a coenzyme required in the tryptophan-to-niacin pathway. As mentioned earlier, pellagra has been considered a disease of the corn-eating populationand has practically disappeared from all affluent societies. It is still encountered in parts of China, among the Bantus of South Africa, in the
Middle East, in Yugoslavia, and in the corn-eating population of India(Gopalan and Rao, 1978).
Poverty and resulting diet selection can greatly influence incidenceof pellagra. The social and economic upheaval of the U.S. Civil War(1860–1865) led to a corn-based diet for large sections of the population, and it was not until the United States entered World War II that in374

NIACIN

creasing employment and a rise in the general standard of living solved
the dietary problem. When cotton prices were high, more money wasavailable to buy foods that prevent pellagra (e.g., animal products) versus subsisting on low-cost corn products. Pellagra was unknown insouthern Africa until the outbreak of rinderpest in 1897, which led towidespread death of cattle and a major change in the dietary habits of
the Bantu. From being a meat-eating and milk-drinking community they
became, and have remained, largely maize eaters, and pellagra continues
to be a major problem of public health nutrition in South Africa.
It had been accepted that the low tryptophan and unavailable niacin
in corn was responsible for pellagra (see Natural Sources). However,identification of pellagra in populations subsisting on sorghum, a millet
in which niacin is not so low or biologically unavailable, raised serious
doubts regarding this concept (Gopalan, 1969). Pellagra is endemic
among the poor communities of India where the millet Sorghum vulgare
is the staple food. The effect of a high intake of leucine, and possibly anisoleucine-leucine imbalance, was suggested as causing pellagra insorghum-eating as well as corn-eating populations (Gopalan, 1969).
Later studies, however, showed conclusively that excess leucine does notcause pellagra, nor does it impact the metabolism of tryptophan toniacin (Manson and Carpenter, 1978; Lowry and Baker, 1989). One
study indicated that concomitant vitamin B6 deficiency, either primaryor brought about by leucine toxicity, may also have an important role
(Rao et al., 1975). Because iron is required by two enzymes in the tryptophan-to-niacin pathway, pellagra in populations having endemic anemia may be due not only to inadequate intake of bioavailable niacin but
also to inadequate intakes of bioavailable iron (Oduho et al., 1994).

Assessment of Status
A functional biochemical test for assessing body reserves has had
only limited success. Determination of blood serum levels of niacin or
niacin-dependent enzymes has not proved to be a reliable or acceptable
method for evaluating niacin status. However, although NAD in liver
was not affected in niacin-deficient quail, the level of pectoral muscle
NAD was markedly reduced (Park et al., 1991). In humans, erythrocyte
NAD may serve as a sensitive indicator for the assessment of niacin status (Fu et al., 1989). Biochemical assessment of niacin nutritional statusis usually based on measurement of urinary metabolites. Measurementof niacin metabolites would be dependent on species, as marked differences in type of metabolites exist among species (see Metabolism). In humans and most monogastric species, niacin is excreted largely as methy375

CHAPTER EIGHT

lated products. Excreted niacin metabolites in a healthy, well-nourishedadult human include 4 to 6 mg of N´-methylnicotinamide. One of thetesting procedures widely used in human population surveys is the estimation of urinary excretion of N´-methylnicotinamide during a periodof 4 to 5 hours after administering a test dose of 50 mg of niacinamide(Narasinga Rao and Gopalan, 1984).

SUPPLEMENTATION
Niacin supplementation must be considered as a possibility for allclasses of livestock and in particular in swine and poultry diets. Much of
the niacin in common feeds (plant sources) is in a bound form that is notavailable to animals (see Natural Sources). Although the bioavailabilityof niacin is suggested to be 100% in soybean meal, it is likely zero inwheat and sorghum and varies from 0 to 30% in corn. In formulating
swine and poultry diets, therefore, niacin values for corn and other cereal grains and their by-product feeds should be almost disregarded. It
is best to assume that these feeds provide no available niacin for the pig
or chick (Cunha, 1982).
The pig and chick and most species can use the essential amino acidtryptophan to synthesize niacin, but they cannot convert niacin back totryptophan. Therefore, if a diet contains enough niacin, the tryptophan
is not depleted for niacin synthesis. Most monogastric diets do not contain large excesses of tryptophan, particularly diets based on corn. Tryptophan concentrations not only are low in corn, but are largely unavailable. Therefore, one should make sure that monogastric diets areadequate in niacin since it is very inexpensive, and it would be poor economics to satisfy niacin needs with the more expensive tryptophan(Cunha, 1982). Also, some species, such as the cat, mink, and fish, apparently lack the ability to convert tryptophan to niacin.
The most critical time for supplementation is during early growth,when requirements are the highest (Table 8.3). Evidence is lacking regarding the need to supplement practical diets of mature poultry (Scottet al., 1982) and swine (Ivers et al., 1993) with niacin. However, niacin
requirements as recommended by the National Research Council foryoung swine and poultry may be insufficient. Cunha for swine and Scottfor poultry have recommended higher niacin requirements as a reasonable safety factor (Cunha, 1982). Higher niacin levels are mainly recommended when subclinical disease level, stress, and higher production
rates are encountered. For poultry, recommended levels are generally 12
to 25% higher than the NRC recommendations. Increased swine re376

NIACINquirements are approximately double those of the NRC for growing
swine under stressful conditions, with breeding animals needing a fourfold increase (Cunha, 1982). Producers must judge their own livestock
operations in relation to stress conditions for animals and other factorsthat may require higher niacin supplementation levels.
Addition of niacin to a milk replacer is recommended at a level of
2.6 ppm because of lack of synthesis in the undeveloped rumen. Inclusion in diets of growing heifers cannot be justified by evidence (Schultz,
1983).
Supplementation of niacin for ruminants can be beneficial. If tryptophan is present in large amounts, the microorganisms can probably
synthesize considerable niacin. However, in urea diets the microbes do
not have enough tryptophan to draw upon and hence use exogenous
sources of niacin. Supplementation appears to be of greatest benefit tostressed animals, such as beef cattle entering the feedlot, dairy cows immediately postpartum, and cows suffering from subclinical ketosis.
Stressed sheep with ruminal acidosis had decreased plasma nicotinicacid from 7.01 to 0.42 μg/ml within 24 hours (Basoglu et al., 1993).
Recommended supplemental niacin levels are estimated to be 50 to 100
ppm niacin for finishing beef cattle and 3 to 12 g daily for milking cows.
Niacin appears to be effective in enhancing acclimation and adaptation to urea-supplemented diets. Minimal amounts of niacin supplied
via microbial synthesis and diet become adequate in the later stages ofthe feedlot period, whereas earlier they were insufficient to meet demands. The large response during the feedlot adaptation phase indicatesthat supplemental niacin assists in overcoming shipping stress and allows for a more rapid adjustment to feedlot conditions and changing diets. Summaries of beef cattle studies over the total feeding period (73 to
176 days) indicate that 50 to 100 ppm of niacin is more effective thanhigher levels of 150, 250, or 500 ppm with respect to gain.
Brent and Bartley (1984) found that a higher milk production response from niacin was greater for early postpartum cows than for thosein midlactation. Niacin response in early postpartum cows might be partially explained via reduction in ketosis. Di Costanzo et al. (1997) reported that niacin did not significantly increase milk production but decreased skin temperatures during periods of mild or severe heat stress.
Although general results of field experiments on niacin supplementation for lactating cows are positive for both milk production and fat
test, the somewhat erratic results and the occasional lack of statisticalsignificance make it difficult to give an unqualified recommendation to
include niacin in diets of all dairy cows in early lactation. However,377

CHAPTER EIGHT

enough positive evidence is available to recommend it for herds withabove-average incidence of ketosis and fat cow problems. For individual
cows with positive tests indicating subclinical ketosis, a level of 12 g per
day is recommended. When used as a general diet additive, a level of 6g per day is suggested, starting 2 weeks before calving. This recommendation to start before calving is based on the evidence that it inhibits
lipid mobilization and thus may prevent abnormal deposition of fat in
the liver at calving time. Most evidence suggests that feeding be continued 8 to 12 weeks after calving (Schultz, 1983).
Niacin supplementation is important for human populations that
depend primarily on corn and sorghum in their diets. Likewise, pellagra
was found in war-torn nations of the Sahel and southeastern parts ofAfrica. Although the condition is largely eliminated in the more technologically advanced countries, it still is found as an undiagnosed complication of malnutrition in today’s urban society (Bosco, 1980). Niacin deficiency is usually found in people with chronically bad dietary habits;
people who consume low-calorie, low-protein, high-fat, and high-carbohydrate diets; alcoholics; and people with diseases that interfere withdigestion and absorption. A large amount of wine (750 ml) markedly
impairs liver protein metabolism, which is counteracted with supplemental nicotinamide (Volpi et al., 1997). Also, some individuals withmetabolic defects (e.g., Hartnup disease, deviation of niacin-tryptophan
metabolism) require supplemental niacin at levels higher than normallyconsumed in typical foods. Cases of pellagra were reported on isoniazid
therapy, which responded to combined vitamin B6 and niacin treatment(DiLorenzo, 1967).
Pharmacological effects from massive supplemental doses of niacin
have been reported in humans. Large doses of nicotinic acid (but not theamide) produce vascular dilatation or “flushing,” with accompanying
sensation of burning of the face and hands. Large doses of nicotinic acid
(3 g or more per day) administered orally result in a significant reduction of serum cholesterol and β-lipoprotein cholesterol (Narasinga Rao
and Gopalan, 1984; Basu and Mann, 1997). The administration of nicotinic acid in the Coronary Drug Project was associated with a reduction
in recurrent myocardial infarctions and in long-term total mortality(Canner et al., 1986). Nicotinic acid given as a drug in doses of 1.5 to 3
g/day decreased concentrations of total and low-density lipoprotein cholesterol and increased concentrations of high-density lipoprotein cholesterol. Niacin has also been used in treatment of schizophrenia. However,according to a review of all the relevant individual controlled studies,
there is no evidence that nicotinic acid is an antipsychotic agent (Petrie
378

NIACIN

and Ban, 1975). Nicotinamide has been shown to have a positive role in
the reduction of insulin-dependent diabetes mellitus (Behme, 1995).
Studies with rats revealed that pharmacological levels of niacin interfered with methionine metabolism, which was counteracted with simultaneous administration of niacin and vitamin B6 (Basu and Mann,
1997). Beneficial effects of pharmacological doses of niacin must be
weighed against the long-term effects of high doses, including laboratory
evidence of diabetes, hepatic injury, and activation of peptic ulcer.
Niacin is supplied in two forms—niacinamide and nicotinic acid—both of which provide available niacin. In studies with chicks, relative
to nicotinic acid used as a standard (100%), nicotinamide activity was
124%. Nicotinamide in NAD was utilized with an efficiency of 95% relative to nicotinamide per se (Oduho and Baker, 1993). Crystalline products are used in feeds and pharmaceuticals, and dry dilutions in feeds.
An additional source of supplemental niacin is from the vitamin K supplement menadione nicotinamide bisulfite (MNB). Results with chicks
suggest that MNB is fully effective as a source of vitamin K and niacin
activity (Oduho et al., 1993).
Commercially produced niacin is quite stable compared to mostother vitamins. Synthetic niacin and the amide were found to be stablein premixes with or without minerals for 3 months (Verbeeck, 1975).
Gadient (1986) reported niacin to be insensitive to heat, oxygen, moisture, and light. The retention of niacin activity in pelleted feeds after 3
months at room temperature should be 95 to 100% as a general rule.
One report demonstrated 98% retention of niacin after 6 months in avitamin premix; however, retention was only 58% when the premix contained choline and trace minerals (Coelho, 1991).

TOXICITY
Harmful effects of niacin occur at levels far in excess of requirements. Nonruminant animals can tolerate oral exposures of at least 10
to 20 times their normal requirements. Limited research indicates that
nicotinic acid and nicotinamide are toxic at dietary intakes greater thanabout 350 mg/kg of body weight per day (NRC, 1987). Short-term intravenous administration of niacin at a dose of 2.5 g/kg was needed before 50% of test mice died. Respective values for subcutaneous and oralroutes of administration were 2.8 and 4.5 g/kg in mice. Nicotinamide is
two to three times more toxic than the free acid (Waterman, 1978).
High levels of nicotinic acid, such as 3 g/day in humans, can cause
vasodilation, itching, heat sensation, nausea, vomiting, headache, and379

CHAPTER EIGHT

occasional skin lesions. The substance can also cause more serious problems, including abnormally high blood sugar levels, elevated uric acidlevels (which people with gout must avoid), peptic ulcers, and liver damage. Niacin therapy was discontinued in eight children because of flushing, abdominal pain, vomiting, headache, or elevated serum aminotransferase levels. It is suggested that although niacin treatment forchildren is efficacious, adverse effects are common (Colletti et al., 1993).
In dogs, oral administration of 2 g of nicotinic acid per day (133 to 145
mg/kg of body weight) produced bloody feces in a few dogs, followedby convulsions and death (NRC, 1987).

■ REFERENCES
Anonymous (1987). Nutr. Rev. 45(5), 142.
Baker, D. H., Allen, N. K., and Kleiss, A. J. (1973). J. Anim. Sci. 36, 299.
Basoglu, A., Turgut, K., Eksen, M., Tras, B., Maden, M., Ok, M., Bas, A. L., andKececi, T. (1993) Vet. Fakultesi. Dergisi 2, 31.
Basu, T. K., and Mann, S. (1997). J. Nutr. 127, 117.
Bechgaard, H., and Jespersen, S. (1977). J. Pharm. Sci. 66, 871.
Behme, M. T. (1995). Nutr. Rev. 53, 137.
Bosco, D. (1980). The People’s Guide to Vitamins and Minerals from A to Zinc.
Beaverbooks, Ltd., Ontario, Canada.
Brent, B. E., and Bartley, E. E. (1984). J. Anim. Sci. 59, 813.
Briggs, G. M., Hill, E. G., and Canfield, T. H. (1953). Poult. Sci. 32, 678.
Byers, F. M. (1979). Anim. Nutr. Health, 35, 20.
Byers, F. M. (1981). Anim. Nutr. Health 36, 36.
Canner, P. L., Berge, K. G., and Wenger, N. K. (1986). J. Am. Cell. Cardiol. 8,1245.
Carpenter, K. J., Schelstraete, M., Vilicich, V. C., and Wall, J. S. (1988). J. Nutr.
118, 165.
Carroll, F. D., Gross, H., and Howell, C. E. (1949). J. Anim. Sci. 8, 290.
Carson, D. A., Seto, S., and Wasson, D. B. (1987). J. Immunol. 138, 1904.
Carter, E. G. A., and Carpenter, K. J. (1982). J. Nutr. 112, 2091.
Cervantes, A., Smith, T. R., and Young, J. W. (1996). J. Dairy Sci. 79, 105.
Chen, B. J., and Austic, R. E. (1989). Poult. Sci. 68(Suppl. 1), 27(Abstr.).
Coelho, M. B. (1991). Vitamin Stability in Premixes and Feeds: A Practical Approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Colletti, R. B., Neufeld, E. J., Roff, N. K., McAuliffe, T. L., Baker, A. L., andNewburger, J. W. (1993). Pediatrics 92, 78.
Cunha, T. J. (1977). Swine Feeding and Nutrition. Academic Press, Inc., NewYork.
Cunha, T. J. (1982). Niacin in Animal Feeding and Nutrition. National Feed Ingredients Association (NFIA), Fairlawn, New Jersey.
Darby, W. J., McNutt, K. W., and Todhunter, E. N. (1975). Nutr. Rev. 33, 289.
Di Costanzo, A., Spain, J. N., and Spiers, D. E. (1997). J. Dairy Sci. 80, 1200.
380

NIACIN

DiLorenzo, P. A. (1967). Acta Dermatol. Venereol. 47, 318.
DiLorenzo, R. N. (1972). Studies on the Genetic Variation in Tryptophan-Nicotinic Acid Conversion in Chicks. Ph.D. Dissertation, Cornell Univ., Ithaca,
New York.
Dittrich, A., Suss, R., and Geissler, C. (1993). In Symposium, Vitamine undweitere Zusatzstoffe bei Mensch und Tier: 4, G. Flachowsky and R. Schubert (eds.), p. 176. Jena, Germany.
Driver, L. S., Grummer, R. R., and Schultz, L. H. (1990). J. Dairy Sci. 73, 463.
Emery, R. S., Burg, N., Braur, L. D., and Blank, G. N. (1964). J. Dairy Sci. 47,
1074.
Ensminger, M. E., Colby, R. W., and Cunha, T. J. (1951). Wash. Agric. Exp. Stn.
Circ. 134, Pullman, Washington.
Erickson, P. S., Murphy, M. R., McSweeney, C. S., and Trusk, A. M. (1991). J.
Dairy Sci. 74, 3492.
Flachowsky, G., Lober, U., Ast, H., Wolfram, D., and Matlhey, M. (1988). Wiss.
Z. Karl-Marx-Univ. Leipzig Math.-Naturwiss R. 37, 55.
Fronk, T. J., and Schultz, L. H. (1979). J. Dairy Sci. 62, 1804.
Fu, C. S., Swendseid, M. E., Jacob, R. A., and McKee, R. W. (1989). J. Nutr.
119, 1949.
Gadient, M. (1986). In Proc. Maryland Nutrition Conference for Feed Manufacturers, p. 73. College Park, Maryland.
Ghosh, H. P., Sarkar, P. K., and Guha, B. C. (1963). J. Nutr. 79, 451.
Gopalan, C. (1969). Lancet 1 (7587), 197.
Gopalan, C., and Rao, K. S. J. (1978). In Handbook Series in Nutrition andFood, Section E: Nutritional Disorders Volume 3 (M. Rechcigl Jr., ed.), p.
23. CRC Press, Inc.,West Palm Beach, Florida.
Halvorsen, K., and Halvorsen, S. (1963). Pediatrics 31, 29.
Hankes, L. V. (1984). In Handbook of Vitamins (L. J. Machlin, ed.), p. 329.
Marcel Dekker, New York.
Haralambie, G. (1976). Nutr. Metab. 20, 1.
Harris, H. F. (1919). Pellagra. MacMillan, New York.
Henderson, L. M., and Gross, C. J. (1979). J. Nutr. 109, 654.
Hoffmann-La Roche (1994). Vitamin Nutrition for Ruminants, RCD 8361/191.
Hoffmann-La Roche Inc., Nutley, New Jersey.
Hopper, J. H., and Johnson, B. C. (1955). J. Nutr. 56, 303.
Horner, J. L., Coppock, C. E., Schelling, G. T., Labore, J. M., and Nave, D. H.
(1986). J. Dairy Sci. 69, 3087.
Hutjens, M. F. (1990). In Proc. 1990 Natl. Feed Ingred. Assoc. Nutr. Inst.: Developments in Vitamin Nutrition and Health Applications. National Feed
Ingredients Association (NIFA), Des Moines, Iowa.
Ikeda, M., Tsuji, H., Nakamura, S., Ichiyama, A., Nishizuka, Y., and Hayaishi,
O. (1965). J. Biol. Chem. 240, 1395.
Ivers, D. J., Rodhouse, S. L., Ellerseick, M. R., and Veum, T. L. (1993). J. Anim.
Sci. 71, 651.
Ivers, D. J., and Veum, T. L. (1993). J. Anim. Sci. 71, 3383.
Jackson, M. (1992). Feeding Layers: Nutritional Considerations. Multi-State
Poultry Meeting, Indianapolis, Indiana.
Jaster, E. H., Hartwell, G. F., and Hutjens, M. F. (1983). J. Dairy Sci. 66, 1046.
381

CHAPTER EIGHT

Klippel, M., Mockel, P., and Flachowsky, G. (1993). In Symposium Vitamine
und weitere Zusatzstoffe bei Mensch und Tier: 4, G. Flachowsky and R.
Schubert (eds.), p. 194. Jena, Germany.
Kodicek, E., Ashby, D. R., Muller, M., and Carpenter, K. J. (1974). Proc. Nutr.
Soc. 33, 105A.
Lanham, J. K., Coppock, C. E., Brooks, K. N., Wilks, D. L., and Horner, J. L.
(1992). J. Dairy Sci. 75, 184.
Leeson, S., Caston, L. J., and Summers, J. D. (1991). Poult. Sci. 70, 1231.
Loosli, J. K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.), Academic Press, San Diego.
Lowry, K. R., and Baker, D. H. (1989). FASEB J. 3, A666(Abstr.).
Luce, W. G., Peo, E. R., and Hudman, D. B. (1967). J. Anim. Sci. 26, 76.
Luecke, R. W., McMillen, W. N., Thorp Jr., F., and Tull, C. (1948). J. Nutr. 36,
417.
Manson, J. A., and Carpenter, K. J. (1978). J. Nutr. 108, 1883.
Mason, J. B., Gibson, N., and Kodicek, E. (1973). Br. J. Nutr. 30, 297.
McCollum, E. V. (1957) In A History of Nutrition, p. 302. Riverside Press,
Cambridge, Massachusetts.
Mertz, W. (1975). Nutr. Rev. 33, 129.
Mizwicki, K.L., Owens, F. N., Issacson, H. R., and Shockey, B. (1975). J. Anim.
Sci. 41, 411(Abstr.).
Narasinga Rao, B. S., and Gopalan, C. (1984). In Nutrition Reviews, Present
Knowledge in Nutrition (R. E. Olson, ed.), p. 318. The Nutrition Foundation, Inc., Washington, D.C.
Nisselbaum, J. S., and Green, S. (1969). Anal. Biochem. 27, 212.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Oduho, G. W., and Baker, D. H. (1993). J. Nutr. 123, 2201.
Oduho, G. W., Chung, T. K., and Baker, D. H. (1993). J. Nutr. 123, 737.
382

NIACIN

Oduho, G. W., Han, Y., and Baker, D. H. (1994). J. Nutr. 124, 444.
Olentine, C. (1984). Feed Manage. 35(4), 18.
Park, I. K., Shin, S., and Marquardt, R. R. (1991). Int. J. Biochem. 23(10),1005.
Petrie, W. M., and Ban, T. A. (1975). Drugs 30, 58.
Porter, J. W. G. (1961). In Digestive Physiology and Nutrition of the Ruminant(D. Lewis, ed.), p. 226. Butterworth, London.
Rao, S. B., Raghuram, T. C., and Krishnaswamy, K. (1975). Nutr. Metab. 18,
318.
Rawling, J. M., Jackson, M., Driscoll, E. R., and Kirland, J. B. (1994). J. Nutr.
124, 1547.
RDA. (1989). Recommended Dietary Allowances, 10th Ed., National Academyof Sciences-National Research Council, Washington, D.C.
Rose, D. P., and Braidman, I. P. (1971). Am. J. Clin. Nutr. 24, 673.
Ruiz, N., and Harms, R. H. (1990a). Poult. Sci. 69, 433.
Ruiz, N., and Harms, R. H. (1990b). Poult. Sci. 69, 2231.
Ruiz, N., and Harms, R. H. (1990c). Poult. Sci. 69, 446.
Schultz, L. H. (1983). In Proceedings Eighteenth Annual Pacific Northwest Animal Nutrition Conference p. 69. Oregon State Univ., Corvallis, Oregon.
Scott, N. L., Nesheim, M. C., and Young, R. J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Shiau, S. Y., and Suen, G. S. (1992). J. Nutr. 122, 2030.
Shields, D. R., and Perry, T. W. (1982). Effect of Supplemental Niacin on Protein Utilization by Lambs National Feed Ingredients Association (NFIA),
Fairlawn, New Jersey.
Stein, J., Daniel, H., Whang, E., Wenzel, U., Han, A., and Rehner, G. (1994). J.
Nutr. 124, 61.
Stocchi, V., Cucchiarini, L., Canestrari, F., Piacentini, M., and Fornaini, G.
(1987). Anal. Biochem. 167, 181.
Vague, P., Vialettes, B., Lassman-Vague, V., and Vallo, J. (1987). Lancet 1(8533), 619.
Verbeeck, J. (1975). Feedstuffs 47(36), 4.
Volpi, E., Lucidi, P., Cruciani, G., Monarchia, F., Reboldi, G., Brunetti, P., Bolli,
G.B., and De Feo, P. (1997). J. Nutr. 127, 2199.
Wall, J. S., and Carpenter, K. J. (1988). Food Technol. 42(10), 198.
Waterman, R. A. (1978). In Handbook Series in Nutrition and Food, Section E:
Nutritional Disorders Volume 1, (M. Rechcigl Jr., ed.), p. 29, CRC Press,
Inc., West Palm Beach, Florida.
Wertz, A. W., Lojkin, M. E., Bouchard, B. S., and Derby, M. B. (1958). J. Nutr.
64, 339.
Wintrobe, M. M., Stein, H. J., Follis, R. H., and Humphreys, S. (1945). J. Nutr.
30, 395.
Zhang, J. Z., Henning, S. M., and Swendseid, M. E. (1993). J. Nutr. 123, 1349.

383

chapter nine

V I T A M I N B6

INTRODUCTION
Vitamin B6 refers to a group of three compounds: pyridoxol (pyridoxine), pyridoxal, and pyridoxamine. Their activities are equivalent inanimals but not in various microorganisms. Vitamin B6 acts as a component of many enzymes that are involved in the metabolism of proteins, fats, and carbohydrates. The vitamin is particularly involved invarious aspects of protein metabolism.
Common feed ingredients for typical poultry and swine diets contain adequate amounts of vitamin B6. Although such conditions are rare,vitamin B6 can be deficient for poultry and swine in a few situations. Although animal diets are generally adequate in vitamin B6, there is evidence of low vitamin B6 status in the human population, especially inyoung and pregnant or lactating women. Vitamin B6 deficiency in humans may be induced by long-term drug therapy, increased requirements
resulting from malignancies, and numerous disease conditions.

HISTORY
The early history of vitamin B6 has been summarized by McCollum(1957), Snell (1986), and Loosli (1991). Clinical signs of what was later
to be known as vitamin B6 deficiency were described in 1926 by Goldberger and Lillie in their attempts to produce pellagra in experimental
animals. Rats developed a severe dermatitis called acrodynia (painful extremities). This dermatitis was also referred to as rat pellagra and wascharacterized by skin lesions, especially on the limbs and trunk, and by
swelling of the paws and ears. These signs were easily distinguished from385

9

CHAPTER NINE

those of riboflavin. In 1934 György first recognized vitamin B6 as a distinct vitamin and showed that it prevented rat acrodynia and that thecondition was not prevented by vitamin B1 (thiamin), vitamin B2 (riboflavin), or vitamin B3 (niacin). Two years later Lepkowsky and associates showed the vitamin to be an essential nutrient for chickens.
Isolation of the vitamin in crystalline form was accomplished independently in five different laboratories in 1938. The vitamin was firstisolated from rice bran and yeast. The structure of the vitamin was first
explained by Kuhn and coworkers, who named it “adermin,” because it
was believed that skin dermatitis was the only clinical sign of the deficiency. The term was discarded when evidence accumulated for nondermal clinical signs for the vitamin deficiency. In view of the chemical
structure of the compound (pyridine structure), György proposed pyridoxine as the vitamin name, and this proposal was widely adopted.
However, by 1945 subsequent bacterial studies identified two additional
compounds—pyridoxal and pyridoxamine. Thus, by official action of
the Society of Biological Chemists and the American Institute of Nutrition, the original term B6 became the approved name for this vitamin.
In 1939 chemical synthesis was accomplished by two independent
laboratories. Therefore, only 5 years elapsed between the discovery of
the vitamin and its chemical synthesis. The speed of isolation indicatesthe intense efforts expended in the mid-1930s to identify new vitamins.
The principal coenzyme form of vitamin B6 pyridoxal 5´phosphate (originally called codecarboxylase) was discovered in bacteria by Umbreit in1944.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Vitamin B6 is a relatively simple compound with three substituted
pyridine derivatives that differ only in functional group in the 4-position: alcohol (pyridoxine or pyridoxol), aldehyde pyridoxal, and amine
pyridoxamine (Fig. 9.1). Pyridoxine is the predominant form in plants,
whereas pyridoxal and pyridoxamine are the forms generally found inanimal products. These three forms have equal activity when administered parenterally to animals but are not equivalent when administeredto various microorganisms. Two additional vitamin B6 forms found infoods are the coenzyme forms of pyridoxal phosphate (PLP) and pyridoxamine phosphate. Various forms of vitamin B6 found in animal tissues are interconvertible, with vitamin B6 metabolically active mainly as
PLP and to a lesser degree as pyridoxamine phosphate.
386

V I T A M I N B6

Fig. 9.1 Structural formulas of pyridoxol, pyridoxal, pyridoxamine, pyridoxal phosphate, and 4pyridoxic acid.

Various forms of vitamin B6 can be destroyed by heat and alkali,
and exposure to light, especially in neutral or alkaline media, is highly
destructive. Forms of vitamin B6 are colorless crystals soluble in waterand alcohol as both free bases and commonly available hydrochlorides.
Commercial preparation of vitamin B6 is almost exclusively the hydrochloride salt of the alcohol form, pyridoxine hydrochloride.
Several vitamin B6 antagonists exist that either compete for reactive
sites of apoenzymes or react with PLP to form inactive compounds. The
majority of natural antagonists are substituted hydrazines and hydroxylamines, that is, substances that form hydrozones or oximes withpyridoxal or PLP. Antagonists frequently react with pyridoxal kinase,thus preventing the phosphorylated form of the vitamin. Deoxypyridoxine is a powerful antagonist to vitamin B6 and is commonly employed inexperiments to accelerate the vitamin deficiency. Fortunately, this antivitamin does not occur in nature. A particularly important binding
compound of vitamin B6 is isonicotinic acid hydrazide (isoniazid), which
is used in tuberculosis treatment. Isoniazid is a strong inhibitor of pyridoxal kinase and results in anemia in humans, probably by inhibiting
the synthesis of δ-aminolevulinic acid and thus of heme. Isoniazid has
been extensively used in experiments designed to clarify functions ofvarious enzymes that are dependent on PLP (Price et al., 1957).
387

CHAPTER NINE

The antihypertensive drugs thiosemicarbizide and hydralazine havealso been shown to interfere with vitamin B6 usage. Penicillamine, which
is used to remove body copper in copper poisoning and Wilson’s disease,is also known to complex PLP. L-Dopa, an antiparkinsonism drug, hasbeen shown to be a vitamin B6 antagonist. Other drugs also form thiazalidine compounds or inhibit pyridoxine kinase, the enzyme that isneeded in formation of PLP (Rothschild, 1982). Oral contraceptives (estrogen component) were shown to be antagonistic to vitamin B6, resulting in deficiency of the vitamin (Salkeld et al., 1973). Presence of a vitamin B6 antagonist in flax (linseed oil meal) is of particular interest to
animal nutritionists. This substance was identified in 1967 as hydrazic
acid and was found to have antibiotic properties (Parsons and Klostermann, 1967). Pesticides (e.g., carbaryl, propoxur, or thiram) can be antagonistic to vitamin B6. Feeding a diet enriched with vitamin B6 prevented disturbances in the active transport of methionine in rats
intoxicated with pesticides (Witkowska et al., 1992). The response of
sensitive persons to administration of large doses of monosodium glutamate (“Chinese restaurant syndrome”) was eliminated by vitamin B6
treatment (Folkers et al., 1981), indicating an antagonist to the vitamin.
High doses (more than 2 g) of niacin have been used as lipid-lowering
agents for humans. Niacin at these large dosages was found to interferewith vitamin B status (Basu and Mann, 1997).

ANALYTICAL PROCEDURES
Methods of analysis for vitamin B6 must be capable of detecting all
forms of the vitamin (Gregory, 1988). In biological systems the vitaminis usually bound to protein, and therefore extraction procedures are required. Bioassay using rat or chick growth has the advantage over microbiological procedures in that it measures all biologically active forms
and does not require extraction of bound forms of the vitamin. However, rat and chick bioassay methods are expensive and time-consuming
and have been largely replaced by other techniques.
The standard method for quantitation of vitamin B6 in foods is microbiological assay. Saccharomyces uvarum (formerly S. carlsbergensis)
is the organism commonly used in the microbiological method. Thisyeast organism, along with Streptococcus faecalis and Lactobacillus casei, makes it possible to differentiate between alcohol, amine, and aldehyde vitamin forms. Disadvantages in microbiological methods include
the following: (1) The procedure is time consuming, (2) variability exists
in growth response of various microorganisms to the vitamin, (3) mi388

V I T A M I N B6

croorganisms can mutate, (4) microbiologically unavailable complexesof the vitamin may be formed, and (5) microbial growth may be retarded by substances in the food extract.
Gas-liquid chromatography is used to analyze vitamin B6, butmainly in pure standards. Combinations of chromatographic separation
methods with fluorometric determinations may be used successfully forenriched foods and feeds. Vanderslice et al. (1980) developed a highpressure liquid chromatography (HPLC) anion-exchange system thatquantitatively separated phosphorylated and unphosphorylated formsof vitamin B6 in pork, hamburger, carp, and nonfat dry milk. Methods
using HPLC with fluorometric detection can serve as a simple and reliable method for the analysis of vitamin B6 in foods and feeds (Van
Schoonhoven et al., 1994).

METABOLISM
Digestion, Absorption, and Transport
Utilization of dietary vitamin B6 by animals necessitates digestion
and absorption of the five forms known to occur in foods: pyridoxine,
pyridoxal, pyridoxamine, PLP, and pyridoxamine phosphate. Digestion
would first involve splitting the vitamin, as it is bound to proteins offoods. Vitamin B6 is absorbed mainly in the jejunum, but also in theileum, by passive diffusion. Absorption from the colon is insignificant,even though colon microflora synthesize the vitamin. However, Durst etal. (1989) administered vitamin B6 in the cecum to sows and concluded
that the vitamin was absorbed at this location. Vitamin B6 compounds
are mainly absorbed from the diet in the dephosphorylated forms, but
phosphorylated forms can be absorbed to a very limited extent (Mehansho et al., 1979). The small intestine is rich in alkaline phosphatases forthe dephosphorylation reaction.
After absorption, B6 compounds rapidly appear in liver, where they
are mostly converted into PLP, considered to be the most active vitaminform in metabolism. Unlike PLP, pyridoxamine phosphate is involved
only in several metabolic reactions. It is widely accepted that the synthesis of PLP and pyridoxamine phosphate from the three unphosphorylated forms is mediated by pyridoxal kinase and pyridoxamine phosphate (pyridoxine phosphate) oxidase. Pyridoxamine phosphate and
PLP are also interconvertible through aminotransferases (formerly referred to as the transaminases). Sakurai et al. (1992) reported that aphysiological dose of pyridoxamine was rapidly transformed to pyri389

CHAPTER NINE

doxal in the intestinal tissues and then released in the form of pyridoxal
into the portal blood.
Both niacin (as the NADP-dependent enzyme) and riboflavin (as theflavoprotein pyridoxamine phosphate oxidase) are important for conversion of vitamin B6 forms and phosphorylation reactions (Wada andSnell, 1961; Kodentsova et al., 1993). Vitamin B6 is found in the bloodlargely as PLP, most of which is derived from the liver after metabolism
by hepatic flavoenzymes. Pyridoxal and PLP found in circulation are associated primarily with plasma albumin and red cell hemoglobin(Mehansho and Henderson, 1980). Pyridoxal phosphate accounts for
60% of plasma vitamin B6.
Pyridoxal crosses the human placenta readily in both directions(Delport et al., 1991; Schenker et al., 1992). Gradients of concentration
between maternal and venous cord plasma were 1:5 for PLP (Zempleni
et al., 1992). Alcohol inhibited the transport of pyridoxal from the maternal to fetal compartments by 42% (Schenker et al., 1992).

StorageOnly small quantities of vitamin B6 are stored in the body. The vitamin is widely distributed in various tissues, mainly as PLP or pyridoxamine phosphate. Reports of PLP content of glycogen phosphorylasesuggest that 90% or more of the vitamin B6 present in muscle might bepresent in this single enzyme (Merrill and Burnhan, 1990). Because themuscle accounts for about 40% of body weight, and because the muscle
contains more vitamin B6 per gram than do other tissues (except kidney), muscle phosphorylase may account for as much as 70 to 80% of
the total body store of the vitamin.

Excretion
When B6 coenzymes are synthesized in excess of the binding capacity of B6-dependent apoenzymes, they are dephosphorylated by alkaline
phosphatase, with the resultant pyridoxal reutilized or oxidized to pyridoxic acid (Fig. 9.1) by aldehyde oxidase and/or NAD-dependent dehydrogenase. Pyridoxic acid is the major route for elimination of vitamin
B6, which is excreted in urine. Also, small quantities of pyridoxol, pyridoxal, and pyridoxamine, as well as their phosphorylated derivatives,
are excreted into the urine (Rabinowitz and Snell, 1949; Merrill and
Burnhan, 1990). Mahuren et al. (1991) suggested that 5-pyridoxic acidand its lactone account for 10 to 20% of urinary vitamin B6 metabolites.
Biliary excretion and enterohepatic circulation of vitamin B6 probably
play only a minor role in the overall economy of the vitamin (Lui et al.,390

V I T A M I N B6

1983). Vitamin B6 metabolism is altered in renal failure, as observed inrats exhibiting plasma PLP 43% lower than that of controls (Wei andYoung, 1994).

FUNCTIONS
Vitamin B6 in the form of PLP (also named codecarboxylase), andto a lesser degree pyridoxamine phosphate, plays an essential role in theinteraction of amino acid, carbohydrate, and fatty acid metabolism andthe energy-producing citric acid cycle. More than 60 enzymes are already known to depend on vitamin B6 coenzymes. They typically bindthe coenzyme tightly in a Schiff base linkage with the ε-amino group ofan active-site lysine, and for most of the enzymes, the incoming substratedisplaces the lysine to form a new Schiff base. This results in a strong
electron sink adjacent to several of the bonds on the substrate and facilitates a variety of chemical reactions (Merrill and Burnhan, 1990). Pyridoxal phosphate functions in practically all reactions involved in aminoacid metabolism, including transamination (aminotransferase), decarboxylation, deamination, and desulfhydration, and in the cleavage orsynthesis of amino acids.
The largest group of the vitamin B6-dependent enzymes are thetransaminases, most of which use α-ketoglutarates as the amino group
acceptor. Aminotransferases are involved in interconversions of a pair ofamino acids into their corresponding keto acids; generally, these are αamino and α-keto acids. For example, amino groups are transferredfrom glutamate to pyruvate with formation of α-ketoglutarate and alanine, or from aspartate to α-ketoglutarate, forming oxaloacetate andglutamate. Aminotransferases function in both amino acid biosynthesis(nonessential amino acids) and catabolism. Each aminotransferase isspecific for a specified pair of amino and keto acids functioning as substrates, but is nonspecific for the other pair. The aminotransferases thus
represent an important link between amino acid, carbohydrate, andfatty acid metabolism and the energy-producing citric acid cycle (Fig.
9.2).
Nonoxidative decarboxylation reactions also involve PLP as a coenzyme. Decarboxylases convert amino acids to biogenic amines, such as
histamine, hydroxytyramine, serotonin, γ-aminobutyric acid, ethanolamine, and taurine, some of which are substances of high physiologicalactivity (regulation of blood vessel diameter, neurohormonal actions,and essential components of phospholipids and bile acids). In accord
with their general importance, these enzymatic reactions take place in391

CHAPTER NINE

Fig. 9.2 Pyridoxal phosphate (PLP) and amino acid metabolism.

virtually all organs, most intensively in the liver, heart, and brain. Neurological disorders, including states of agitation and convulsions, result
from reduction of B6 enzymes in the brain, including glutamate decarboxylase and γ-aminobutyric acid transaminase. Maternal restriction of
B6 in rats adversely affected synaptogenesis, neurogenesis and neuron
longevity, and differentiation of the progeny (Groziak and Kirksey,392

V I T A M I N B6

1987, 1990). Research in animal models suggests that vitamin B6 deficiency during gestation and lactation alters the function of N-methyl-Daspartate receptors, a subtype of receptors of the glutamatergic neurotransmitter system thought to play an important role in learning andmemory (Guilarte, 1993).
Vitamin B6 is involved in many additional reactions, particularly
those involving proteins. The vitamin participates in the following functions (Bräunlich, 1974; Marks, 1975; LeKlem, 1991):
1. Deaminases—for serine, threonine, and cystathionine.
2. Desulfhydrases and transulfhyurases—interconversion and metabolism of sulfur-containing amino acids.
3. Synthesis of niacin from tryptophan—hydroxykynurenine is notconverted to hydroxyanthranilic acid but rather to xanthurenic acid becauseof lack of the B6-dependent enzyme kynureninase (see Chapter 8, Functions).
4. Formation of δ-aminolevulinic acid from succinyl-CoA and glycine,the first step in porphyrin synthesis.
5. Conversion of linoleic to arachidonic acid in the metabolism of essential fatty acids (this function is controversial).
6. Glycogen phosphorylase catalyzes glycogen breakdown to glucose
l-phosphate. Pyridoxal phosphate does not appear to be a coenzyme for thisenzyme but rather affects the enzyme’s conformation.
7. Synthesis of epinephrine and norepinephrine from either phenylalanine or tyrosine—both norepinephrine and epinephrine are involved in carbohydrate metabolism as well as in other body reactions.
8. Racemases—PLP-dependent racemases enable certain microorganisms to utilize D-amino acids. Racemases have not yet been detected in mammalian tissues.
9. Transmethylation by methionine.
10. Incorporation of iron in hemoglobin synthesis.
11. Amino acid transport—all three known amino acid transport systems—(a) neutral amino acids and histidine, (b) basic amino acids, and (c)proline and hydroxyproline—appear to require PLP.
12. Formation of antibodies—B6 deficiency results in inhibition of thesynthesis of globulins, which carry antibodies.

Animal and human studies suggest that vitamin B6 deficiency affects
both humoral and cell-mediated immune responses. In humans, vitamin
B6 depletion significantly decreased percentage and total number of lymphocytes, mitogenic responses of peripheral blood lymphocytes to T393

CHAPTER NINE

and B-cell mitogens, and interleukin 2 production (Meydani et al.,1991). Additional human studies indicate that vitamin B6 status may influence tumor growth and disease processes. Deficiency of the vitaminhas been associated with immunological changes observed in the elderly,
persons infected with human immunodeficiency virus (HIV), and thosewith uremia or rheumatoid arthritis (Rall and Meydani, 1993).
Vitamin B6 has other functions, some of which relate directly or indirectly to overall protein metabolism. The vitamin has been shown tofunction in blood pressure regulation (Heseker and Kübler, 1992), reduction of edema and inflammation in rats (Lakshmi et al., 1991), carnitine biosynthesis (Cho and LeKlem, 1990), conversion of selenomethionine to glutathione peroxidase and distribution and transport ofselenium in the body (Beilstein and Whanger, 1989; Yin et al., 1992),
collagen maturation for the connective tissue matrix (Masse et al.,1994), modulate gene expression or transcriptional activation of glycogen phosphorylase in RNA in liver of rats (Oka et al., 1994), detoxification of albizia versicolor poisoning in sheep (Gummow et al., 1992),
and rat enterocyte calcium homeostasis—with a 69% reduction in total
exchangeable calcium and a 56% reduction in cytosol ionized calcium
concentration as a result of vitamin B6 deficiency (Matyaszczyk et al.,1993).

REQUIREMENTS
Requirement for vitamin B6 has been found generally to depend on
species, age, physiological function, dietary components, the intestinal
flora, and other factors that are not yet fully understood. Table 9.1
summarizes the vitamin B6 requirements for various animal species andhumans, with a more complete listing given in the appendix, Table A1.
Because of microbial synthesis, ruminants have no dietary requirement
for vitamin B6. Young ruminants that do not have a fully developed rumen, however, require a dietary source. Considerable quantities of vitamin B6 are synthesized in the relatively large intestinal tract of thehorse, but whether this is adequately absorbed to meet requirements is
controversial. Vitamin B6 is also produced by microorganisms of the intestinal tracts of other animals and humans, but whether significant
quantities are absorbed and utilized is in doubt. Roth-Maier andKirchgessner (1993) suggested that adult sows are able to maintain
optimal metabolic functions over 8 weeks by utilizing bacterially synthesized vitamin B6. Cellulose supplement to these pigs increased total
vitamin B6 excretion from 3.4 to 5.2 mg. In rats administered sul394

V I T A M I N B6
■ Table 9.1

Vitamin B6 Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Leghorn, 0–18 weeks
Laying (100-g intake)
Broilers, 0–8 weeks
All classes
All classes
All classes
Adult
Growing-finishing, 3–5 kg
Growing-finishing, 5–120 kg
Adult
Adult
Adult
Growing
Growing
Growing
GrowingCatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
Children
AdultsMicrobial synthesis
6.5 ppm milk replacer
Microbial synthesis
3.0 mg/kg
2.5 mg/kg
3.0–3.5 mg/kg
2.5–3.0 mg/kg
3.0 mg/kg
3.0–4.5 mg/kg
Microbial synthesis
2.0 mg/kg
1.0–1.5 mg/kg
1.0 mg/kg
Microbial synthesis
Microbial synthesis
1.6 mg/kg
2.0 mg/kg
4.0 mg/kg
60 μg/kg body wt
3.0 mg/kg
5 mg/kg
1–10 mg/kg
6.0 mg/kg
8.0 mg/kg
0.3–0.6 mg/day
1.0–1.4 mg/day
1.7–2.2 mg/day

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1981)
NRC (1982a)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)

ChickenDuck (Pekin)Japanese quail
Turkey
Sheep
Swine
Horse
Goat
Mink
Fox
Cat
Dog
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg/day.

fasalazine, vitamin B6 deficiency was aggravated, suggesting that the intestinal synthesis of the vitamin was affected (Trumbo and Raidi,1991). Animals practicing coprophagy would obviously be receiving vitamin B6 from this source.
Breed of animal and environmental temperature have been shown to
influence vitamin B6 requirements. Lucas et al. (1946) found that crossbred chicks (Rhode Island Red ↔ Barred Plymouth Rock) showed aconsiderably higher requirement for pyridoxine than had previously
been found for White leghorn chicks, and a Japanese strain of chickens
was also shown to have a higher B6 requirement (Scott et al., 1982). Twostrains of mice have likewise been shown to have greatly different requirements for vitamin B6 (Hoover-Plow and Rice, 1992). Regarding
ambient temperature, when rats were housed at 33oC, they needed twice
as much vitamin B6 as when they were housed at 19oC (Bräunlich,1974).
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Quantity of dietary protein affects requirement for vitamin B6 inboth animals and humans. Vitamin B6 requirement is increased whenhigh-protein diets are fed. For example, when feed contained 60% casein instead of 20%, the level of pyridoxine required by mice was threetimes as high (Miller and Baumann, 1945). Gries and Scott (1972) founda much higher vitamin B6 requirement in chicks receiving 31% protein
than in those receiving a normal 22% protein diet. A number of studies
have suggested that amino acid imbalance has an adverse effect on vitamin B6 status in that weight gain was depressed and survival was decreased when large amounts of a single amino acid were added to rat diets limited in the vitamin. High tryptophan, methionine, and otheramino acids increase the need for vitamin B6 (Scott et al., 1982). Fisher
et al. (1984) reported a consistent deleterious effect of a low-quality protein on vitamin B6 status in rats. Certain feed antagonists (see Chemical
Structure, Properties, and Antagonists), bioavailability of B6 in feeds (seeNatural Sources), and nutrients other than protein influence the B6 requirement. Niacin and riboflavin are needed for interconversions of different forms of vitamin B6, with an overdose of thiamin reported to produce deficiency of the vitamin in rats (LeKlem, 1991).
In humans, vitamin B6 requirements are increased with higher dietary protein, pregnancy, lactation, oral contraceptives, alcoholic beverages, liver disease, and certain drugs (see Chemical Structure, Properties,
and Antagonists), in the elderly population, and for some individuals
with inborn errors of metabolism. The allowance for vitamin B6 is approximately 2.0 mg/day for adult males and 1.6 mg/day for adult females (RDA, 1989). The RDA for infants begins at 0.3 mg/day, increases
to 0.6 mg/day for older infants consuming a mixed diet, and increases
from 1 to 2 mg/day from childhood through adolescence. Pregnancy andlactation add 0.6 and 0.5 mg/day, respectively, to the RDA for women.
Low milk vitamin B6 has been associated with long-term use of oral contraceptives (estrogen component). Both the use of oral contraceptives
and alcoholism result in lowered plasma PLP levels. Low plasma PLPlevels in alcoholics are believed to result from acetaldehyde, which reduces the protein binding of PLP, thereby promoting its dephosphorylation and oxidation to pyridoxic acid. In studies with vitamin B6-deficient
rats, alcohol affected fetal weight and reduced maternal liver PLP by
22% (Lin, 1989). Some individuals have inborn errors of metabolismthat greatly increase their B6 requirement. Fortunately, the number ofsuch individuals appears to be small, but unless they take doses of B6
starting at birth, they develop convulsions and brain damage, and die(Scriver and Whelan, 1969).
396

V I T A M I N B6

NATURAL SOURCES
Vitamin B6 is widely distributed in foods and feeds (Table 9.2). In
general, muscle meats, liver, vegetables, whole-grain cereals and theirby-products, and nuts are among best sources; few materials are reallypoor sources, with the exception of fruits. The vitamin present in cereal
grains is concentrated mainly in bran, the rest containing only small
amounts. The richest source is royal jelly produced by bees (5,000 μg/g).
Most vitamin B6 in animal products is in the form of pyridoxal and pyridoxamine phosphates. In plants and seeds the usual form is pyridoxol.
Many analytical figures for vitamin B6, especially older ones, were too
low because the assays did not measure all the biologically active forms(Scott et al., 1982).
The level of vitamin B6 contained in all feeds is affected by processing and subsequent storage. Of the several forms, pyridoxine is far morestable than either pyridoxal or pyridoxamine. Therefore, the processing
losses of vitamin B6 tend to be highly variable (0 to 40%), with plantderived foods (which contain mostly pyridoxine) losing little if any ofthe vitamin and animal products (mostly pyridoxal and pyridoxamine)
losing large quantities. Vitamin B6 loss during cooking, processing, refining, and storage has been reported to be as high as 70% (Shideler,1983) or in the range of 0 to 40% (Birdsall, 1975). Losses may be■ Table 9.2

Vitamin B6 in Foods and Feedstuffs (mg/kg, dry basis)

Alfalfa leaves, sun cured
4.4
Barley, grain
7.3
Bean, navy (seed)
0.3
Blood meal
4.8
Brewer's grains
0.8
Buttermilk (cattle)
2.6
Copra meal (coconut)
4.8
Corn, gluten meal
8.8
Corn, yellow grain
5.3
Cottonseed meal, solvent extracted 6.8
Crab meal
7.2
Fish meal, anchovy
5.0
Fish meal, menhaden
5.1
Horse meat
0.7
Linseed meal, mechanically
6.1
extracted
Liver, cattle
18.0
Milk, skim, cattle
4.5

Molasses, sugarcane
Oat, grain
Oat, groutsPea, seeds
Peanut meal, solvent extracted
PotatoRice, grain
Rice, polished
Rye, grain
Sorghum, grain
Soybean meal, solvent extracted
Spleen, cattle
Wheat, bran
Wheat, grain
Whey
Yeast, brewer's
Yeast, torula

5.7
2.8
1.2
1.7
6.9
15.5
5.0
0.4
2.9
5.0
6.7
1.3
9.6
5.6
3.6
39.8
38.9

Source: NRC (1982b).

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caused by heat, light, and various agents that can promote oxidation.
Two-hour sunlight exposure may destroy half the vitamin B6 in milk.
Blanching of rehydrated lima beans resulted in a loss of 20% vitamin B6,
but more significantly, the availability of the vitamin was reduced by almost 50% (Ekanayake and Nelson, 1990). Irradiation as a potential
method for microbial control of poultry feed results in a loss of 15% vitamin B6 potency (Leeson and Marcotte, 1993).
In the early 1950s several infants in the United States were diagnosed as deficient in vitamin B6 as a result of consuming a commerciallysterilized, liquid milk formula. Later studies showed that the vitamin underwent rapid destruction during the autoclaving procedure used in the
production of formulas. Also, human milk often does not contain sufficient vitamin B6 for the newborn (see Deficiency, Effects of Deficiency,
Humans). However, human milk could be an adequate source of vitamin
B6 because vitamin levels respond rapidly to dietary intake (Kirksey,
1981; Kang-Yoon et al., 1992). Milk vitamin B6 concentrations fromgoats and cows were approximately twice that of human values (Coburn
et al., 1992).
Data on vitamin B6 content of feeds are generally insufficient and information on the vitamin’s bioavailability is lacking. Bioavailability of
B6 was found to be greater from beef than from corn meal, spinach, or
potatoes (Nguyen and Gregory, 1983). LeKlem et al. (1980) reported
that in young adult men, vitamin B6 bioavailability in whole-wheat
bread is lower than that in white bread enriched with pure pyridoxine.
Vitamin B6 in whole-wheat bread and peanut butter was 75 and 63%,
respectively, as available as that from canned tuna (Kabir et al., 1983).
In studies with young men based upon plasma PLP level or urinary B6(i.e., 4-pyridoxic acid) excretion, B6 bioavailability in the average American diet was estimated to be about 75% (Tarr et al., 1981).
Pyridoxine-5´-β-D-glucoside (PNG), a conjugated form of vitamin
B6, has been shown to be abundant in various plant-derived foods. This
form of B6 may account for up to 50% of the total vitamin B6 content
of oilseeds such as soybeans and sunflower seeds. The utilization of dietary PNG, relative to pyridoxine, has been shown to be 30% in rats
(Trumbo et al., 1988) and 50% in humans (Gregory et al., 1991). In
suckling rats, the availability of vitamin B6 derived from PNG is only
25% compared with pyridoxine (Trumbo and Gregory, 1989). Pyridoxine-5´-β-D-glucoside seems to be well absorbed, but the hydrolysis
to pyridoxine is poor. The primary site of limited conversion to pyridoxine is the intestinal mucosa, where β-glucosidase activity toward
PNG has been demonstrated (Gregory et al., 1991). The glycosylated398

V I T A M I N B6

PNG can quantitatively alter the metabolism of pyridoxine in vivo;
hence, it partially impairs the metabolic utilization of co-ingested nonglycosylated forms of vitamin B6 (Gilbert and Gregory, 1992; Zhang etal., 1993; Nakano and Gregory, 1995; Hansen et al., 1996; Nakano etal., 1997).

DEFICIENCY
Effects of Deficiency
Characteristics of vitamin B6 deficiency are retarded growth, dermatitis, epileptic-like convulsions, anemia, and partial alopecia. Becauseof the predominant function of the vitamin in protein metabolism, in vitamin B6 deficiency a fall in nitrogen retention is observed, feed protein
is not well utilized, nitrogen excretion is excessive, and impaired tryptophan metabolism may result.

Ruminants
Amounts of vitamin B6 that are normally synthesized by microorganisms in the digestive tract of ruminants, when fully developed, areenough to meet needs of these animals. McElroy and Goss (1939) reported vitamin B6 to be 6 to 10 mg/kg of diet in dried rumen contents
of mature sheep, even though the diet contained only 1.5 mg/kg. Therefore, no deficiency signs of vitamin B6 have yet been observed in matureruminants. However, stressed cattle, as a result of long-distance transportation, had very low blood pyridoxine concentrations (Dubeski andOwens, 1993).
Vitamin B6 has been shown to be essential for the young calf whenselected experimental diets are used. Calves that were reared on a “milksubstitute” lost appetite within 2 to 4 weeks; their growth was impaired;
and they progressively showed apathy, diarrhea, loss of appetite, andloss of coordination. In the last stages, convulsions were soon followed
by death (Johnson et al., 1947). The convulsions included thrashing of
the legs and head and grinding of the teeth (Johnson et al., 1950).
Postmortem examination of vitamin B6 deficient calves revealed
hemorrhages in the epicardium and in the kidneys, demyelination of theperipheral nerves, proliferation of Schwann cells, desquamation of theintestinal mucosa, and pneumonia (Johnson et al., 1950). Analyses of
the urine from the vitamin B6-deficient calves showed greatly reduced
excretion of pyridoxine, pyridoxal, pyridoxamine, and pyridoxic acid(major excretory metabolite).
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Swine
In growing pigs, clinical signs of vitamin B6 deficiency include apoor appetite, slow growth (Fig. 9.3), microcytic hypochromic anemia,
epileptic-like seizures or convulsions (Fig. 9.4), fatty infiltration of theliver, diarrhea, rough hair coats, scaly skin, a brown exudate around theeyes, demyelination of peripheral nerves, and subcutaneous edema(Bauernfeind, 1974; Bräunlich, 1974; Cunha, 1977). Like some other vitamins, vitamin B6 deficiency reduces the immune responses of the pig(Harmon et al., 1963). The first and most conspicuous sign in baby pigs
that vitamin B6 is insufficient is a loss of appetite that, if the deficiency
is severe, may appear in less than 2 weeks. This is accompanied by reduced growth, vomiting, diarrhea, and a peculiar compulsion to lick.
When deficiency of vitamin B6 reaches an advanced stage (probably
due to degeneration of the peripheral nerves), disordered movement andataxia appear. Finally, convulsions develop at irregular intervals, but are,apparently, stimulated by excitement, because they are most often observed at feeding time. Between these convulsions, pigs lie down and areapathetic and unresponsive (Bräunlich, 1974). Bräunlich suggested thata vitamin B6 deficiency may go unnoticed in swine because of a lack ofvisible signs associated specifically with the deficiency. Metabolic disor-

Fig. 9.3 A 6-week-old B6-deficient pig weighing only 3.6 kg. (Courtesy ofR.W. Luecke and E.R. Miller, Michigan Agriculture Experiment Station, andJ. Nutr. 62 [1957], 405.)
400

V I T A M I N B6

Fig. 9.4 This pig is having an epileptic-like seizure while receiving a diet low invitamin B6. (Courtesy of E.H. Hughs and H. Heitman, California Agriculture
Experiment Station.)ders may be revealed only by poor appetite, slow growth, and inefficient
feed utilization. In some experiments with vitamin B6, protein retention
by pigs deficient in the vitamin was reduced to less than half.
During reproduction and lactation, sows fed a corn-sorghum-soybean meal diet responded to vitamin B6 supplementation at a level of 4.4
mg/kg of feed (Adams et al., 1967). Vitamin B6 supplementation of 11
mg/kg of feed produced a slightly superior daily gain in weight, more
piglets born alive, and a smaller number of resorbed fetuses comparedwith control sows that received only 1 mg/kg of vitamin B6 (Ritchie etal., 1960).

Poultry
Chicks fed a vitamin B6-deficient diet have little appetite and growslowly, with plumage failing to fully develop. Chicks maintained on a
B6-deficient diet exhibited general weakness after a few days of deprivation. The birds squat in a characteristic posture, with wings slightly
spread and head resting on the ground (Bräunlich, 1974). Miller (1963)
observed high proportions of pendulous crops with vitamin B6-deficient
chicks.
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A more specific sign of B6 deficiently is the nature of the nervous
conditions that develop. Deficient chicks are abnormally excitable. As
deprivation continues, nervous disorders become increasingly severe(Bräunlich, 1974). There is a trembling and vibration of the tip of thetail, with movement stiff and jerky. Chicks will run aimlessly about withlowered head and drooping wings. Finally, convulsions develop, during
which chicks fall on their side or back, with the legs scrabbling. Violent
convulsions cause complete exhaustion and may lead to death. These
clinical signs may be distinguished from those of encephalomalacia by
the relatively greater intensity of activity during a seizure (Scott et al.,1982).
Blood alterations are also typical of a vitamin B6 inadequacy inchicks. An extreme deficiency leads to a microcytic, polychromatic
hypochromic anemia in conjunction with atrophy of the spleen and thymus (Asmar et al., 1968). Marginal deficiencies provoke a microcytic,
normochromic polycythemia (Blalock and Thaxton, 1984), and deficient chicks show decreased immunoglobulin M and immunoglobulin G
response to antibody challenge (Blalock et al., 1984).
Similar signs of a vitamin B6 deficiency have been observed in turkey
poults—loss of appetite, poor growth (Fig. 9.5), oversensitivity andcramps, and finally death. Ducklings not receiving enough vitamin B6
grow slowly, and development of plumage is poor. At 5 days of age,
ducklings showed retarded growth (Yang and Jeng, 1989). Clinical signs
were first observed at 7 days of age and were characterized by decreased
appetite, extreme weakness, hyperexcitability, convulsion, and death.
Hematological examination at 3 weeks of age indicated that pyridoxine
deficiency in ducklings resulted in microcytic, hypochromic anemia.
Signs of B6 deficiency in chicks will appear very rapidly after introduction of a B6-deficient feed. Fuller and Kifer (1959) reported that signsof a deficiency appeared on the eighth day. Chronic, borderline B6 deficiency produces perosis. Usually one leg is severely crippled, and one orboth of the middle toes may be bent inward at the first joint (Gries andScott, 1972). A marked increase in gizzard erosion was found in vitamin
B6-deficient chicks (Daghir and Haddad, 1981).
In adult poultry, vitamin B6 deficiency results in reduced egg production and hatchability as well as decreased feed consumption, weight
loss, and death. Robel and Christensen (1991) reported that hatchability of pyridoxine-injected white turkey hen eggs was 4.2% higher than
non-injected eggs. A severe deficiency (levels of vitamin B6 below 0.5
mg/kg of diet) causes rapid involution of the ovary, oviduct, comb, and402

V I T A M I N B6

Fig. 9.5 Vitamin B6-deficient poult (about 4 weeks old) on left and a normal
poult on right. (Courtesy of T.W. Sullivan, University of Nebraska.)

wattles in mature laying hens. Involution of testes, comb, and wattle occurs in vitamin B6-deficient adult cockerels (Scott et al., 1982).

HorsesNo experimental information is available on vitamin B6 requirements or deficiency in the horse. It has been shown that the horse synthesizes vitamin B6 in the intestinal tract, with the assumption that adequate quantities are absorbed. However, some researchers believe that
racehorses need vitamin B6 supplementation because when these horses
undergo intensive training, they need a high proportion of protein in
their diets that would increase B6 requirements (Bräunlich, 1974).

Other Animal Species
DOGS AND CATS
A typical consequence of B6 deficiency in both young and old dogs
is a microcytic, hypochromic anemia. In addition to decreased appetiteand body weight loss, pathological changes include ataxia, cardiac dilatation and hypertrophy, congestion of various tissues, and demyelina-403

CHAPTER NINE

tion of peripheral nerves (NRC, 1985a). Acute deficiency of vitamin B6
in growing puppies may lead to sudden death with clinical signs of only
anorexia, slow growth, or body weight loss.
In kittens deficient in B6, there is growth depression, a mild microcytic, hypochromic anemia, convulsive seizures, and irreversible kidney
lesions consisting of areas of tubular atrophy and dilatation (NRC,1986). Vitamin B6 deficiency has been reported to produce behavioral,
neurophysiological, and neuropathological abnormalities in a variety of
species, including cats. Buckmaster et al. (1993) used brainstem auditory
evoked potentials and determined that vitamin B6 deficiency in cats affected peripheral and brainstem auditory pathways. The finding of prolonged interwave intervals in vitamin B6-deficient cats was consistent
with slowed axonal conduction velocity secondary to defective myelination.

FISH
Most fish are sensitive to vitamin B6 deficiency and show neurological disorders in 3 to 8 weeks (Lovell, 1987). Clinical signs in catfish include hyperirritability, erratic swimming, anorexia, tetany, and greenishblue coloration. Erratic swimming, hyperirritability, and convulsions
have been observed in salmonids, gilthead sea bream, common carp, yellowtail, and Japanese eel (NRC, 1993). Other deficiency signs, such as
anorexia and poor growth, usually appear in the fish within 3 to 6
weeks after they have been fed a vitamin B6-deficient diet. Pyridoxine
deficiency has been reported to cause various histopathological changes
in rainbow trout liver and kidney and in the intestinal tissue of both
rainbow trout and gilthead sea bream.

FOXES AND MINK
Signs of deficiency in growing kits appeared after about 2 weeks ona B6-deficient diet and included reduced feed intake, weight loss, diarrhea, brown exudate around the nose, excessive lacrimation, swellingand puffiness around the nose and face region, apathy, muscular incoordination, convulsions, and finally death (NRC, 1982a). Reproductionis also affected in mink with testes degeneration and resorption of embryos. Vitamin B6 deficiency in the fox results in anorexia, growth cessation, and a decrease in hemoglobin.

LABORATORY ANIMALS
Rats fed diets deficient in vitamin B6 develop symmetrical scalingdermatitis (acrodynia) on the tail, paws, face, and ears; microcytic ane404

V I T A M I N B6

mia; hyperexcitability; convulsions; and reduced reproductive performance in both sexes (NRC, 1995). Edema was increased 54% in vitamin
B6-deficient rats (Lakshmi et al., 1991). Subtle abnormalities of gait, consisting of a reduced width of step was detected early in vitamin B6-deficient rats (Schaeffer et al., 1990). Sexual behavior of vitamin B6-deficientmale rats was exhibited as reduced post-ejaculatory intervals and an increased proportion of animals ejaculating (Gorzalka and Moe, 1994).
For mice, vitamin B6 deficiency signs include poor growth, hyperirritability, posterior paralysis, necrotic degeneration of the tail, alopecia,
and progressive hypochromic anemia (NRC, 1995). Loss of appetite isan early and constant result of B6 deficiency in hamsters. Although acrodynia is characteristic of vitamin B6 deficiency in rats, it is not found in
hamsters. The fur of B6-deprived hamsters has an unkempt appearance,and crusted lesions are occasionally observed on lips and mouth (NRC,1995).

NONHUMAN PRIMATES
Vitamin B6 deficiency signs in rhesus monkeys consist of weight
loss, hypochromic, microcytic anemia, apathy, and ataxia. There is
widespread arteriosclerosis, fatty metamorphosis of liver, hepatic necrosis and cirrhosis, dental caries, oxaluria, and neuronal degeneration of
the cerebral cortex (NRC, 1978). With cebus monkeys, deficiency consists of weight loss, a profound hypochromic, microcytic anemia, hairloss, and dermatitis, especially about the hands and toes.

RABBITSIn rabbits, vitamin B6 deficiency signs include a scaly thickening of
the skin of ears and inflammation around eyes and nose. In a few cases
there is a pronounced acrodynia, with severe encrustation and inflammation of nose, eyes, and paws. Alopecia develops most often on
forelegs and is accompanied by skin inflammation and desquamation(Bräunlich, 1974). Neurological signs observed include convulsions,partial paralysis, and contracture of extremities. In the advanced stage,some rabbits are unable to move their hind legs at all, the muscles of thehind leg having become severely atrophied.

HumansA high proportion of the human population receives inadequate dietary vitamin B6, particularly young and pregnant or lactating women.
Typical vitamin B6 deficiency symptoms in humans include
hypochromic, microcytic anemia, loss of weight, abdominal distress,405

CHAPTER NINE

vomiting, hyperirritability, epileptic-type convulsions in infants, depression and confusion followed by convulsions in adults, and electroencephalographic abnormalities (LeKlem, 1991). Seborrhea-like lesions
developed about the eyes, nose, and mouth, and some subjects showedcheilosis and glossitis. A depression of the lymphocyte count was a fairly
constant finding.
There are a large number of conditions in which vitamin B6 metabolism is altered. Conditions in which tryptophan metabolism has beenaltered and in which vitamin B6 administration was used include
asthma, diabetes, certain cancers, pellagra, and rheumatoid arthritis,
while diseases and pathological conditions in which plasma PLP levels
have been shown to be depressed include asthma, diabetes, renal disorders, alcoholism, heart disease, pregnancy, breast cancer, Hodgkin’s disease, and sickle-cell anemia (LeKlem, 1991; Driskell, 1994). Animal andhuman studies suggest that vitamin B6 deficiency affects both humoral
and cell-mediated immune responses. Lymphocyte differentiation andmaturation are altered by deficiency, delayed-type hypersensitivity responses are reduced, and antibody production may be indirectly impaired. Deficiency of the vitamin has been associated with immunological changes observed in the elderly, persons infected with HIV, and thosewith uremia or rheumatoid arthritis (Rall and Meydani, 1993).
Studies conducted during the 1990s have suggested that in the
United States, a large percentage of women of child-bearing age andpregnant and lactating women have vitamin B6 intakes significantly
lower than the recommended dietary allowance (Guilarte, 1993). Vitamin B6 deficiency has been reported in 83.4% of Turkish children(Wetherilt et al., 1992) and 34% of adolescents in Nigeria (Korede,1990). In the elderly Greek population, dietary intake of vitamin B6 wasless than 50% of the U.S. RDA (Kafatos et al., 1993).
Convincing evidence of the essential nature of vitamin B6 in humannutrition was first provided by reports in the United States of infants
who developed irritability and convulsive seizures (and sometimes died)when fed autoclaved infant formulas. This autoclaving destroyed muchof the vitamin B6, and infants did not receive adequate quantities of thevitamin. Infants receiving human milk are also sometimes at risk as generally this is a poor source of the vitamin. Other conditions responsive
to B6 first seen in infants are several inborn errors of metabolism resulting in vitamin B6 dependency.
Reports indicate that unsupplemented mothers have a milk vitamin
B6 level that in some cases is lower than 100 μg/L, a concentration thatplaces infants at risk for development of seizures (Styslinger and Kirk406

V I T A M I N B6

sey, 1985; Guilarte, 1993). Although this concentration of vitamin B6 inmilk does not always result in clinical signs of apparent vitamin B6 deficiency, it may influence the normal development of the child. Evidence
for such an effect was noted in the Egyptian study (McCullough et al.,1990), in which abnormalities in behavior were observed in infants
whose mothers had vitamin B6 levels in milk below 85 μg/L. Likewise,
these mothers were less responsive to their infants’ vocalizations,showed less effective intervention to infant distress, and were more
likely to use older siblings as caregivers than were mothers of better vitamin status.
Both pregnancy and use of oral contraceptives increased the vitamin
B6 requirement (Miller, 1986). Contractor and Shane (1970) discoveredthat pregnant women had lower blood levels of PLP than did nonpregnant controls. Brin (1971) reported that pregnant women had increased
urinary metabolites of tryptophan. He also found fetal blood levels of B6
two to three times greater than the maternal level, indicating a depletionof the mother’s B6 stores. According to Heller et al. (1973), vitamin B6
supplementation is necessary in approximately 50% of pregnant women
to maintain normal B6 coenzyme saturation of aspartate aminotransferase.
Alcohol and certain drugs (see Chemical Structure, Properties, andAntagonists) interfere with the metabolic functions of vitamin B6. Vitamin B6 deficiency is viewed as an important nutritional complication inalcoholism, and it has been established that alcoholics absorb less vitamin B6 than do control subjects. Alcoholics have a decreased ability toliberate vitamin B6 from its bound form, and several studies suggest that
alcohol interferes with conversion of pyridoxine to PLP (Bonjour, 1980).
Certain drugs, including the antituberculosis drug isonicotinic acid hydrazide (INH), have been shown to increase need of vitamin B6. INH
forms hydrazone complexes with PLP, which inactivates the coenzyme(Roe, 1973). Studies suggest that vitamin B6 availability is limited in hyperthyroidism, and consequently B6 should be added to antithyroid
medication for hyperthyroid patients (Wohl et al., 1960). The use of interleukin-2 for treatment of various cancers depressed vitamin B6 bloodconcentration to subnormal in 90% of patients (Baker et al., 1992).
Vitamin B6 is affected by uremia and urinary calculi and has been
associated with inadequacy of the vitamin. Many symptoms of uremia
are quite similar to vitamin B6 deficiency. Convulsions, peripheral neuritis, and depression of the immune response are seen in both conditions.
Dobbelstein and coworkers (1974) suggested that uremic toxins exert an
inhibitory effect on PLP. The increase in oxalic as well as xanthurenic407

CHAPTER NINE

acid in the urine of B6-deficient humans has been suggested as a cause
for urinary calculi. In a study by Lilum et al. (1981), 85% of 100 rats
produced calcium oxalate stones within 6 weeks of being placed on a vitamin B6-free diet.

Assessment of Status
Several biochemical procedures have been developed and utilized
for assessment of vitamin B6 status of humans and animals. Indices forevaluating vitamin B6 status and suggested values for adequate status inadult humans have been reported (LeKlem, 1990). Vitamin B6 deficiency
leads to an impairment of protein utilization even before visible signs occur. A widely used approach to assess vitamin B6 status in humans ismeasuring urinary metabolites of tryptophan following a tryptophanload test. In a B6 deficiency, tryptophan is not converted to niacin and ismetabolized to other intermediate products at a greater rate than in a
nondeficient state. Metabolites most often measured are xanthurenic
acid and kynurenic acid. Although the tryptophan load test has been
widely used in assessment of vitamin B6 status, results obtained via thismethod should be interpreted with care, as several other metabolic andhormonal factors are known to be involved in tryptophan metabolism.
The tryptophan load test is a good parameter for the assessment of thevitamin B6 status of population groups. Some research suggests that thistest may be less useful in detecting marginal vitamin B6 intake.
Direct measurements of one or more forms of vitamin B6 in plasma,urine, or erythrocytes are relatively reliable indicators of vitamin B6 status. Urinary 4-pyridoxic acid (4-PA) excretion is considered a short-term
indicator of vitamin B6 status due to the fact that 4-PA reflects recent vitamin B6 intake rather than the underlying state of tissue reserve (Lui etal., 1985). Often 4-PA is not detectable in the urine of vitamin B6-deficient subjects. One of the most commonly used measures of vitamin B6
status is the measurement of erythrocyte alanine and aspartic acidtransaminase(s) activity and/or stimulation (LeKlem, 1990). Because of
the nature of the life span of the erythrocyte, they are considered longterm measures of vitamin B6 status. The specific tests employed are themeasurement of the basal activity of these transaminases and the activity in the presence of excess PLP.
Over the past 20 years, plasma PLP has been the most frequently
used direct measure of vitamin B6 status. This measure has replaced thetryptophan load test as the method of choice in evaluating status.
Plasma PLP is the primary form of circulating vitamin B6 and comprises
70 to 90% of the total vitamin B6 in the plasma. Plasma pyridoxal has
408V I T A M I N B6

been suggested as an additional index of vitamin B6 status (LeKlem,
1991).
Other factors may influence plasma PLP and should be consideredwhen using this index as a measure of vitamin B6 status. These include
the physiological variables of age, exercise, pregnancy, and dietary components, particularly protein. Plasma PLP has been shown to decrease
with advancing age (Löwik et al., 1989), with an age-dependent difference in protein intake related to vitamin B6 requirements. Elderly subjects apparently need less vitamin B6 at higher protein intake based on
plasma PLP, pyridoxal, and total vitamin B6 concentrations (Pannemans
et al., 1994).
Other vitamin B6 status indicators might include concentrations of
the vitamin in milk and eggs. Pyridoxine intakes of breast-fed infants reflected the amount of their mother’s supplement (Kang-Yoon et al.,1992). Pyridoxine intake of mothers was a strong indicator of infant
pyridoxine status. Concentrations of vitamin B6 in hen egg albumen increased with four incremental dietary pyridoxine levels (Robel, 1992).

SUPPLEMENTATION
Vitamin B6 is one of the B vitamins that is least likely to be deficientin livestock diets, particularly ruminants. However, a supplemental useof vitamin B6 was reported in that 40 to 50 mg pyridoxine hydrochloride/kg body weight administered to sheep was successful in counteracting toxicity of the toxic plant Albezia versicolor (Gummow et al., 1992).
Only before the rumen is developed should B6 supplementation for ruminants be considered. Young ruminants receiving mother’s milk areprovided with sufficient quantities of the vitamin. Bräunlich (1974) suggested that the best possible concentration of vitamin B6 in milk substitutes for calves is probably higher than 2.4 mg/kg. Because of its wide
dietary distribution, vitamin B6 is seldom deficient in typical swine andpoultry diets. Evidence indicates that corn, soybean meal, and other ingredients used to supply energy and protein in practical diets for these
species usually also provide sufficient vitamin B6. Consequently, it is notoften added in supplemental form. However, the bioavailability of vitamin B6 in corn and soybean meal ranges from only 45 to 65% (Hoffmann-La Roche, 1979).
Under certain conditions vitamin B6 supplementation is warranted
for practical growing and breeding diets for poultry and other monogastric or simple-stomached animals. Fuller et al. (1961) believed that
although corn-soybean meal practical poultry breeder diets probably
409

CHAPTER NINE

contain sufficient vitamin B6 to support hatchability, there is little margin of safety. The amount of supplemental vitamin B6 recommended formonogastric species varies from 1 to 10 mg/kg of diet depending on
species, age, activity, stress of performance, and field use experience(Bauernfeind, 1974). Reasons for needed supplementation of vitamin B6
include the following (Perry, 1978): (1) great variations in amounts of B6
in individual ingredients, (2) variable bioavailability of this vitamin in
ingredients, (3) losses reported during processing of ingredients, (4) discrepancies in activity for test organisms versus those for animals, (5) ahigher pyridoxine requirement due to a marginal level of methionine in
the diet, and (6) high-protein diets.
Variability of vitamin B6 in feeds depends on the sample origin, conditions of growth, climate, weather conditions, and other local factors(see Natural Sources). For products of animal origin, concentration depends on level of nutrition and environment of original animals. Yen etal. (1976) determined available vitamin B6 in corn and soybean meal using a chick growth assay. Corn was found to be 38 to 45% available,
and B6 in soybean meal 58 to 62% available. There was little differencein availability between corn samples not heated and those heated to
120oC. However, corn heated at 160oC contained significantly lessavailable B6. Level of vitamin B6 contained in feedstuffs is also affectedby processing and subsequent storage. In one report a loss of 30% of B6
contained in alfalfa meal during the coarse-milling and pelleting
processes was observed (Bräunlich, 1974). Ink and Henderson (1984)
reported that bioavailability can be as low as 40 to 50% after heat processing of foods.
Predominant losses of vitamin B6 activity in food occur in the pyridoxal and pyridoxamine forms, with pyridoxine being the more stable
form. Supplemental vitamin B6 is reported to have a higher bioavailability and stability than the naturally occurring vitamin, which in retorted milk products exhibited only 50% of the bioavailability of synthetic B6 or B6 in formulas that were fortified with the vitamin prior tothermal processing (Tomarelli et al., 1955). Heat sterilization of milk reduced vitamin B6 activity by irreversible binding to protein, as well as bypartial formation of 4-PA (Pingali and Trumbo, 1992).
Commercially, vitamin B6 is available as crystalline pyridoxine hydrochloride and various dilutions. Pyridoxine hydrochloride contains
82.3% vitamin B6 activity. The dry dilution is used in feeds, while thecrystalline product is used in parenteral and oral pharmaceuticals as wellas in feeds. For human nutrition, pyridoxine hydrochloride is used by410

V I T A M I N B6

pharmaceutical companies in the preparation of capsules, tablets, andampoules. The tablets used for prophylactic purposes usually contain 2
mg per daily dose. For therapeutic purposes, 10- to 150-mg tablets aretaken one to three times daily.
The recovery of vitamin B6 as pyridoxine hydrochloride in a multivitamin premix not containing trace minerals was 100%, even after 3
months in storage at 37oC. However, stability in a premix containing
trace minerals was poor, with only 45% recovery after 3 months at 37oC
(Adams, 1982). Verbeeck (1975) found vitamin B6 to be stable in premixes with minerals as sulfates. However, if minerals in the form of carbonates and oxides are used, 25% of the vitamin can be lost over a 3month period. Stress agents such as choline chloride help catalyze thisdestruction. Gadient (1986) considers pyridoxine to be very sensitive toheat, slightly sensitive to moisture and light, and insensitive to oxygen.
Retention of B6 activity in pelleted feeds after 3 months at room temperature should be 80 to 100% as a general rule. The retention of pyridoxine in an extruded fish meal fed after 1 month at room temperature
was found to be 56%.
Data have accumulated for the need of vitamin B6 supplementation
for humans, especially for young and pregnant or lactating women. Requirements for B6 are higher for individuals during pregnancy, use oforal contraceptives, certain drug therapy, radiation sickness, overuse ofalcohol, hyperthyroidism, uremia, urinary calculi, and in errors of metabolism (see Deficiency). Vitamin B6 supplements are commonly given
when isoniazid is used in tuberculosis treatment and when penicillamine
is used in the treatment of Wilson’s disease. Vitamin B6 supplements arealso frequently given along with most anticonvulsant drugs. Reports indicate that mothers unsupplemented with vitamin B6 produce milk low
in the vitamin (Guilarte, 1993). Based on these studies, it is apparent
that some degree of vitamin B6 deficiency may be present in infants
whose sole source of nutrients is breast milk and whose mothers are notsupplemented with the vitamin (see Deficiency). Supplemental vitamin
B6 is needed by persons with the “Chinese restaurant syndrome,” a condition in which individuals are sensitive to foods heavily seasoned withmonosodium glutamate (Folkers et al., 1981). Individuals with carpal
tunnel syndrome (pain and/or numbness in hands) require vitamin B6
well in excess of the RDA requirement (LeKlem, 1991). Pyridoxine inpharmacological doses was useful in the management of kidney stones,
decreasing urinary oxalate excretion in patients with recurrent oxalate
renal calculi (Mitwalli, 1989). In doses of 10 to 25 mg, vitamin B6 in-411

CHAPTER NINE

creases the conversion of L-dopa to dopamine, which, unlike its precursor, is unable to cross the blood-brain barrier. Thus, vitamin B6 interfereswith L-dopa in the management of Parkinson’s disease.
Vitamin B6 is one of several vitamins currently popular in “orthomolecular” megadose therapy. It has been used for a variety of conditions, including premenstrual syndrome and behavioral disorders. Pyridoxine tablets are available over the counter in dosages of 30 to 500 mg.

TOXICITY
In common with other members of the vitamin B complex, vitamin
B6 has a very low toxicity. Prolonged feeding of pyridoxine-HCl to rats
(2.5 mg per day), puppies (20 mg per kilogram body weight per day),and monkeys (10 mg per kilogram body weight per day) produced no
toxic signs (Unna and Antopol, 1940).
Large doses of vitamin B6 can produce detrimental effects in experimental animals and humans. Signs of toxicity occur most obviously inthe peripheral nervous system and include changes in gait and peripheral
sensation (Krinke and Fitzgerald, 1988). Changes in central nervous system function were detected in rats fed excess vitamin B6 using techniquesof startle behavior measurement (Schaeffer, 1993).
Rats, rabbits, and dogs tolerated 1 g/kg body weight without ill effects; however, larger doses produced adverse manifestations in all three
species. Marked impairment of coordination and/or righting reflexes
were observed within about 3 days, followed by severe convulsions,paralysis, and death (Unna and Antopal, 1940). Neurohistological examination of rats and dogs that had received 2 to 6 g/kg body weight ofthe vitamin revealed degeneration of the spinal cord and peripheral
nerves (Antopol and Tarlov, 1992). Testicular damage from high dosesof vitamin B6 to rats was reported (Kaido et al., 1991), including retardation in spermiation and Sertoli cell alterations.
Pyridoxal is apparently twice as toxic as pyridoxine or pyridoxamine. Evidence from dog and rat studies suggests that more than 1,000
times the requirement would be needed in diets to produce signs of toxicity (NRC, 1987).
Humans are reported to tolerate daily doses of 20 to 1,000 mg per
day of pyridoxine-HCl for prolonged periods without deleterious effects. These doses are 10 to 500 times greater than the recommended
daily dietary allowance of vitamin B6 for adults (Haskell, 1978). Vitamin B6 is one of several vitamins currently popular in megadose therapy(see Supplementation). Humans have developed a sensory neuropathy as412

V I T A M I N B6

a direct result of taking 2,000 to 6,000 mg of pyridoxine daily (Schaumburg et al., 1983). Subjects developed ataxia and severe sensory nervous
system dysfunctions, but substantial improvement occurred with pyridoxine withdrawal.

■ REFERENCES
Adams, C.R. (1982). In Vitamins—The Life Essentials. Nutrition Institute, National Feed Ingredient Association, NI-82, 1, Des Moines, Iowa.
Adams, C.R., Richardson, C.E., and Cunha, T.J. (1967). J. Anim. Sci. 26, 903(Abstr.).
Antopol, W., and Tarlov, I.M. (1992). J. Neuropathol. Exp. Neurol. 1, 330.
Asmar, J.A., Daghirn, N.J., and Azar, H.A. (1968). J. Nutr. 95, 153.
Baker, H., Marcus, S.L., Petrylak, D.P., Frank, O., DeAngelis, B., Baker, E.R.,
Dutcher, J.P., and Wiernik, P.H. (1992). J. Am. College Nutr. 11, 482.
Basu, T.K., and Mann, S. (1997). J. Nutr. 127, 117.
Bauernfeind, J.C. (1974). Feedstuffs, 46, 30.
Beilstein, M.A., and Whanger, P.D. (1989). J. Nutr. 119, 1962.
Birdsall, J.J. (1975). Technology of Fortification of Foods, p. 19. National Academy of Sciences, Washington, D.C.
Blalock, T.L., and Thaxton, J.P. (1984). Poult. Sci. 63, 1243.
Blalock, T.L., Thaxton, J.P., and Garlich, J.D. (1984). J. Nutr. 114, 312.
Bonjour, J.P. (1980). Int. J. Vitam. Nutr. Res. 50, 215.
Bräunlich, K. (1974). Vitamin B6, No. 1451. Hoffmann-La Roche, Basel,
Switzerland.
Buckmaster, P.S., Holliday, T.A., Bai, S.C., and Rogers, Q.R. (1993). J. Nutr.
123, 20.
Brin, M. (1971). Am. J. Clin. Nutr. 24, 704.
Cho, Y.O., and LeKlem, J.E. (1990). J. Nutr. 120, 258.
Coburn, S.P., Mahuren, J.D., Pauly, T.A., Ericson, K.L., and Townsend, D.W.
(1992). J. Nutr. 122, 2348.
Contractor, S., and Shane, B. (1970). Am. J. Obstet. Gynecol. 107, 635.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Daghir, N.J., and Haddad, K.S. (1981). Poult. Sci. 60, 988.
Delport, R., Ubbink, J.B., Vermoak, W.J.H., Shaw, A., Bissbort, S., and Becker,
P.J. (1991). Nutr. Burbank 7, 260.
Dobbelstein, H., Korner, W., Mempel, W., Grosse-Wilde, H., and Edel, H.
(1974). Kidney Int. 5, 233.
Driskell, J.A. (1994). Nutr. Res. 14, 293.
Dubeski, P.L., and Owens, F.N. (1993). In Animal Science Res. Rept. No. P-933.
Stillwater, Oklahoma.
Durst, L., Kirchgessner, M., and Roth-Maier, D.A. (1989). J. Anim. Physiol.
Anim. Nutr. 62, 85.
Ekanayake, A., and Nelson, P.E. (1990). J. Food Sci. 55, 154.
Fisher, J.H., Willis, R.A., and Haskell, B.E. (1984). J. Nutr. 114, 786.
Folkers, K., Shizukuishi, S., Scudder, S.L., Willis, R., Takemura, K., and Longe413

CHAPTER NINE

necker, J.B. (1981). Biochem. Biophys. Res. Commun. 100, 972.
Fuller, H.L., and Kifer, P.E. (1959). Poult. Sci. 38, 255.
Fuller, H.F., Field, R.C., Roncalli-Amici, A., Dunahoo, W.S., and Edwards, H.M.
(1961). Poult. Sci. 40, 249.
Gadient, M. (1986). Proc. Nutr. Conf. Feed Manuf. College Park, Maryland, p.
73.
Gilbert, J.A., and Gregory, J.F. III. (1992). J. Nutr. 122, 1029.
Gorzalka, B.B., and Moe, I.V. (1994). Nutr. Res. 14, 279.
Gregory, J.F. III. (1988). J. Food Comp. Anal. 1, 105.
Gregory, J.F. III, Trumbo, P.R., Bailey, L.B., Toth, J.P., Baumgartner, T.G., andCerda, J.J. (1991). J. Nutr. 121, 177.
Gries, C.L., and Scott, M.L. (1972). J. Nutr. 102, 1269.
Groziak, S.M., and Kirksey, A. (1987). J. Nutr. 117, 1045.
Groziak, S.M., and Kirksey, A. (1990). J. Nutr. 120, 485.
Guilarte, T.R. (1993). Nutr. Rev. 51, 193.
Gummow, B., Bastianello, S.S., Labuschagne, L., and Erasmus, G.L. (1992). Onderstepoort J. Vet. Res. 59, 111.
Hansen, C.M., LeKlem, J.E., and Miller, L.T. (1996). J. Nutr. 126, 2512.
Harmon, B.G., Miller, E.R., Hoefer, J.A., Ullrey, D.E., and Leucke, R.W. (1963).
J. Nutr. 79, 263.
Haskell, B.E. (1978). In Handbook Series in Nutrition and Food, Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 1, p. 43. CRC Press, Boca Raton, Florida.
Heller, S., Salkeld, R.M., and Korner, W.F. (1973). Am. J. Clin. Nutr. 26, 1330.
Heseker, H., and Kübler, W. (1992). In Nutrition of the Elderly (H.N. Munro
and G.J. Schlierf, eds.), Nutrition Workshop Series 29. Raven Press, NewYork.
Hoffmann-La Roche. (1979). Rationale for Roche Recommended Vitamin Fortification-Poultry Rations. RCD 5692/979. Hoffmann-La Roche, Nutley,
New Jersey.
Hoover-Plow, J., and Rice, S. (1992). Nutr. Res. 12, 773.
Ink, S.L., and Henderson, L.M. (1984). Annu. Rev. Nutr. 4, 455.
Johnson, B.C., Mitchell, H.H., Hamilton, T.S., and Nevens, W.B. (1947). Fed.
Proc. Am. Soc. Exp. Biol. 6, 410.
Johnson, B.C., Pinkos, J.A., and Burke, K.A. (1950). J. Nutr. 40, 309.
Kabir, H., LeKlem, J.E., and Miller, L.T. (1983). J. Nutr. 113, 2412.
Kafatos, A., Diacatou, A., Labadarios, D., Kounali, D., Apostolaki, J., Vlachonikolis, J., Mamalakis, G., and Megremis, S. (1993). J. Am. Coll. Nutr.
12, 685.
Kaido, M., Mori, K., Ide, Y., Inoue, N., and Koide, O. (1991). Exp. Mol. Pathol.
55, 63.
Kang-Yoon, S.A., Kirksey, A., Giacoia, G., and West, K. (1992). Am. J. Clin.
Nutr. 56, 548.
Kirksey, A. (1981). In Vitamin B6 Methodology and Nutrition Assessment (J.E.
LeKlem and R.D. Reynolds, eds.), p. 269. Plenum, New York.
Kodentsova, V.M., Yakushina, L.M., Vrzhesinskaya, O.A., Beketova, N.A., andSpirichev, V.B. (1993). Voprosy Pitaniya 5, 32.
Korede, O. (1990). Ann. Nutr. Metab. 34, 273.
414

V I T A M I N B6

Krinke, G.J., and Fitzgerald, R.E. (1988). Toxicology 49, 171.
Lakshmi, R., Lakshmi, A.V., Divan, P.V., and Bamji, M.S. (1991). Indian J.
Biochem. Biophys. 28, 481.
Leeson, S., and Marcotte, M. (1993). World’s Poultry Sci. J. 49, 120.
LeKlem, J.E. (1990). J. Nutr. 120, 1503.
LeKlem, J.E. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 341. Marcel Dekker, New York.
LeKlem, J.E., Miller, L.T., Perera, A.D., and Peffers, D.E. (1980). J. Nutr. 110,1819.
Lilum, D., Hammond, W., Krauss, D., and Schoonmaker, J. (1981). Invest. Urol.
18, 451.
Lin, G.W.J. (1989). Alcohol. Clin. Exp. Res. 13, 236.
Loosli, J.K. (1991). In Animal Science Handbook (P.A. Putnam, ed.). Academic
Press, San Diego, California.
Lovell, R.T. (1987). Feed Manage. 38, 28.
Löwik, M.R.H., van den Berg, H., Westenbrink, S., Wedel, M., Schrijver, J., andOckhuizen, T. (1989). Am. J. Clin. Nutr. 50, 391.
Lucas, H.L., Heuser, G.F., and Norris, L.C. (1946). Poult. Sci. 25, 137.
Lui, A., Lumeng, L., and Li, T. (1983). J. Nutr. 113, 893.
Lui, A., Lumeng, L., Aronoff, G.R., and Li, T. (1985). J. Lab. Clin. Med. 106,
491.
Mahuren, J.D., Pauly, T.A., and Coburn, S.P. (1991). J. Nutr. Biochem. 2, 449.
Marks, J. (1975). A Guide to the Vitamins, Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
Masse, P.G., Pritzker, K.P., Mendes, M.G., Boskey, A.L., and Weiser, H. (1994).
Brit. J. Nutr. 71, 919.
Matyaszczyk, M., Karczmarewicz, E., Czarnowska, E., Reynolds, R.D., andLorenc, R.S. (1993). J. Nutr. 123, 204.
McCullough, A.L., Kirksey, A., and Wacks, T.C. (1990). Am. J. Clin. Nutr. 51,
1067.
McCollum, E.V. (1957). In A History of Nutrition. The Riverside Press, Cambridge, Massachusetts.
McElroy, L.W., and Goss, H. (1939). J. Biol. Chem. 130, 437.
Mehansho, H., and Henderson, L.M. (1980). J. Biol. Chem. 255, 11901.
Mehansho, H., Hamm, M.W., and Henderson, L.M. (1979). J. Nutr. 109, 1542.
Merrill, A.H., and Burnhan, F.S. (1990). In Nutrition Reviews, Present Knowledge in Nutrition (M.L. Brown, ed.), p. 155. The Nutrition Foundation,
Washington, D.C.
Meydani, S.N., Ribaya-Mercado, J.D., Russell, R.M., Sahyoun, N., Morrow,
F.D., and Gershoff, S.N. (1991). Am. J. Clin. Nutr. 53, 1275.
Miller, B.F. (1963). Poult. Sci. 42, 795.
Miller, E.E., and Baumann, C.A. (1945). J. Biol. Chem. 157, 551.
Miller, L.T. (1986). J. Nutr. 116, 1344.
Mitwalli, A. (1989). Ann. Saudi Med. 6, 541.
Nakano, H., and Gregory, J.F. III. (1995). J. Nutr. 125, 926.
Nakano, H., McMahon, L.G., and Gregory, J.F. III. (1997). J. Nutr. 127, 1508.
Nguyen, L.B., and Gregory, J.F. III. (1983). J. Nutr. 113, 1550.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sci415

CHAPTER NINE

ences- National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Oka, T., Komori, N., Kuwahata, M., Suzuki, I., Okada, M., and Natori, Y.
(1994). Experientia 50, 127.
Pannemans, D.L., van den Berg, H., and Westerterp, K.R. (1994). J. Nutr. 124,
1207.
Parsons, J.L., and Klostermann, H.J. (1967). Feedstuffs 39(45), 74.
Perry, S.C. (1978). Proc. Roche Vitam. Nutr. Update Meeting, p. 29. Hot
Springs, Arkansas.
Pingali, A.V., and Trumbo, P.R. (1992). J. Agr. Food. Chem. 40, 1860.
Price, J.M., Brown, R.R., and Larson, F.C. (1957). J. Clin. Invest. 36, 1600.
Rabinowitz, J.C., and Snell, E.E. (1949). Proc. Soc. Exp. Biol. Med. 70, 235.
Rall, L.C., and Meydani, S.N. (1993). Nutr. Rev. 51, 217.
RDA. (1989). Recommended Dietary Allowances, 10th Ed. National Academyof Sciences-National Research Council, Washington, D.C.
Ritchie, H.D., Miller, E.R., Ullrey, D.E., Hofer, J.A., and Lucke, R.W. (1960). J.
Nutr. 70, 491.
Robel, E.J. (1992). Poultry Sci. 71, 1733.
Robel, E.J., and Christensen, V.L. (1991). Brit. Poult. Sci. 32, 509.
Roe, D.A. (1973). Drug Ther. 3, 23.
Roth-Maier, D.A., and Kirchgessner, M. (1993). J. Anim. Physiol. Anim. Nutr.
70, 6.
Rothschild, B. (1982). Arch. Intern. Med. 142, 840 (Abstr.).
Sakurai, T., Asakura, T., Mizuno, A., and Matsuda, M. (1992). J. Nutr. Sci. Vitaminol. 38, 227.
Salkeld, R., Knorr, K., and Korner, W. (1973). Clin. Chim. Acta 49, 195.
Schaeffer, M.C. (1993). J. Nutr. 123, 1444.
Schaeffer, M.C., Cochary, E.F., and Sadowski, J.A. (1990). J. Am. Coll. Nutr. 9,120.
Schaumburg, H., Kaplan, J., Windebank, A., Vick, N., Rasmug, S., Pleasure, D.,
and Brown, M.J. (1983). N. Engl. J. Med. 309, 445.
416

V I T A M I N B6

Schenker, S., Johnson, R.F., Mahuren, J.D., Henderson, G.I., and Coburn, S.P.
(1992). Am. J. Physiol. 262, R966.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Scriver, C.R., and Whelan, D.T. (1969). Ann. N Y Acad. Sci. 166, 83.
Shideler, C.E. (1983). Am. J. Med. Technol. 49, 17.
Snell, E.E. (1986). In Pyridoxal Phosphate: Chemical, Biochemical and Medical
Aspects, Part A (D. Dolphin, R. Poulson and O. Avramovic, eds.), Vol. 1A,
p. 1. John Wiley and Sons, New York.
Styslinger, L., and Kirksey, A. (1985). Am. J. Clin. Nutr. 41, 21.
Tarr, J.B., Tamura, T., and Stokstad, E.L.R. (1981). Am. J. Clin. Nutr. 34, 1328.
Tomarelli, R.M., Spence, E.R., and Bernhart, F.W. (1955). J. Agric. Food Chem.
3, 338.
Trumbo, P.R., and Gregory, J.F. III. (1989). J. Nutr. 119, 36.
Trumbo, P.R., and Raidi, M.A. (1991). Nutr. Res. 11, 53.
Trumbo, P.R., Gregory, J.F. III, and Sartain, D.B. (1988). J. Nutr. 118, 170.
Unna, K., and Antopol, W. (1940). Proc. Soc. Exp. Biol. Med. 43, 116.
Vanderslice, J.T., Maire, C.E., Doherty, R.F., and Beecher, G.R. (1980). J. Agric.
Food Chem. 28, 1145.
Van Schoonhoven, J., Schrijver, J., van den Berg, H., and Haenen, G. (1994). J.
Agr. Food Chem. 42, 1475.
Verbeeck, J. (1975). Feedstuffs 47(36), 4.
Wada, H., and Snell, E.E. (1961). J. Biol. Chem. 236, 2089.
Wei, I.L., and Young, T.K. (1994). Nutr. Res. 14, 271.
Wetherilt, H., Ackurt, F., Brubacher, G., Okan, B., Aktas, S., and Turdu, S.
(1992). Int. J. Vitam. Nutr. Res. 62, 21.
Witkowska, D., Sedrowicz, L., and Oledzka, R. (1992). Bromatologia Chemia
Toksykoleziczna 25, 25.
Wohl, M., Levy, H., Szutha, A., and Maldia, G. (1960). Proc. Soc. Exp. Biol.
Med. 105, 523.
Yang, C.P., and Jeng, S.L. (1989). J. Chinese Agr. Chem. Soc. 27, 450.
Yen, J.T., Jensen, A.H., and Baker, D.H. (1976). J. Anim. Sci. 42, 866.
Yin, S., Sato, I., and Yamaguchi, K. (1992). J. Nutr. Biochem. 3, 633.
Zempleni, J., Link, G., and Kubler, W. (1992). Int. J. Vitam. Nutr. Res. 62, 165.
Zhang, Z., Gregory, J.F. III, and McCormick, D.B. (1993). J. Nutr. 123, 85.

417

chapter ten

PA N T O T H E N I C A C I D

INTRODUCTION
After the discovery of thiamin, riboflavin, and niacin as B-complex
vitamins, researchers realized that at least one other unidentified factor
remained. Two additional vitamins, pyridoxine (vitamin B6) and pantothenic acid, were found in the late 1930s as fractions of yeast and liver.
Tissue extracts from a variety of biological materials provided a growth
factor for yeast that was identified as pantothenic acid, derived from theGreek word pantos, meaning “found everywhere.”
Pantothenic acid is found in two enzymes, coenzyme A and acyl carrier protein, which are involved in many reactions in carbohydrate, fat,and protein metabolism. Although this vitamin occurs in practically all
feedstuffs, the quantity present is generally insufficient for optimum performance of poultry and swine and other monogastric species. There are
no reports of deficiency in adult ruminants because of microbial synthesis. Pantothenic acid deficiency occurs only rarely in humans.

HISTORY
Historical aspects of pantothenic acid have been reviewed (Sauberlich, 1973; Fox, 1991; Loosli, 1991). During the 1930s several independent research programs concentrated on either a growth factor for
microorganisms or a chick antidermatitis factor. Research to identify
pantothenic acid was also closely associated with studies of vitamin B6(pyridoxine). Both vitamins were fractions of the “vitamin B2” complex
and were found associated in biological materials. Since pyridoxine wasadsorbed on Fuller’s earth, from which it could be eluted, and pantothenic acid was not, pantothenic acid was sometimes referred to as
419

10CHAPTER TEN“filtrate factor” and pyridoxine as “eluate factor.” Pantothenic acid deficiency was first described in the chick as a pellagra-like dermatitis by
Norris and Ringrose in 1930. Since the filtrate factor cured dermatitis inchicks, but not rats, one early name of the vitamin was “chick antidermatitis factor.”
In independent studies in 1933, Williams and associates fractionated “bios,” a growth factor for yeast, particularly Saccharomyces cerevisiae. They concentrated the factor, determined many of its properties,
and provided the name pantothenic acid. In 1937 Snell and associates
independently studied the nature of an essential factor for lactic andpropionic acid bacteria. It was soon found that the filtrate factor, thechick antidermatitis factor, and the unknown factor required by yeast
and lactic acid bacteria were all pantothenic acid. In 1939 Williams isolated pantothenic acid and determined the structure in 1940. Also in
1940 the vitamin was synthesized independently in three different laboratories. The active form of pantothenic acid, coenzyme A, was discovered as an essential cofactor in acetylation of sulfonamides in the liverby Lipmann and Kaplan in 1947.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Pantothenic acid is an amide consisting of pantoic acid (α, γ-dihydroxy-β, β-dimethylbutyric acid) joined to β-alanine. Pantothenic acidand the biologically active coenzyme A, which contains the vitamin as
an essential component, are shown in Fig. 10.1. The vitamin is derivatized at its carboxyl end by β-mercaptoethylamine and at its alcohol end
by phosphate to form a pseudodinucleotide containing phosphoadenylic
acid. Therefore coenzyme A contains the vitamin combined with adenosine 3′-phosphate, pyrophosphate, and β-mercaptoethylamine. Anothermetabolically active form of pantothenic acid, panthenol, is more easily
absorbed and is converted to the acid in vivo.
Analogs of pantothenic acid that have replaced the β-alanine withother amino acids, such as α-alanine, β-aminobutyric acid, aspartic acid,leucine, or lysine, are inactive. The most common antagonist of pantothenic acid is ω-methyl-pantothenic acid, which has been used to produce a deficiency of the vitamin in humans (Hodges et al., 1958). Other
antivitamins include pantoyltaurine, phenylpantothenate hydroxycobalamine (c-lactam) (analog of vitamin B12), and antimetabolites of the vitamin containing alkyl or aryl ureido and carbamate components in theamide part of the molecule (Fox, 1991; Brass, 1993). Feeding chickens
420

PA N T O T H E N I C A C I D

Fig. 10.1 Structures of pantothenic acid and coenzyme A.

on high intakes of copper results in reduced formation of coenzyme A,
by increasing the oxidation of cysteine to cystine, and also by the formation of copper-cysteine and copper-glutathione complexes, which
render the amino acid unavailable for coenzyme A synthesis (Latymer
and Coates, 1981).
The free acid of the vitamin is a viscous, pale yellow oil readily soluble in water and ethyl acetate. The oil is extremely hygroscopic and iseasily destroyed by acids, bases, and heat. Maximum heat stability occurs at pH 5.5 to 7.0. Calcium pantothenate, the form used in commerce, crystallizes as white needles from methanol and is reasonably
421

CHAPTER TEN

stable to light and air. Pantothenic acid is optically active (characteristicof rotating a polarized light). It may be prepared as either the pure dextrorotatory (d-) form or the dl-form; the racemic form has approximately one-half the biological activity of d-calcium pantothenate. Only
the dextrorotatory form, d-pantothenic acid, is effective as a vitamin.
The l-pantothenic acid is inactive for organisms requiring the intact vitamin.

ANALYTICAL PROCEDURES
Pantothenic acid content of various substances has been analyzed by
microbiological assay, animal bioassay, radioimmunoassay, and chemical methods (Fox, 1991). The complex nature of natural products has
hindered development of chemical methods for assay of the vitamin.
Chick growth bioassays have been used extensively and measure boundforms as well as free pantothenic acid.
The rat and chick tests have gradually been replaced by more rapid,
simpler, and cheaper microbiological tests. Microbiological proceduresfor determination of pantothenic acid are widely employed but requirethat the vitamin be freed from the coenzyme form. Values for pantothenic acid content of feedstuffs obtained by microbiological procedures performed before proper methods were devised for liberating thevitamin from its bound form are much too low. Procedures for cleavageof bound forms to obtain free pantothenic acid can include use of (1) intestinal phosphatase to cleave the phosphate linkage and (2) pigeon orchick liver enzyme preparation to break the linkage between mercaptoethylamine and pantothenic acid. Bound forms cannot be hydrolyzed
as this will destroy the vitamin. A number of organisms can be used in
the assay method; Lactobacillus plantarum (formerly Lactobacillus arabinosus) is widely used.
Radioimmunoassay and an automated fluorometric assay technique(Roy and Buccafuri, 1978) have been reported to be successful for pantothenic acid determination. Pantothenic acid levels in blood have been
determined by radioimmunoassay (Wyse and Hansen, 1977). After
treatment with alkaline phosphatase and liver extract, the pantothenic
acid content of whole blood analyzed by radioimmunoassay comparedfavorably to the microbiological assay with L. plantarum. An indirect
enzyme-linked immunosorbent assay (ELISA) has been developed for
determining pantothenic acid in plasma (Song et al., 1990). The ELISA
was based on the competition for the antibody between immobilized422

PA N T O T H E N I C A C I D

pantothenate adsorbed on polystyrene microwells and free pantothenate
in samples. This method is easy to use, less expensive, and sensitive.

METABOLISM
Pantothenic acid is found in feeds in both bound and free forms.
The bound coenzyme forms are principally coenzyme A and acyl carrier
protein (ACP). It is necessary to liberate the pantothenic acid from thebound forms in the digestive process prior to absorption. Coenzyme A
and other bound forms are hydrolyzed in the intestinal lumen to 4′phosphopantetheine. This form is dephosphorylated to yield pantothenate, which is rapidly converted by the intestinal enzyme pantetheinase topantothenic acid. Pantothenic acid, its salt, and the alcohol are absorbed
primarily in the jejunum by a specific transport system that is saturable
and sodium ion dependent (Fenstermacher and Rose, 1986). The alcohol form, panthenol, which is oxidized to pantothenic acid in vivo, appears to be absorbed somewhat faster than the acid form. After absorption, pantothenic acid is transported to various tissues in the plasma,from which it is taken up by most cells via another active-transport
process involving cotransport of pantothenate and sodium in a 1:1 ratio(Olson, 1990).
Within all tissues pantothenic acid is converted to coenzyme A andother compounds in which the vitamin is a functional group (Sauberlich,1985). Free pantothenate appears to be efficiently absorbed; in the dog,
between 81 and 94% of an oral dose of sodium [14C] pantothenate wasabsorbed (Taylor et al., 1974). Measurement of pantothenic acidbioavailability in adult men consuming a typical United States dietranged from 40 to 61%, with an average of 50% (Sauberlich, 1985).
Urinary excretion represents the major route of body loss of absorbed
pantothenic acid, with prompt excretion when taken in excess. Most
pantothenic acid is excreted as the free vitamin, but some species (e.g.,dog) excrete it as β-glucuronide (Taylor et al., 1972). An appreciable
quantity of pantothenic acid (approximately 15% of daily intake) is oxidized completely and is excreted across the lungs as CO2 (Combs,
1992).
Animals and humans do not appear to have the ability to store appreciable amounts of pantothenic acid, with organs such as the liver andkidney having the highest concentrations. Most pantothenic acid inblood exists in red blood cells as coenzyme A; serum contains no coenzyme A but does contain free pantothenic acid. A number of human423

CHAPTER TEN

studies suggest that stores of pantothenic acid are not great, as tissue levels dropped dramatically and/or clinical signs developed after 7 to 12
weeks on diets low in the vitamin (Fry et al., 1976). Urinary pantothenic
acid concentrations dropped from 3 to 8.0 mg in just 63 days in humansubjects fed low quantities of the vitamin (Fry et al., 1976). Coenzyme
A is synthesized by cells from pantothenic acid, ATP, and cysteine. Pantothenic acid kinase, a cytosolic enzyme, is rate limiting for the overall
pathway of coenzyme A biosynthesis (Brass, 1993).

FUNCTIONS
Pantothenic acid functions as a constituent of two important coenzymes—coenzyme A and ACP. Coenzyme A is found in all tissues and is
one of the most important coenzymes for tissue metabolism. The important role of coenzyme A is summarized in Fig. 10.2; biochemical reactions catalyzed by coenzyme A are presented in Table 10.1. The coenzymes are known to be involved in more than 100 different metabolic
pathways involving the metabolism of carbohydrates, proteins, andlipids, and the synthesis of lipids, neurotransmitters, steroid hormones,
porphyrins, and hemoglobin.
The most important function of coenzyme A is to act as a carrier
mechanism for carboxylic acids (Marks, 1975). Such acids, when boundto coenzyme A, have a high potential for transfer to other groups andare normally then referred to as “active.” The most important of thesereactions is the combination of coenzyme A with acetate to form “active
acetate” with a high-energy bond that renders acetate capable of further
chemical interactions. As an example, it is utilized directly by combination with oxaloacetic acid to form citric acid, which enters the Krebs citric acid cycle. Its combination with two-carbon fragments from fats, carbohydrates, and certain amino acids to form acetyl coenzyme A is an
essential step in their complete metabolism, because the coenzyme enables these fragments to enter the citric acid cycle.
Coenzyme A functions as a carrier of acyl groups in enzymatic reactions involved in synthesis of fatty acids, cholesterol, and sterols; inthe oxidation of fatty acids, pyruvate, and α-ketoglutarate; and in biological acetylations. In the form of active acetate, acetic acid can also
combine with choline to form acetylcholine, the chemical transmitter at
the nerve synapse, and can be used for detoxification of drugs including
sulfonamides.
Coenzyme A has an essential function in lipid metabolism, fatty
acids are activated by formation of the coenzyme derivative, and degra424

PA N T O T H E N I C A C I D

Fig. 10.2 The important role of coenzyme A in metabolism. (Modified fromMarks, 1975.)

dation by removal of acetate fragments in beta oxidation also uses another molecule of coenzyme A. These active acetate fragments may directly enter the citric acid cycle or combine to form ketone bodies. A
pantothenic acid-dependent enzyme, ACP, that replaces coenzyme A
during building of the carbon chain in synthesis of fatty acids, was recognized in 1965 (Pugh and Wakil, 1965). ACP is a protein with asulfhydryl group covalently attached to acetyl, malonyl, and intermediate-chain acyl groups. The sulfhydryl group at the binding site of the
ACP was identified as a cysteine residue, thioethanolamine. Discovery ofthioethanolamine and β-alanine residues suggested that both ACP andcoenzyme A have similar acyl-binding sites.
425

CHAPTER TEN■ Table 10.1

Selected Biochemical Reactions Catalyzed by Coenzyme A
Pantothenate
Derivative

Reactant

Product

Site

Pyruvic
dehydrogenaseCoA

Pyruvate

Acetyl-CoA

Mitochondria

α-Ketoglutarate
dehydrogenaseCoA

α-Ketoglutarate

Succinyl-CoA

Mitochondria

Fatty acid oxidase CoA

Palmitate

Acetyl-CoA

Mitochondria

Fatty acidsynthetase

Acyl carrier protein

Acetyl-CoA,
Malonyl-CoA

Palmitate

Microsomes

Propionyl-CoA
carboxylase

CoA

Propionyl-CoA,carbon dioxide

MethylmalonylCoA

Microsomes

Acyl-CoAsynthetase

Phosphopanthetheine Succinyl-CoA,
GDP + P1

Succinate,
GTP + CoA

Mitochondria

EnzymeSource: Modified from Olson (1990).

Decarboxylation of ketoglutaric acid in the citric acid cycle yields
succinic acid, which is then converted to the active form by linkage withcoenzyme A. Active succinate and glycine are together involved in thefirst step of heme biosynthesis. Pantothenic acid also stimulates synthesis of antibodies, which increase resistance of animals to pathogens. It
appears that when pantothenic acid is deficient, the incorporation ofamino acids into the blood albumin fraction is inhibited, which would
explain why there is a reduction in the titer of antibodies (Axelrod,
1971). The 4′-phosphopantetheine is the prosthetic group of an enzyme
system that synthesizes peptide antibodies, such as gramicidin in bacteria (Olson, 1990).

REQUIREMENTS
Pantothenic acid requirements for selected animals and humans arepresented in Table 10.2. For growth and reproduction, the majority of
species have a dietary requirement between 5 and 15 mg/kg of diet. Foregg production in chickens, the pantothenic acid requirement is very low(2.2 mg/kg) compared to a requirement of 10 mg/kg for growth and reproduction. Requirements are based on typical consumption levels.
When energy density of diets is increased, intake is reduced, so higher
dietary concentrations of pantothenic acid and other vitamins are required. When the level of energy in the rations of broilers was raised426

PA N T O T H E N I C A C I D■ Table 10.2

Pantothenic Acid Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Leghorn, 0–18 weeks
Laying (100-g intake)
Broilers, 0–8 weeks
0–7 weeks
All classes
0–4 weeks
4–24 weeks
Adult
Growing-finishing, 3–5 kg
Growing-finishing, 5–110 kg
Adult
Adult
Adult
Growing
Growing
GrowingCatfishPacific salmon
Rainbow trout
All classes
All classes
Infants
AdultsMicrobial synthesis
13 ppm milk replacer
Microbial synthesis
10.0 mg/kg
1.7–2.5 mg/kg
10.0 mg/kg
11.0 mg/kg
10.0–15.0 mg/kg
10.0 mg/kg
9.0 mg/kg
Microbial synthesis
12.0 mg/kg
7.0–10.0 mg/kg
12.0 mg/kg
Microbial synthesis
Microbial synthesis
8.0 mg/kg
5.0 mg/kg
400 μg/kg body wt.
10–15 mg/kg
17–50 mg/kg
10–20 mg/kg
8.0 mg/kg
16.0 mg/kg
2–3 mg/day
4–7 mg/day

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1981b)
NRC (1982a)
NRC (1986)
NRC (1985a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)

ChickenDuck (Pekin)Japanese quail
Turkey
Sheep
Swine
Horse
Goat
Fox
Cat
Dog
Fish
Rat
Mouse
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as mg/day.

from 2,870 to 3,505 kcal/kg, intake of pantothenic acid fell by 19.1%
because appetite is mainly controlled by intake of energy (Friesecke,1975). For the blue tilapia (Tilapia aurea) fish, the growth requirement
for pantothenic acid was 6 mg/kg, but based on survival and lack ofpathology, a value of 10 mg/kg was recommended (Roem et al., 1991).
Research has shown a wide variation in pantothenic acid requirements, even among animals within a breed. Data from Michigan showthat for one-half of growing pigs, 9.13 mg/kg of pantothenic acid was
sufficient for growth, whereas the remaining one-half required morethan 9.13 but less than 13.5 mg/kg (Luecke et al., 1953).
Antibiotics have a sparing effect on the pantothenic acid requirement. A dietary level of 22 mg/kg aureomycin for weanling pigs (McKigney et al., 1957) and 10 mg/kg of procaine penicillin to turkey poults(Slinger and Pepper, 1954) reduced the pantothenic acid requirement forthese species.
427

CHAPTER TEN

If the rumen is functioning normally, ruminal microflora will synthesize enough pantothenic acid to satisfy ruminant needs; however,biosynthesis will depend on the composition of feeds. Vitamin synthesis
is reduced with diets high in cellulose but increases with higher quantities of easily soluble carbohydrates (Virtanen, 1966). Certain amountsof B-complex (including pantothenic acid) vitamins are synthesized in
the large intestine of monogastric animals. It is doubtful, however,whether much benefit is derived, as only limited pantothenic acid absorption occurs in the large intestine, with greatest benefit found in animals that practice coprophagy (Friesecke, 1975).
Interrelationships with other vitamins (e.g., vitamin B12, vitamin C,
and biotin) on pantothenic acid requirements are known (Scott et al.,1982). Pantothenic acid requirement of chicks from B12-depleted hens
was found to be greater than that of chicks from normal hens. A fivefold increase in coenzyme A content of liver was found in B12-deficient
chicks and rats. There is no evidence of sparing action of B12 on pantothenic acid needs for swine as has been observed in poultry (Luecke etal., 1953). There is some evidence that pantothenic acid is involved inascorbic acid synthesis in plants and animals, with ascorbic acid sparing
the pantothenic acid requirement in rats (Barboriak and Krehl, 1957).
Also, there have been suggestions of a possible interrelationship between
folacin and biotin with pantothenate. Both vitamins were found necessary for pantothenic acid utilization in the rat (Wright and Welch,1943). The inclusion of biotin in the diet of a pantothenic acid-deficient
pig was effective in prolonging the life of the pig, but caused the pantothenic acid deficiency signs to appear in half the time (Colby et al.,1948).
Dietary fat and protein influence the pantothenic acid requirement.
Pigs fed a diet deficient in pantothenic acid and high in fat failed to gain
weight, exhibited lower feed efficiency, and developed deficiency signs
more quickly than those fed diets low in fat (Sewell et al., 1962). Nelson
and Evans (1945) found that pantothenic acid-deficient rats fed a highprotein diet excreted more pantothenic acid and had accelerated growthand survival rates compared with rats fed a low-protein diet. The superiority of the high-protein diet may be due to the decreased level of dietary carbohydrate, which would presumably require coenzyme A for
metabolism.
The Food and Nutrition Board of the National Research Council(RDA, 1989) has not established a recommended human dietary allowance for pantothenic acid but states that 4 to 7 mg per day for adults
and 2 to 3 mg per day for infants are safe and adequate intakes. Women428

PA N T O T H E N I C A C I D

who are pregnant or lactating may have an increased requirement of
about 30% (Plaut et al., 1974).

NATURAL SOURCES
Pantothenic acid is widely distributed in foods of animal and plantorigin. Alfalfa hay, peanut meal, cane molasses, yeast, rice bran and polishings, green leafy plants, wheat bran, brewer’s yeast, and fish solubles
are good sources for animals. For human foods, in general, liver, kidney,
yeast, egg yolk, and fresh vegetables (e.g., mushrooms, avocados, andbroccoli) are the best sources, followed by milk and meat; grains, fruits,
and nuts contain the lowest levels. Royal jelly of bees is a rich source (510ppm), with ovaries of the cod containing more than 2,000 ppm. Analysis of feeds for pantothenic acid are sometimes conflicting depending onwhether data were determined by microbiological assays or more recent
radio immunoassay procedures. Different methods of sample hydrolysis
may also contribute to the variation (see Analytical Procedures). The
pantothenic acid content of foods and feedstuffs is shown in Table 10.3.
Many swine and poultry diets are borderline in supplying pantothenic acid requirements, and many are deficient in this vitamin. Dietsbased on corn and soybean meal are apt to be deficient in pantothenic
acid. Milling by-products such as rice bran and wheat bran are good

■ Table 10.3

Pantothenic Acid in Foods and Feedstuffs (mg/kg, dry basis)

Alfalfa hay, sun cured
28.6
Alfalfa leaves, sun cured
32.4
Barley, grain
9.1
Bean, navy (seed)
2.3
Blood meal
2.6
Brewer's grains
8.9
Buttermilk (cattle)
40.1
Citrus pulp
14.3
Clover hay, ladino (sun cured)
1.1
Copra meal (coconut)
6.9
Corn, gluten meal
11.2
Corn, yellow grain
6.6
Cottonseed meal, solvent extracted 15.4
Eggs
27.0
Fish meal, anchovy
10.9
Fish meal, menhaden
9.4
Fish, sardine
11.8
Linseed meal, solvent extracted
16.3
Liver, cattle
164.9
Milk, skim, cattle
38.6

Molasses, sugarcane
Oat, grain
Pea seeds
Peanut meal, solvent extracted
PotatoRice, branRice, grain
Rice, polished
Rye, grain
Sorghum, grain
Soybean meal, solvent extracted
Soybean seedSpleen, cattle
Timothy hay, sun cured
Wheat, bran
Wheat, grain
Whey
Yeast, brewer's
Yeast, torula

50.3
8.8
21.0
50.7
22.0
25.2
9.1
3.9
9.1
12.5
18.2
17.3
8.2
7.9
33.5
11.4
49.6
118.4
100.6

Source: NRC (1982b).

429

CHAPTER TEN

sources, being two to three times higher than the respective grains. Biological availability of pantothenic acid is high from corn and soybean
meal, but low from barley, wheat, and sorghum (Southern and Baker,
1981). Changing processing methods can greatly alter vitamin feed levels. As an example, with changes in sugar technology, literature values
for pantothenic acid content of beet molasses have decreased from 50 to
110 mg/kg in the 1950s to about 1 to 4 mg/kg (Palagina et al., 1990).
Pantothenic acid is reported to be fairly stable in feedstuffs during
long periods of storage (Scott et al., 1982). The authors indicate thatheating during processing may cause considerable losses, especially iftemperatures attain 100 to 150oC for long periods and pH values areabove 7 or below 5.
Gadient (1986) considers pantothenic acid to be slightly sensitive toheat, oxygen, or light, but very sensitive to moisture. Pelleting was reported to cause only small losses of the vitamin. As a general guideline,
pantothenic acid activity in normal pelleted feed over a period of 3
months at room temperature should be 80 to 100%. Pantothenic acidloss during processing of human foods is significant and can amount toas much as 70% in frozen meat and 80% in canned legumes. Loss of thevitamin in dairy products during processing and storage is about 30 to
35% (Song, 1990).

DEFICIENCY
Effects of Deficiency
On the basis of observations on pantothenic acid-deficient animals
and studies in human volunteers, deficiency of the vitamin is shown in
the following signs and symptoms:
1. Reduced growth and feed conversion efficiency.
2. Lesions of skin and its appendages.
3. Disorders of the nervous system.
4. Gastrointestinal disturbances.
5. Inhibition of antibody formation and thus decreased resistance to infection.
6. Impairment of adrenal function.

Clinical signs of pantothenic acid deficiency take many forms anddiffer from one animal species to another. In humans, additional emotional and neurological symptoms include hyperventilation, irritability,430

PA N T O T H E N I C A C I D

insomnia, depression, headache, and dizziness. Since pantothenic acid is
so widely distributed in nature, clear-cut deficiency symptoms in humans are rarely found in practice. However, pantothenic acid does occur in animals with certain feeding regimens.

Ruminants
Pantothenic acid is not required in the diet of adult ruminants, because ruminal microorganisms synthesize this vitamin in adequate
amounts. Sheppard and Johnson (1957) experimentally produced pantothenic acid deficiencies in calves. Major clinical signs were anorexia,
reduced growth, rough hair coat, dermatitis, diarrhea, and eventual
death. The most characteristic pantothenic acid deficiency sign in thecalf is scaly dermatitis around the eyes and muzzle. Anorexia and diarrhea followed after 11 to 20 weeks on a deficient diet, and calves become weak and unable to stand and may develop convulsions. They aresusceptible to mucosal infection, especially in the respiratory tract. Postmortem studies have revealed moderate sciatic and peripheral nerve demyelination. If deficient calves are treated with calcium pantothenate,
they respond with increased appetite and weight gains and subsequent
reversal of dermatitis.

Swine
Many swine diets are borderline in supplying pantothenic acid, andmany are deficient in the vitamin (Cunha, 1977). Nonspecific signs ofpantothenic acid deficiency in swine include reduced growth rate,
bloody diarrhea, loss of appetite, poor general condition, dermatosis,
lowered blood pantothenic acid, and reduced feed conversion.
A characteristic sign of a pantothenic acid deficiency in the pig is locomotor disorder (especially of hindquarters), which has been described
by Goodwin (1962). In the early stages of such a deficiency, the movement of back legs becomes stiff and jerky. Standing animals show aslight tremor of the hindquarters. When deficiency persists, this particular action of the back legs grows more exaggerated and resembles acharacteristic military gait, or goose step (Fig. 10.3) (Luecke et al.,1953). The condition may be so severe that, as the pig moves forward,
the back legs will touch the belly. Finally, increasingly severe paralysis of
the hindquarters develops. Affected pigs frequently fall sideways or, with
their back legs spread apart, assume a posture that resembles that of asitting dog (Fig. 10.4). The chief microscopic lesion is a chromatolysis of
isolated cells of the dorsal root ganglia, followed by a demyelinatingprocess in brachial and sciatic nerves.
431

CHAPTER TEN

Fig. 10.3 Goose-stepping pig with pantothenic acid deficiency. (Courtesy ofR.W. Luecke, Michigan Agriculture Experiment Station.)

Pigs with pantothenic acid deficiency have scurfy skin, thin hair, andbrownish secretion around the eyes. The dermatosis associated with deficiency appears principally on the shoulders and behind the ears; theskin appears dirty and scaly. Skin becomes reddened, and the bristles on
the rump and along the spine loosen and fall. The dermatosis extends tothe mucosa, where it becomes manifest as necrotic enteritis, ulceration,and hemorrhages in the large intestine (Ullrey et al., 1955). As a consequence, the feces contain blood. Goodwin (1962) observed various degrees of gastritis and, occasionally, peritonitis and intestinal fissures.
Ullrey et al. (1955) observed pathological changes in other organsof sows and baby pigs maintained on pantothenic acid-deficient diets.
These changes included fatty liver degeneration, enlargement of adrenals, enlargement of heart (with some related flaccidity of the myocardium), and intramuscular hemorrhages. Histopathological studies
showed degenerative changes and necroses of the tissue cells.
Pantothenic acid is particularly important in sow fertility, with insufficient quantities of the vitamin resulting in complete reproductive
failure. Estrus may occur, but the sows fail to retain the embryos following breeding. Female hogs fed low-pantothenic acid diets exhibited
fatty livers, enlarged adrenal glands, intramuscular hemorrhage, heart
dilation, small ovaries, and improper uterus development (Ullrey et al.,1955). Ensminger et al. (1951) reported that although gilts on a lowpantothenic acid diet became pregnant, they did not farrow or show any
signs of pregnancy. Necropsy of gilts revealed macerating fetuses in the432PA N T O T H E N I C A C I D

ABFig. 10.4 Pantothenic acid deficiency. (A) Locomotor incoordination (goosestepping). (B) An affected pig often falls sideways or, with its back legs spreadapart, assumes a posture that resembles that of a sitting dog. (Courtesy of L.R.
McDowell, University of Florida.)

uterine horns in all cases. Minimal pantothenic acid sufficient to resultin normal farrowing may still result in an abnormal locomotion in suckling pigs from sows that had received diets low in the vitamin (Teague etal., 1971). Sucking movements in the deficient piglets are impaired, as is
the use of the tongue.

Poultry
The major lesions of pantothenic acid deficiency for poultry appear to involve the nervous system, adrenal cortex, and skin (Scott et433

CHAPTER TEN

Fig. 10.5 Pantothenic
acid deficiency in achick, with dermatitis
around beak. (Courtesyof G.F. Combs, University of Maryland.)al., 1982). Pantothenic acid deficiency reduces normal egg production
and hatchability. Subcutaneous hemorrhage and severe edema aresigns of pantothenic acid deficiency in the developing chick embryo.
In chickens, there is first a decline of growth, followed by decline in
feed conversion and retardation of feather growth. Plumage becomesrough and ruffled, and feathers become brittle and may fall; next, dermatosis rapidly develops in chicks. Corners of the beak and the area
below the beak are the most affected, but the disorder is also observedin feet. Outer layers of skin between toes and on bottoms of feet peeloff, and small cracks and fissures appear. In some cases, skin layers offeet thicken and cornify, and wart-like protuberances develop on ballsof the feet. The foot problem is usually exacerbated by bacterial invasion of the lesions. Within 12 to 14 days of receiving a deficientdiet, the margins of the eyelids are sealed closed by a viscous discharge. (Fig. 10.5 illustrates the typical deficiency syndrome in thechick.)
Pantothenic acid concentrations in liver are reduced during deficiency. Liver is hypertrophied and varies in color from a faint yellow toa dirty yellow. Nerves and fibers of the spinal cord show myelin degeneration, with these degenerating fibers occurring in all segments of thecord down to the lumbar region (Scott et al., 1982).
In young chicks, clinical signs of pantothenic acid deficiency are difficult to differentiate from those of biotin deficiency—both cause severedermatitis, broken feathers, perosis, poor growth, and mortality (seeChapter 11, Deficiency). In pantothenic acid deficiency, dermatitis of the434

PA N T O T H E N I C A C I D

Fig. 10.6 Pantothenic
acid deficiency in aturkey with dermatitis
on lower beak and atangle of mouth (lower
turkey). Sticky exudatethat formed on the eyelid resulted in encrustation and caused swollen
eyelids to remain stuck
together. Normal turkey
above is the control.
(Courtesy of T.M. Ferguson [deceased] andJ.R. Couch, Texas
A&M University.)feet is evident over the toes, in contrast to biotin deficiency, which primarily affects the foot pads and is often more severe than pantothenic
acid deficiency.
Signs of pantothenic acid deficiency in young turkeys are similar
to those in young chickens and include general weakness, dermatitis,and sticking together of eyelids (Fig. 10.6). Young ducks do not show
the usual signs seen in chickens and turkeys, except for retarded
growth; however, their mortality rate is very high. Poor feathering is
the most prevalent deficiency sign in pheasants and quail (Scott et al.,1964).
Pantothenic acid deficiency does not normally affect egg production
but severely depresses hatchability, and chicks that hatch may be too
weak to survive. Embryonic mortality in pantothenic acid deficiency occurs usually during the last few days of incubation. A direct linear relationship exists between diet pantothenic acid and hatchability. Beer et al.
(1963) fed a purified diet to White Leghorn hens that contained 0.9 mg
of pantothenic acid per kilogram of diet. They found that the hens required addition of 1.0 mg/kg for optimum egg production, at least 4.0
mg/kg for maximum hatchability, and 8.0 mg/kg for optimum hatchability and viability of offspring. Dawson et al. (1962) reported thatturkey breeder hens fed a diet deficient in pantothenic acid demonstrated a high embryonic mortality during the first week of development. After 17 days, the surviving embryos were small and poorly feathered, and showed signs of edema, hemorrhaging, fatty livers, and pale
dilated hearts.
435

CHAPTER TEN

Horses
Pantothenic acid deficiency has not been reported in the horse(NRC, 1989b). Likewise, no requirement for the vitamin has been reported. Pantothenic acid is synthesized in the intestinal tract of adulthorses. Carroll et al. (1949) fed a diet containing 0.8 mg of pantothenic acid per kilogram of dry matter and found the following pantothenic acid concentrations (ppm dry matter) in ingesta: duodenum,
11.7; ileum, 9.2; cecum, 39.2; anterior large colon, 34.4; and anterior
small colon, 20.5. No signs of pantothenic acid deficiency were observed in horses on this diet or on a second diet containing less than
0.2 mg/kg.

Other Animal Species
Experimental pantothenic acid deficiencies have been induced inmost animal species studied. However, the deficiency is not present inanimals with sufficient microbial fermentation (e.g., ruminants andhorses) or animals that practice sufficient coprophagy (e.g., rabbits).

DOGS AND CATS
Pantothenic acid-deficient dogs exhibit erratic appetites, reduced
rates of growth, lowered antibody response, and reduced blood concentrations of cholesterol and total lipids. Diarrhea becomes severe andbloody just prior to death. Vomiting is frequent. Animals become quiet
and very weak prior to death. Schaeffer et al. (1942) reported that vomiting was sometimes so severe in pantothenic acid-deficient dogs that feces occurred in the vomit. In terminal stages of deficiency, dogs exhibited spasticity of the hindquarters, sudden prostration, or coma, usuallyaccompanied by rapid respiratory and cardiac rates and possibly convulsions (NRC, 1985a). Deficient dogs have reduced pantothenate concentrations in urine, blood, liver, muscle, and brain.
In kittens, pantothenic acid deficiency is characterized chiefly byemaciation (NRC, 1986). Moderate to marked fatty metamorphosis of
the liver occurs, with both fine and coarse vacuolar formation. Giant,blunted villi are seen in some areas of the jejunum and upper ileum, withthe tops of the villi in some animals necrotic.

FISH
In rapidly growing fish (e.g., fingerling yellowtail), pantothenic aciddeficiency has been found to appear within 10 to 14 days. Gill lamellar
hyperplasia, or clubbed gills, is a characteristic sign of pantothenic acid436

PA N T O T H E N I C A C I D

deficiency in most fish (Lovell, 1987; NRC, 1993). In addition to
clubbed gills, anemia and high mortality have been observed in pantothenic acid-deficient salmonids. In blue tilapia and rainbow trout, deficiency signs included exudated gills, fusion of gill filaments and lamellae, severe fin erosion, and abnormal swimming (Roem et al., 1991;
Masumoto et al., 1994). Pantothenic acid-deficient Japanese parrot fishexhibited anorexia, convulsions, cessation of growth, and a high mortality rate (Ikeda et al., 1988). Poor growth, hemorrhage, skin lesions,and abnormal swimming were found in Japanese eels receiving pantothenic acid-deficient diets (NRC, 1993).

FOXES AND MINKDeficiency signs in pantothenic acid-deprived mink include severe
emaciation, enlarged adrenals, and general gastric ulceration, with petechial (minute) hemorrhages in the jejunum (NRC, 1982a). No definitive pantothenic acid studies in foxes have been completed.

LABORATORY ANIMALS
Pantothenic acid deficiency retards the development of weight in alllaboratory animals. Feed consumption is reduced because of appetite
loss. Animals become apathetic and noticeably less active (Friesecke,1975). In mild pantothenic acid deficiency, serum triglyceride and freefatty acid levels are elevated prior to weight loss and more severe signsof the deficiency (Wittwer et al., 1990).
In rats, lack of pantothenic acid results in premature graying of hair(achromotrichia). For this reason pantothenic acid has been called the“anti-gray-hair factor.” Another term applied to rats on a deficient dietis “bloody whiskers.” Porphyrin from the lachrymal glands of the pantothenic acid-deficient rat accumulates on the whiskers and gives the appearance of blood. The same condition can be produced when water intake is reduced to 50% of normal, so it may be due to adrenal
degeneration. Pantothenic acid deficiency also induces exfoliative dermatitis, oral hyperkeratosis, necrosis, and ulceration of the gastrointestinal tract. Focal or generalized hemorrhagic necrosis of the adrenals
may occur, and death results after 4 to 6 weeks of deficiency (NRC,1995). Pantothenic acid-deficient rats have impaired antibody synthesis,
decreased serum globulins, and decreased antibody-forming cells in response to antigen. Reproduction is affected, with litter size and birthweight reduced. Newborn rats exhibit neurological defects in the formof locomotor disturbances, incoordination of movements, and motor
spasms.
437

CHAPTER TEN

Signs of pantothenic acid deficiency in growing mice are reported to
be weight loss, hair loss (particularly on the ventral surface, flanks, andlegs), dermatosis, partial posterior paralysis, other neurological abnormalities, and achromotrichia. Young guinea pigs fed a purified, pantothenic acid-deficient diet developed signs of deficiency such as decreased growth rate, anorexia, weight loss, rough hair coat, diarrhea,
weakness, and death. Adult guinea pigs fed a pantothenic acid-deficient
diet died within 10 to 41 days. Many of them had adrenal and gastrointestinal hemorrhages (NRC, 1995).

NONHUMAN PRIMATES
Pantothenic acid deficiency in rhesus monkeys results in reduced
growth, anemia, hair loss, and ataxia (NRC, 1978).

HumansIn humans, pantothenic acid deficiency has not been observed undernatural conditions except when associated with severe malnutrition. The
main reason is that pantothenic acid deficiency is rare because of the vitamin’s widespread distribution in foods. Deficiency has been producedin volunteers consuming a pantothenic acid-deficient diet, but more rapidly by also including the antagonist ω-methylpantothenic acid (seeChemical Structure, Properties, and Antagonists) in the diet. The deficient diet alone may require about 12 weeks to produce recognizable
symptoms (Hodges et al., 1958). Symptoms most commonly present
were persistent and annoying fatigue, headache, muscle weakness, andcramps. Some subjects demonstrated impaired motor coordination andpeculiar gait. The neuromotor disorders included paresthesia of thehands and feet with hyperactive deep tendon reflexes. Neuromotor disorders could be explained by the role of acetyl coenzyme A in the synthesis of the neurotransmitter acetylcholine.
Other signs of pantothenic acid deficiency are gastrointestinal disturbances, apathy, depression, cardiovascular instability, increased susceptibility to infections, and impaired adrenal function. Increased susceptibility to upper respiratory tract infections presumably reflects theimpairment of immune response.
Deficiency of the vitamin was reported to cause “burning feet syndrome” during World War II among prisoners in Japan and the Philippines (Gopalan, 1946). This syndrome is occasionally described in malnourished people in developing countries. Symptoms include abnormalskin sensations of the feet and lower legs, which were increased by438

PA N T O T H E N I C A C I D

warmth and lessened by cold. Supplementation with large quantities of
pantothenic acid increased the ability of subjects to withstand stress.

Assessment of Status
Urinary excretion has been used in humans to estimate status ofpantothenic acid as well as the vitamin content of the diet. Urinary excretions of less than 1 mg/day are considered to be in the deficiency zone
corresponding to intakes of possibly less than 4 mg/day (Olson, 1990).
Like urine, the pantothenic acid content of human milk reflects the pantothenic acid content of the diet. Johnston et al. (1981) reported a correlation of 0.62 between dietary and milk content of pantothenic acid.
Total pantothenic acid levels in whole blood below 100 μg/dL would besuggestive of deficiency (Fox, 1991).
Studies of pantothenic acid deficiency in various animal species indicate lowered tissue levels and decreased urinary excretion of the vitamin, and decreased tissue coenzyme A (Nelson, 1978). However, data
are insufficient to establish pantothenic acid status for animals based on
critical concentrations. There is a close relationship between theamounts of dietary pantothenic acid in hen diets, egg production, eggconcentrations, and newly hatched chicks and poults (Pearson et al.,1945; Robel, 1993), indicating potential for status determination.

SUPPLEMENTATION
Monogastric animal diets based on grains, particularly corn, areroutinely supplemented with pantothenic acid. Scott et al. (1982) concludes that practical diets usually contain sufficient pantothenic acid for
all classes of chickens, but a number of factors may influence the requirement for this vitamin. Because of this, supplemental calcium pantothenate is usually added to diets for chicks and breeding hens. Increasing supplemental pantothenic acid to turkey breeding hens
increased the transfer of pantothenic acid in eggs (Robel, 1993). Scott(1966) indicated that the pantothenic acid requirement for poultry may
have to be increased 60 to 80% because of a lack of availability frombound forms in feeds. Southern and Baker (1981) estimated that pantothenic acid present in both corn and dehulled soybean meal was 100%
bioavailable. The bioavailability of pantothenic acid in barley, wheat,and sorghum, however, was estimated to be only 60% available based
upon chick growth bioassay. Nevertheless, corn and sorghum contained
less bioavailable total pantothenic acid than barley and wheat because439

CHAPTER TEN

of less total content of the vitamin. Clinical pantothenic acid deficiency
signs appear to be completely reversible, if not too far advanced, by oral
treatment or injection with the vitamin followed by restoration of an adequate level of pantothenic acid in the diet.
Type of diet will influence need for pantothenic acid supplementation. High-protein diets would reduce pantothenic acid needs due to thedecreased level of dietary carbohydrate, which would presumably require coenzyme A for metabolism. Pigs fed a pantothenic acid-deficient,high-fat diet failed to gain weight, exhibited a lower feed-efficiency ratio, and developed deficiency signs more quickly than pigs fed a diet low
in fat (Sewell et al., 1962). Biotin and folacin have been found necessary
for pantothenic acid utilization, and vitamin B12 and antibiotics (see Requirements) have a sparing effect for chicks (Latymer and Coates, 1981)
and pigs (Latymer et al., 1985).
Pantothenic acid is available as a commercially synthesized productfor addition to feed. It is available as d- or dl-calcium pantothenate. One
gram of d-calcium pantothenate is equivalent to 0.92 g of d-pantothenic
acid activity, while the combination of 1 g of the dl-form has 0.46 g ofd-pantothenic acid activity. A racemic mixture (equal parts of d- and lcalcium pantothenate) is generally sold to the feed industry. Because livestock and poultry can biologically utilize only the d-isomer of pantothenic acid, nutrient requirements for the vitamin are routinely
expressed in the d-form.
Feed-grade pantothenic acid products are available in a number ofpotencies. Products that are sold on the basis of racemic mixture content
can be misleading and confusing to a buyer who is not fully aware of thebiological activity supplied by d-pantothenic acid. To avoid confusion,
the label should clearly state the grams of d-calcium pantothenate or its
equivalent per unit weight and the grams of d-pantothenic acid.
A straight racemic mixture (90%) is available to people in the feedindustry, but its hygroscopic and electrostatic properties contribute tohandling problems. Because it readily picks up moisture, it sticks tobags, cans, and scoops and can become hard after prolonged exposure
to air. Its electrostatic properties cause it to cling to metallic and otherobjects, and losses can be significant. Through complexing procedures,several companies now market free-flowing and essentially nonhygroscopic and nonelectrostatic products.
Verbeeck (1975) reported calcium pantothenate to be stable in premixes with or without minerals and regardless of the mineral form.
Losses of calcium pantothenate may occur in premixes that are extremely acidic in nature, however. If a calcium pantothenate–calcium
440

PA N T O T H E N I C A C I D

chloride complex is used instead of the plain calcium pantothenate, thisproblem should be alleviated.
In the United States, estimates of pantothenic acid dietary intakes in
the human adult population range between 5 and 20 mg/day (Chung etal., 1961). These levels are higher than estimated requirements of 4 to 7
mg/day for children and adults (RDA, 1989). Therefore, supplementation would not normally be recommended unless associated with general malnutrition. Humans at greatest risk and needing supplementation
include alcoholics and diabetics; adrenal and digestive dysfunctions also
influence need for the vitamin (Fox, 1991).

TOXICITY
Pantothenic acid toxicity has not been reported in humans. The
LD50 in milligrams per kilogram of body weight for mice and rats has
been reported to range from 0.83 to 10.0 for calcium pantothenate(Unna and Greslin, 1941). In rats, 100 times the dietary requirement resulted in nonfatal liver damage (NRC, 1987).
Pantothenic acid has only limited pharmacological effects when administered to animals or humans. Dexpanthenol, the alcohol synthetically derived from d-pantothenic acid, has been administered to increase
gastrointestinal peristalsis and applied topically to alleviate itching andimprove minor skin irritations (Fox, 1991). Pantothenic acid has beenreported to have a protective effect against radiation sickness (Egarova
and Perepelkin, 1979). Some work suggests that pantothenic acid may
accelerate wound healing in rabbits by increasing the fibroblast content
of scar tissue (Aprahamian et al., 1985).

■ REFERENCES
Aprahamian, M., Dentinger, A., Stock-Damge, C., Kouassi, J.C., and Grenier,
J.F. (1985). Am. J. Clin. Nutr. 41, 578.
Axelrod, A.E. (1971). Am. J. Clin. Nutr. 24, 265.
Barboriak, J.J., and Krehl, W.A. (1957). J. Nutr. 63, 601.
Beer, A.E., Scott, M.L., and Neshein, M.C. (1963). Br. Poult. Sci. 4, 243.
Brass, E.P. (1993) J. Nutr. 123, 1801.
Carroll, F.D., Gross, H., and Howell, C.E. (1949). J. Anim. Sci. 8, 290.
Chung, A.S.M., Pearson, W.N., Darby, W.J., Miller, O.N., and Goldsmith, G.A.
(1961). Am. J. Clin. Nutr. 9, 573.
Colby, R.W., Cunha, T.J., Lindley, C.E., Cordy, D.R., and Ensminger, M.E.
(1948). J. Am. Vet. Med. Assoc. 113, 589.
Combs, G.F. Jr. (1992). The Vitamins. Academic Press, San Diego, California.
441

CHAPTER TEN

Cunha, T. J. (1977). Swine Feeding and Nutrition. Academic Press, Inc., New
York. 7, 1074.
Dawson, E., Ferguson, T.M., Deyoe, C.W., and Counch, J.R. (1962). Poult. Sci.
41, 1639.
Egarova, N.D., and Perepelkin, S.R. (1979). Gig. Sanit. 10, 25.
Ensminger, M. E., Colby, R. W., and Cunha, T. J. (1951). Wash. Agric. Exp. Stn.
Circ. 134, Pullman, Washington.
Fenstermacher, D.K., and Rose, R.C. (1986). Am. J. Physiol. 250, G155.
Fox, H.M. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 437. Marcel Dekker, New York.
Friesecke, H. (1975). In Pantothenic Acid, No. 1533. Hoffmann-La Roche,
Basel, Switzerland.
Fry, P.C., Fox, H.M., and Tao, H.G. (1976). J. Nutr. Sci. Vitaminol. 22, 339.
Gadient, M. (1986). In Proc. Maryland Nutrition Conference for Feed Manufacturers, p. 73. College Park, Maryland.
Goodwin, R.F.W. (1962). J. Comp. Pathol. 72, 214.
Gopalan, C. (1946). Indian Med. Gaz. 81, 23.
Hodges, R.E., Ohison, M.A., and Bean, W.B. (1958). J. Clin. Invest. 37, 1642.
Ikeda, M., Ishibashi, Y., Murata, O., Nasu, T., and Harada, T. (1988). Bull. Jpn.
Soc. Sci. Fish 54, 2029.
Johnston, L., Vaughan, L., and Fox, H.M. (1981). Am. J. Clin. Nutr. 34, 2205.
Latymer, E.A., and Coates, M.E. (1981). Br. J. Nutr. 45, 431.
Latymer, E.A., Mitchell, G., Coates, M.E., Keol, H.D., Thomas, J., and Woodley, S.C. (1985). Livest. Prod. Sci. 12, 265.
Loosli, J. K. (1991). In Handbook of Animal Science (P. A. Putnam, ed.). Academic Press, San Diego, California.
Lovell, R.T. (1987). Feed Manage. 38, 28.
Luecke, R. W., Hoefler, J.A., and Thorpe, F. Jr. (1953). J. Anim. Sci. 12, 605.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
Masumoto, T., Hardy, R.W., and Stickney, R.R. (1994). J. Nutr. 124, 430.
McKigney, J.I., Wallace, H.D., and Cunha, T.J. (1957). J. Anim. Sci. 16, 35.
Nelson, M.M., and Evans, H.M. (1945). Proc. Soc. Exp. Biol. Med. 60, 319.
Nelson, R.A. (1978). In Handbook Series in Nutrition and Food. Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 3, p. 33. CRC Press, Boca Raton, Florida.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
442

PA N T O T H E N I C A C I D(1995). Nutrient Requirements of Laboratory Animals.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Olson, R.E. (1990). In Nutrition Reviews, Present Knowledge in Nutrition (R.E.
Olson, ed.), p. 208. Nutrition Foundation, Washington, DC.
Palagina, N.K., Meledina, T.V., and Karpisheva, I.A. (1990). Appl. Biochem.
Microbiol. 26, 688.
Pearson, P.B., Melass, V.H., and Sherwood, R.M. (1945). Arch. Biochem. 7,
353.
Plaut, G.W.E., Smith, C.M., and Alworth, W.L. (1974). Annu. Rev. Biochem.
43, 899.
Pugh, E.L., and Wakil, S.J. (1965). J. Biol. Chem. 240, 4727.
RDA. (1989). Recommended Dietary Allowances, 10th Ed., National Academyof Sciences-National Research Council, Washington, D.C.
Robel, E.J. (1993). Poult. Sci. 72, 1740.
Roem, A.J., Stickney, R.R., and Kohler, C.C. (1991). Prog. Fish Culturist 53,
216.
Roy, R.B., and Buccafuri, A. (1978). J. Assoc. Off. Anal. Chem. 61, 720.
Sauberlich, H.E. (1973). In Modern Nutrition in Health and Disease, 5th ed.
(R.S. Goodhart and M.E. Shils, eds.), p. 203. Lea and Febiger, New York.
Sauberlich, H.E. (1985). Prog. Food Nutr. 9, 1.
Schaefer, A.E., McKibben, J.M., and Elvehjem, C.A. (1942). J. Biol. Chem. 143,
321.
Scott, M.L. (1966). Proc. Cornell Nutr. Conf. p. 35.
Scott, M.L., Holm, E.R., and Reynolds, R.E. (1964). Poult. Sci. 43, 1543.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken.
Scott & Associates, Ithaca, New York.
Sewell, R.F., Price, D.G., and Thomas, M.C. (1962). Fed. Proc. Fed. Am. Soc.
Exp. Biol. 21, 468(Abstr.).
Sheppard, A.J., and Johnson, B.C. (1957). J. Nutr. 61, 195.
Slinger, S.J., and Pepper, W.F. (1954). Poult. Sci. 33, 633.
Song, W.O. (1990). Nutr. Today, March-April, 19.
Song, W.O., Smith, A.,Wittwer, C.,Wyse, B., and Hansen, G. (1990). Nutr. Res.
10, 439.
Southern, L.L., and Baker, D.H. (1981). J. Anim. Sci. 53, 403.
Taylor, T., Hawkins, D.R., Hathway, D.E., and Partington, H. (1972). Br. Vet.
J. 128, 500.
Taylor, T., Cameron, B.D., Hathway, D.E., and Partington, H. (1974). Res. Vet.
Sci. 16, 271.
Teague, H.S., Grifo, A.P. Jr., and Palmer, W.M. (1971). J. Anim. Sci. 33, 239(Abstr.).
Ullrey, D.E., Becker, D.E., Terrill, S.W., and Notzold, R.A. (1955). J. Nutr. 57, 40.
Unna, K., and Greslin, J.C. (1941). J. Pharmacol. Exp. Ther. 73, 85.
443

CHAPTER TENVerbeeck, J. (1975). Feedstuffs 47 (36), 4.
Virtanen, A.I. (1966). Science 153, 1603.
Wittwer, C.T., Beck, S., Peterson, M., Davidson, R., Wilson, D.E., and Hansen,
R.G. (1990). J. Nutr. 120, 719.
Wright, L.D., and Welch, A.D. (1943). Science 97, 426.
Wyse, B.W., and Hansen, R.G. (1977). Fed. Proc. 36, 1169.

444

chapter eleven

BIOTININTRODUCTION
For many years it was believed that supplemental biotin was not required in swine and poultry diets because of its wide distribution in feedstuffs and because of known biotin synthesis by the intestinal microflora. However, in the mid-1970s, field cases in these species were
found under modern production systems. These conditions, characterized by specific clinical signs, responded to supplemental biotin. On thebasis of these findings, nutritionists have had to reexamine the role ofbiotin in livestock and poultry diets. In many different animal species,
biotin deficiency is usually associated with epidermal lesions.
Incidence of biotin deficiency has been found occasionally when humans and animals have consumed excessive quantities of raw eggs,
which contain a biotin complexing factor (avidin). Likewise, biotin deficiency is reported in children with inborn errors of metabolism when
there are insufficient biotin-dependent enzymes. Such cases in children
respond dramatically to high-level dietary supplementations with biotin.

HISTORY
Discovery of the physiological significance of biotin is the history of
the merging of different lines of investigation that appeared to be unrelated (Scott et al., 1982; Bonjour, 1991; Loosli, 1991). Biotin was thename given to a substance isolated from egg yolk by Kögl and Tönnis in1936 that was necessary for yeast growth. This substance was discovered to be identical to a growth factor named coenzyme R that was required by legume nodule bacteria.
The toxic properties of feeding raw egg white to animals were first445

11CHAPTER ELEVEN

observed by Bateman in 1916. Clinical signs of dermatitis and hair loss
due to egg-white injury were prevented by several researchers by feedingcertain foods, notably liver and kidney. György studied the chemistry ofthis protective factor in certain foods, which he named factor H in 1937.
The term vitamin H was chosen by György because the factor protected
the haut, the German word for skin.
In 1940 György and associates found that biotin, vitamin H, andcoenzyme R were the same substance. Other names given to this factor
were protective factor X, egg-white injury protection factor, factor S,
and factor W (or vitamin Bw). Egg-white injury resulted from its inactivating dietary biotin because of a specific constituent, avidin. Heat was
found to destroy avidin, therefore revealing that this biotin antagonistic
factor was heat labile. Avidin is a protein that is a secretory product ofthe mucosa of the oviduct and therefore is found in the albuminous partof eggs.
The structure and properties of biotin were established by U.S. andEuropean investigators between 1940 and 1943. The first chemical synthesis was completed by Harris and associates of the Merck Company
in 1943. In 1949 Goldberg and Sternbach developed a technique for industrial production of biotin.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTSThe chemical structure of biotin in metabolism includes a sulfur
atom in its ring (like thiamin) and a transverse bond across the ring (Fig.
11.1). The empirical formula for biotin is C11H18O3N2S. Biotin is a fusion of an imidazolidone ring with a tetrahydrothiophene ring bearing avaleric acid side chain. It is a monocarboxylic acid with sulfur as athioether linkage. Biotin, with its rather unique structure, contains three
asymmetric carbonations, and therefore eight different isomers are possible. Of these isomers only one contains vitamin activity, d-biotin. The
stereoisomer l-biotin is inactive.
Biotin crystallizes from water solution as long, white needles. Its
melting point is 232 to 233oC. Free biotin is soluble in dilute alkali andhot water and practically insoluble in fats and organic solvents. Biotin isquite stable under ordinary conditions. It is destroyed by nitrous acid,other strong acids, strong bases, and formaldehyde and is inactivated by
rancid fats and choline (Scott et al., 1982). It is gradually destroyed byultraviolet radiation.

446

BIOTIN

Fig. 11.1 Structures of biotin and some derivatives.

Structurally related analogs of biotin (Fig. 11.1) can vary from noactivity to partial replacement value and to antibiotin activity. Mild oxidation converts biotin to sulfoxide, and strong oxidation converts it tosulfone. Strong agents result in sulfur replacement by oxygen, resultingin oxybiotin and desthiobiotin. Oxybiotin has some biotin activity forchicks (one-third) but less for rats (one-tenth). Both desthiobiotin andbiotin sulfone are active for yeast but are inhibitory to bacteria. Biocytin, a bound form of biotin, is also biologically active in several
species, including animals. This naturally occurring compound has beenshown to be ε-N-biotinyl-L-lysine. Besides these biotin analogs, othercompounds exist that can bind biotin to form a stable complex, thus
preventing the utilization of the vitamin by animals and/or microorganisms. The microorganism Saccharomyces avidinii produces a biotinbinding protein streptavidin and other compounds that can inactivate
free biotin and apparently inhibit biotin synthesis in susceptible microorganisms. Oleic acid and related compounds, in the presence of aspartic acid, satisfy the biotin requirement of many microorganisms, but
are inactive for some and actually toxic for others.

447

CHAPTER ELEVENANALYTICAL PROCEDURES
A number of methods are available for estimating biotin content offeeds and supplements. An enzyme-linked immunosorbent assay has
been used to determine biotin in blood and other fluids (Wellenberg andBanks, 1993). This assay utilizes the strong binding property of streptavidin. Bioassay procedures with animals (rats and chicks) and microorganisms are employed because of the difficulty of establishing chemicalor physical methods suitable for natural materials. However, Achuta
Murthy and Mistry (1977) reported progress in the use of colorimetric,gas chromatographic, and polarographic methods of analyses. Isotope
dilution and radiotracers have been used for biotin analysis and are sensitive, yielding results comparable to the Lactobacillus plantarum assay(Bonjour, 1991). Nevertheless, such analytical methods are not capableof identifying amounts of this vitamin that are biologically available tothe animal.
In the biological estimation, rats and chicks are made biotin deficientby the use of special diets. Rats are made deficient by resorting to the useof sulfonamides, preventing coprophagy, feeding egg white (avidin), or
combinations of these techniques. However, deficiency in the chick canbe produced by a diet low in biotin alone. For both species, biotin content of the test sample can be calculated from growth-response curves.
The healing of skin lesions in avidin-fed rats is a sensitive indicator of biotin availability. An additional test to evaluate feeds is based on the pyruvate carboxylase activity in poultry or rat blood, either directly or by assaying the in vitro activity before and after addition of biotin (Whitehead
et al., 1982; Bonjour, 1991). Most studies of biotin content of feedstuffs
have been conducted by microbiological assay. Lactobacillus plantarum
is often employed with other suitable organisms, including L. casei and
Saccharomyces cerevisiae. The procedure with microorganisms includes
heating with H2SO4 to liberate bound forms and removing unsaturated
fatty acids that interfere with the assay. A simple, rapid, and economic
method for assessing biotin availability of feedstuffs was developed using
egg yolk and plasma biotin concentrations after hens were fed specificfeeds (Buenrostro and Kratzer, 1984).

METABOLISM
Biotin exists in natural materials in both bound and free forms, withmuch of the bound biotin apparently not available to animal species. For
448

BIOTIN

poultry, often less than one-half of the microbiologically determined biotin in a feedstuff is biologically available (Scott, 1981). Naturally occurring biotin is found partly in the free state (fruit, milk, vegetables)and partly in a form bound to protein in animal tissues, plant seeds, andyeast. Variation in availability appears to be due to differential susceptibilities to digestion of the biotin-protein linkages in which the vitamin isfound in natural products. Those linkages involve the formation of covalent bonds between the carboxyl group of the biotin side-chain withthe amino acid lysine or to protein.
Biotinidase, present in pancreatic juice and intestinal mucosa, releases biotin from biocytin during the luminal phase of proteolysis. In
most species that have been investigated, physiological concentrations ofbiotin are absorbed from the intestinal tract by a sodium-dependent active transport process, which is inhibited by desthiobiotin and biocytin(Said and Derweesh, 1991). Absorption of biotin by a Na+-dependent
process was noted to be higher in the duodenum than the jejunum,
which was in turn higher than that in the ileum, and it was concluded
that the proximal part of the human small intestine was the site of maximum transport of biotin (Said et al., 1988).
The few studies conducted in animals on biotin metabolism revealed
that biotin is absorbed as the intact molecule in the first third to half ofthe small intestine (Bonjour, 1991). There is also absorption of biotinfrom the hind gut of the pig, with disappearance of between 50 and
61% of infused biotin between the cecum and feces that was accompanied by more than a fourfold increase of plasma biotin concentrationand more than a sixfold increase of urinary biotin excretion (Barth et al.,1986). Kopinski and Leibholz (1985) reported that postileal absorption
was 10 to 15% of that from the small intestine after oral ingestion.
Eighty percent of a labeled biotin dose infused into the cecum of minipigs appeared in portal blood (Drouchner and Volker, 1984) with thelargest proportion appearing in feces. Using 14C-labeled biotin, Kopinski et al. (1989a,b) reported similar findings in that absorption of freebiotin in the postileal digestive tract was about 8% as efficient as thatfrom a similar labeled dose of orally administered biotin. Kopinski et al.
(1989c) observed that even with extensive microbial synthesis of biotinin the postileal tract, low concentrations of biotin in plasma and tissue,
and the presence of deficiency signs indicated that postileal synthesizedbiotin is of limited benefit to the pig. Scholtissek et al. (1990) suggested
that under basal conditions, 1.7 to 17% of the requirement for biotin isprovided by colonic bacteria.
Biotin appears to circulate in the bloodstream both free and bound449

CHAPTER ELEVEN

to a serum glycoprotein, which also has biotinidase activity, catalyzing
the hydrolysis of biocytin. In humans, 81% of biotin in plasma was free
and the remainder bound (Mock and Malik, 1992). Information is very
limited on biotin transport, tissue deposition, and storage in animals andhumans. Mock (1990) reported that biotin is transported as a free water-soluble component of plasma, is taken up by cells via active transport, and is attached to its apoenzymes. Said et al. (1992) reported thatbiotin is transported into human liver via a specialized carrier-mediatedtransport system. This system is Na+-gradient dependent and transports
biotin via an electroneutral process.
Disappearance of an intravenous dose of radioactive biotin fromblood of biotin-deficient rats was more rapid than for controls (Petrelli
et al., 1979). Also, rate and extent of deposition into deficient liver, particularly mitochondrial and cytosolic fractions, were favored. This research supports the concept of homeostatic mechanisms responding toprovide biotin in relation to needs. All cells contain some biotin, withlarger quantities in the liver and kidneys. Intracellular distribution of biotin corresponds to known localization of biotin-dependent enzymes(carboxylases).
Investigations into biotin metabolism in animals and humans are
difficult to interpret, as biotin-producing microorganisms exist in the intestinal tract distal to the cecum. Often, the amount of biotin excreted
in urine and feces together exceeds total dietary intake, whereas urinary
biotin excretion is usually less than intake. 14C-labeled biotin showedthat the major portion of intraperitoneally injected radioactivity was excreted in the urine and none in the feces or expired as CO2 (Lee et al.,1973). In rats and pigs, biliary excretion of biotin and metabolites was
negligible (Zempleni et al., 1997).
The brush border of the kidney cortex has a sodium-biotin transportsystem similar to that in the intestinal mucosa, thus providing for resorption of free biotin filtered into the urine. It is only when this resorption mechanism is saturated that there will be significant excretionof biotin. Efficient conservation of biotin, together with the recycling ofbiocytin released from the catabolism of biotin-containing enzymes, may
be as important as intestinal bacterial synthesis of the vitamin in meeting biotin requirements (Bender, 1992).

FUNCTIONS
Biotin is an essential coenzyme in carbohydrate, fat, and protein metabolism. It is involved in conversion of carbohydrate to protein and vice450

BIOTIN

versa as well as conversion of protein and carbohydrate to fat. Biotin
also plays an important role in maintaining normal blood glucose levels
from metabolism of protein and fat when dietary intake of carbohydrate
is low. Biotin functions as a carboxyl carrier in four carboxylase enzymes: pyruvate carboxylase, acetyl CoA carboxylase, propionyl CoA
carboxylase, and 3-methylcrotonyl CoA carboxylase. As a componentof these carboxylating enzymes, there is the capacity to transport carboxyl units and to fix carbon dioxide (as bicarbonate) in tissue. Biotin
serves as a prosthetic group in a number of enzymes in which the biotin
moiety functions as a mobile carboxyl carrier. The biotin prosthetic
group is linked covalently to the ε-amino group of a lysyl residue of thebiotin-dependent enzyme.
In carbohydrate metabolism, biotin functions in both carbon dioxide fixation and decarboxylation, with the energy-producing citric acidcycle dependent on the presence of this vitamin. It is suggested that partof the effect of biotin on energy production is through stimulation ofphosphorylation pathways (Moretti et al., 1990). Glucokinase activity isaffected by biotin, with activity of the enzyme decreasing with biotin deficiency (Li-Hsieh and Mistry, 1992). Specific biotin-dependent reactions in carbohydrate metabolism are the following:■ Carboxylation of pyruvic acid to oxaloacetic acid.
■ Conversion of malic acid to pyruvic acid.
■ Interconversion of succinic acid and propionic acid.
■ Conversion of oxalosuccinic acid to α-ketoglutaric acid.

In protein metabolism, biotin enzymes are important in protein synthesis, amino acid deamination, purine synthesis, and nucleic acid metabolism. Biotin is required for transcarboxylation in degradation ofvarious amino acids. Deficiency of the vitamin in mammals hinders thenormal conversion of the deaminated chain of leucine to acetyl-CoA.
Depletion of hepatic biotin results in reduction of hepatic activity of
methylcrotonyl-CoA carboxylase, which is needed for leucine degradation (Mock and Mock, 1992). Ability of liver homogenates to incorporate labeled CO2 was directly related to biotin level of the animal (Terroine, 1960). Likewise, ability to synthesize citrulline from ornithine
was reduced in liver homogenates from biotin-deficient rats. The urea
cycle enzyme ornithine transcarbamylase was significantly lower in livers of biotin deficient rats (Maeda et al., 1996).
Acetyl-CoA carboxylase catalyzes addition of CO2 to acetyl-CoA toform malonyl-CoA. This is the first reaction in the synthesis of fatty451

CHAPTER ELEVEN

acids. A cytosolic multienzyme complex, fatty acid synthetase, then accomplishes synthesis of palmitate from malonyl-CoA. Biotin is required
for normal long-chain unsaturated fatty acid synthesis and is importantfor essential fatty acid metabolism (Kramer et al., 1984). Deficiency inrats inhibited arachidonic acid (20:4 ω-6) synthesis from linoleic acid(18:2 ω-6) while increasing linolenic acid (18:3 ω-3) and its metabolite(22:6 ω-3) (Kramer et al., 1984; Watkins and Kratzer, 1987a). Biotin deficiency resulted in reduced arachidonic acid (20:4) in chicks, and therefore reduced plasma prostaglandin E2 (PGE2) as a result of having lessof the prostaglandin precursor (20:4) (Watkins and Kratzer, 1987b;
Watkins, 1989).

REQUIREMENTS
Estimates of biotin requirements for various animals and humans
are presented in Table 11.1. Biotin requirements are difficult to establish
because of variability in feed content and bioavailability. Likewise, it isdifficult to obtain a quantitative requirement for biotin because the vitamin is synthesized by many different microorganisms and certain
fungi. These microorganisms are found in the lower part of the intestinal
tract, a region in which absorption of nutrients is generally reduced.
However, it is believed that intestinal microflora make a significant contribution to the body pool of available biotin. In general, combined urinary and fecal excretion of biotin exceeds the dietary intake. What is notknown for the various species is the extent of microbial synthesis or thebiotin availability to the host.
It is concluded that microorganisms contribute to animal and human requirements because the use of some sulfa drugs, such as Sulfathalidine, can induce deficiency under some circumstances. Microorganisms that provide significant quantities of biotin to most species
apparently supply a variable and undependable amount of biotin for
poultry (Scott, 1981). Biotin deficiency was less severe in cecectomized
than normal chickens, indicating that cecal microorganisms do not supply chickens with significant amounts of biotin but instead compete withthe host animal for dietary biotin, thus increasing the requirement(Sunde et al., 1950).
Rate and extent of biotin synthesis may be dependent on the level ofother dietary components. In poultry and rats, it has been shown thatpolyunsaturated fats, ascorbic acid, and other B vitamins may influence
the demand for biotin. Addition of polyunsaturated fatty acids to fat-452

BIOTIN■ Table 11.1

Biotin Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf
Adult
Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100-g intake)
Broilers, 0–6 weeks
Starting and growing
Breeding
0–4 weeks
4–8 weeks
Breeding hens
Adult
Growing-finishing
Breeding-lactating
Adult
Adult
All classes
Growing
All classes
Common carp
Pacific salmon
Rainbow trout
Infants
AdultsMicrobial synthesis
0.1 ppm milk replacer
Microbial synthesis
0.15 mg/kg
0.10 mg/kg
0.10 mg/kg
0.15 mg/kg
0.3 mg/kg
0.15 mg/kg
0.25 mg/kg
0.20 mg/kg
0.10 mg/kg
Microbial synthesis
0.05 mg/kg
0.20 mg/kg
Microbial synthesis (?)
Microbial synthesis
0.07 mg/kg
0.12 mg/kg
0.6 mg/kg
1 mg/kg
1–1.5 mg/kg
0.05–0.14 mg/kg
10–15 μg/day
30–100 μg/day

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC(1981b)
NRC (1986)
NRC (1982a)
NRC (1978a)
NRC (1993)
NRC (1993)
NRC (1993)
RDA (1989)
RDA (1989)

ChickenJapanese quail
Turkey
Sheep
Swine
Horse
Goat
Cat
Mink
HamsterFish
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as μg/day. Often requirements are unknown, so the best recommendation is used.

free, biotin-deficient diets increased severity of dermal lesions (Roland
and Edwards, 1971). Biotin is rapidly destroyed as feeds become rancid.
Pure biotin was inactivated to the extent of 96% in 12 hours whenlinoleic acid of a high peroxide number was added to the diet (Pavcek
and Shull, 1942). In the presence of α-tocopherol, this destruction
amounted to only 40% after 48 hours.
Verification that humans require biotin was documented in two situations: prolonged consumption of raw egg white and parenteral nutrition without biotin supplementation in patients with short-gut syndrome and other causes of malabsorption (Mock, 1990). Generalrecommendations for human biotin requirements vary from 10 to 100
μg/day; uncertainty of requirements is due to lack of information on the
nutritional significance of biotin synthesis and utilization of biotin byenteric bacteria (RDA, 1989).

453

CHAPTER ELEVEN

NATURAL SOURCES
Biotin is present in many foods and feedstuffs (Table 11.2). The
richest sources of biotin are royal jelly, liver, kidney, yeast, blackstrap
molasses, peanuts, and eggs. Most fresh vegetables and some fruits are
fairly good sources. Corn, wheat, other cereals, meat, and fish are relatively poor sources. In comparison to cereal grains, oilseed meals are
better sources of total biotin. Soybean meal, for instance, contains amean biotin content of 270 ppb, with a range of 200 to 387 ppb (Frigg
and Volker, 1994).
Animal proteins, on the other hand, are rather unreliable sources of
biotin. For instance, highly variable levels are found in fish meals (135ppb on average, ranging from 11 to 421 ppb) and meat meals (88 ppb
on average, ranging from 17 to 322 ppb).
The biotin content of cereal grains appears to be influenced by variety, season, and yield (in particular, the endosperm-pericarp ratio)(Brooks, 1982). Harvest conditions, origin of sample, year of collection,
postharvest treatment, and storage conditions all appear to play a partin determining biotin content and may also affect the availability. Thereis considerable variation in biotin content within individual sources. For
example, 65 samples of corn analyzed for biotin varied between 0.012
and 0.072 ppm, and 20 samples of meat meal varied between 0.008 and0.20 ppm (Brooks, 1982).
Less than one-half of the biotin in various feedstuffs is biologicallyavailable (Table 11.2) as determined by microbiological assay (Frigg,
1976, 1984; Frigg and Volker, 1994). Thus, it is important to know thechemical form of biotin (e.g., bound or unbound) as well as its overall
content in feed. Oilseed meals, alfalfa meals, and dried yeast are excellent sources of biologically available biotin. Meat meal and fish meals
contain biotin of relatively poor biological availability. Sauer et al.
(1988) determined that an apparent biotin digestibility of soybean meal
was 55.4%, while that of meat and bone meal was only 2.7%, andcanola meal was 3.9%. Grains are poor sources of biotin, with corn andoats more available than wheat, hull-less barley, or regular barley. Frigg(1976) reported that the biotin content of wheat and barley may be totally unavailable. The excretion of biotin in feces of pigs given wheat
was 10 times greater than from corn, indicating greater utilization ofcorn biotin (Bryden, 1989).
Biotin in feedstuffs may be destroyed by heat curing, solvent extraction, and improper storage conditions, while pelleting has little ef454

BIOTIN■ Table 11.2

Typical Biotin Contents and Bioavailability in Various Foods and Feedstuffs (as-fed basis)

Food or Feedstuff
Alfalfa meal, dehydrated
BarleyBeef, steak
Cabbage
Carrots
ChickenCorn, yellow
Corn gluten meal
Cottonseed meal, solvent extracted
Distiller’s solubles, dried
Eggs, whole
Fish meal
Liver, beef
Meat and bone meal
Milk, cow’s
Molasses, blackstrap
Oats
Peanut meal
Rice bran
Rice polishings
Skim milk, dried
Sorghum
Soybean meal
Wheat
Wheat bran
Yeast, brewer’s

Biotin Levels (μg/g)
0.33
0.14
0.04
0.02
0.03
0.10
0.08
0.19
0.08–0.47
0.44–1.1
0.25
0.14
1.00
0.08
0.05
0.70
0.25
1.63
0.42
0.37
0.25
0.29
0.27
0.10
0.36
0.63

Biotin Availability (%)
75–100
10
—
—
—
—
100
100
100
90
—
100
—
100
—
—
35
53
—
20
65
20
100
—
20
100

Sources: Modified from NRC (1982b) and Frigg and Volker (1994).

fect on biotin content of feed (McGinnis, 1986a,b). Milling of wheat or
corn and canning of corn, carrots, spinach, or tomatoes reduced biotin
concentrations (Bonjour, 1991). Biotin is unstable to oxidizing conditions and therefore is destroyed by heat, especially under conditions that
support simultaneous lipid peroxidation. These losses can be reduced by
the use of an antioxidant (e.g., vitamin C, vitamin E). After baby foods
were stored for 6 months, an approximate 15% reduction in biotin content was found (Karlin and Foisy, 1972).

DEFICIENCY
Effects of Deficiency
Biotin is important for normal function of the thyroid and adrenal
glands, the reproductive tract, and the nervous system. However, the ef455

CHAPTER ELEVEN

fect of biotin deficiency on the cutaneous system is most dramatic sincea severe dermatitis is the major obvious clinical sign of deficiency in livestock and poultry.

Ruminants
Biotin deficiency, identified by hindquarter paralysis, decreased urinary excretion of biotin, and correction of the problem with biotin injections was reported in calves (Wiese et al., 1946). Flipse et al. (1948)
reported a potassium-biotin interrelationship in calves, with the resultthat calves fed purified diets low in potassium and biotin developed progressive paralysis of the hind legs that spread to the forelegs, neck, andrespiratory system. Death may result within 12 to 24 hours of the firstsigns; however, the condition can be cured by injections of potassium
salts or biotin.
Due to ruminal and intestinal synthesis of biotin, a need for supplemental sources was at one time not expected for ruminants. Nevertheless, promising preliminary results in preventing lameness in dairy cattlewith biotin supplementation were reported (Frigg et al., 1993). Successful biotin treatment of dairy cows with claw problems was reported by
Nietlis-Bash and Triebel (1988). In biotin-deficient dairy cows, the hoof
horn is of poor quality, soft, and crumbling, with no distinct separationof keratinizing and cornified cells. This results in the omission of the
granular layer at the epidermis of the bulb of the heel. Decreased stabilizing filaments in the upper spiny layer of the hoof corium in biotin-deficient cows reveals the depressed hormone-like activity of biotin in thesynthesis of protein (Budras et al., 1997).
Increased plasma biotin levels have been associated with hardness
and conformational changes in the bovine hoof. Dairy cows supplemented with 20 mg of biotin per cow over an 11-month period expressed a steepened angle of the dorsal border and height of the heel;
length of the diagonal and size of the ground surface increased (Distl andSchmid, 1994). The hardness of the hoof was also significantly greaterin the biotin-treated group. Greenough et al. (1999) reported that biotin
supplementation (20 mg/day) to dairy cows not only reduced hoof lesions, but significantly increased milk production.
Beef cattle suffer from a common hoof defect known as sandcracks,
which are vertical fissures in the hoof. These tend to be more prevalent
in older, heavier cows and often result in chronic lameness problems inbeef cattle. Biotin supplementation in 15 beef cow herds in which 37.2%of the cows were affected with sandcracks dramatically reduced the proportion of affected cows (Campbell et al., 1995). This study indicates456BIOTINthat biotin supplementation appears to improve hoof quality by lowering the number of sandcracks per cow and decreases occurrence of otherhoof defects, such as broken toes and abnormal coronary bands. Feeding dairy and beef cows 20 mg/day of supplemental biotin resulted in reduced incidence of hoof lesions and increased milk production (Seymour, 1999).

Swine
Biotin deficiency was produced in 1946 in swine by feeding a purified diet containing Sulfathalidine or raw egg white (Cunha et al., 1946;
Lindley and Cunha, 1946). For many years it was concluded that biotin
supplementation was not needed since it is synthesized by intestinal microorganisms and is widely distributed in feed. Nevertheless, clinicalsigns of deficiency were observed by feed company personnel and scientists under field conditions. However, it wasn’t until the 1970s that agreater awareness of the problem of biotin field deficiency became apparent (Cunha, 1984).
Biotin deficiency results in reduced growth rate and impaired feed
conversion as well as a wide variety of clinical signs. These include
alopecia (hair loss); dermatitis characterized by dryness, roughness, anda brownish exudate; ulceration of the skin; inflammation of the mouth
mucosa; hind-leg spasticity; and transverse cracking of the soles andtops of hooves (Cunha, 1977). Clinical signs of biotin deficiency inswine are shown in Figs. 11.2 and 11.3.
Growth depression may become evident in biotin-deficient swine
before clinical signs are seen. The first clinical signs are generally excessive hair loss and dermatitis, with complete hair loss in severe cases. Dermatitis first appears as scaly skin, often starting on the ears, neck, shoulder, and tail and eventually spreading over the entire body. In laterstages, crust and cracks appear on the face and extremities.
After 5 to 7 weeks on a biotin-deficient diet, swine may show claw
defects. Foot lesions may be the most characteristic sign of biotin deficiency. The hoof horn becomes soft and rubbery and is poorly resistantto abrasions. The slow growth-and-repair process in the hoof tissue andthe considerable weight on the feet add to the problem. Depending on
the floor type on which the animal is kept, this may have little effect or
may lead to the development of cracks and necrotic lesions, resulting in
extreme lameness (Glättli, 1975). Secondary infections may gain entry
through claw cracks and result in infected joints, which may lead to premature removal from the herd. Feeding and breeding activities are also
adversely affected; in particular, the sow becomes unable to support the457

CHAPTER ELEVEN

Fig. 11.2 The two middle pigs are biotin deficient. Note the hair loss and dermatitis. (Courtesy of T.J. Cunha and Washington State University.)

weight of the boar. Also, because the hog’s ability to eat may be impaired, these problems will obviously lead to economic losses.
Biotin supplementation improved the strength of the pig claw horn
(Kempson et al., 1989). Supplementary biotin affected the structure of thecoronary epidermis; there was an increase in the density of the horntubules in the stratum medium, the horny squames in the stratum medium
were more tightly packed, and the tubules were more clearly defined in thepigs receiving biotin. The body condition of sows during lactation, and in
the period from weaning to remating, was improved by biotin supplementation (Greer et al., 1991). Twelve months after the experiment commenced, the percentage of sows with foot lesions was lower in the biotinsupplemented than the unsupplemented group (42 versus 55).
Investigations of lameness in sows showed that pigmented claws were lessliable to develop lesions than unpigmented claws, that claws with highmoisture content were relatively weak, that feeding biotin increased claw
hardness under certain conditions, and that the frequency of claw lesions
increased during pregnancy (Kroneman et al., 1992).
458

BIOTIN

Fig. 11.3 Biotin-deficient pigs. Note transverse cracking of the soles and thetops of the hooves. (Courtesy of T.J. Cunha and Washington State University.)

Supplementation of the diet of breeding sows with biotin from anearly stage of development made a significant contribution to the maintenance of hoof-horn integrity (Simmins and Brooks, 1988). Tagwerker(1983) noted that foot lesions were responsible for 4 to 8% of all sowsculled in Europe. Also, he noted a Denmark study that reported 8.5%of hoof lesions in biotin-supplemented sows compared to 25% for controls. After biotin supplementation, in Holland, culling rate due tolameness was decreased from 25 to 14% (de Jong and Sytsema, 1983).
Cunha (1984) noted that in most of the 40 countries he visited during the past 30 years, biotin deficiency signs were observed in swine operations. These signs, observed under field conditions, occurred in only
10 to 20% of sows or less. Baby pigs nursing these sows usually showedno biotin deficiency signs but responded to biotin supplementation. Unfortunately, many swine producers are of the opinion that it is naturalfor a swine herd to have a few animals with hair loss, dermatitis, andcracked feet and therefore are not overly concerned when a small percentage of sows exhibit these clinical signs (Cunha, 1984).
459

CHAPTER ELEVEN

Biotin supplementation of sow diets has significantly improved reproductive performance, including the number of pigs farrowed andweaned, litter weaning weight, and number of days from weaning to estrus (Brooks et al., 1977; Simmins and Brooks, 1983; Misir and Blair,
1984; Kornegay, 1986).
In a field study, sows had severe lameness and impaired reproduction (Fonge, 1977). After these sows received supplemental biotin, normal foot health and normal reproductive performance were restored.
Researchers also found that sows housed in total confinement showed apositive response to conception rate and interval from weaning to first
estrus and a trend to larger litters when supplemented with biotin(Bryant et al., 1985). In an earlier study (Brooks et al., 1977), sows fed
supplemental biotin had more pigs born alive (9.8 versus 8.1), more pigs
weaned (7.8 versus 6.8), increased litter weight at weaning (71.0 versus
64.5 kg), and reduced time interval from weaning to first estrus after
weaning (6.2 versus 15.3 days) compared to unsupplemented controls.

Poultry
Biotin requirement in the turkey is higher than that of the chick, somore field problems with biotin deficiency have arisen in turkeys. Biotin
deficiency in chicks and poults results in a wide range of clinical signs(Figs. 11.4 through 11.7), with considerable variation in time of appearance of individual signs (NRC, 1994). The principal effects in both
species are reduced growth rate and feed efficiency, disturbed and broken feathering, dermatitis, and leg and beak deformities. First signs ofdeficiency are usually growth depression and loose feathering; signs ofdermatitis then appear and, finally, disorders of the leg (perosis) andbeak become apparent. With dermal lesions, bottoms of feet becomerough and calloused and contain deep fissures that show some hemorrhaging. Feet problems are usually exacerbated by bacterial invasion oflesions. Toes may become necrotic and slough off. Tops of feet and legs
usually show only a dry scaliness. Lesions appear in the corner of themouth and slowly spread to the whole area around the beak. Eyelids
eventually swell and stick together.
Dermal lesions have a characteristic order of appearance, although
speed of onset depends on deficiency severity. For chicks fed severely deficient diets, dryness and flakiness of the feet first become noticeable atabout 14 days of age, and slight encrustations and superficial fissures develop in the undersurfaces of the feet at about 18 days (Whitehead,1978). These increase in severity until, by about 25 days, the fissures be-460

Fig. 11.4 Perosis (bone deformities) as a result of biotin deficiency. Chicks
showed perosis as early as 17 days of age, with rigid limb joints that resulted
in a stilted walk. (Courtesy of Hoffmann-La Roche Inc., Nutley, New Jersey.)

Fig. 11.5 Normal (left) and biotin-deficient (right) Broad-Breasted Bronze maleturkeys at 3 weeks of age. (Courtesy of D.C. Dobson, Utah State University.)461

CHAPTER ELEVEN

Fig. 11.6 Biotin deficiency in a poult. Notelesion at apex of mouth.
(Courtesy of L.S. Jensen
and Washington State
University).

come hemorrhagic. Between 3 and 4 weeks, dermatitis may also appear
on the eyelids and, as this develops, the bird becomes unable to keep thelids apart and they eventually become stuck together.
Dermal lesions are similar to those of pantothenic acid deficiency(see Chapter 10). However, with biotin deficiency, lesions occur first in
the feet and later around the beak and eyes, whereas in pantothenic aciddeficiency, the signs occur first in the corners of the mouth and eyes andonly in prolonged cases appear in the feet. Because of the difficulty ofmaking a differential diagnosis between the two vitamins, it is often necessary to examine the diet composition and decide which is most likely
to be deficient.
Biotin deficiency is a cause of hock disorders in both poults andchicks. The major deficiency sign affecting market turkeys is severe leg
weakness. Lesions caused by biotin deficiency are brought about by462

BIOTIN

Fig. 11.7 Severe foot-pad lesions in the growing turkey as a result of biotin deficiency. Less severe lesions are more common. (Courtesy of Richard Miles,
University of Florida.)

chondrodystrophy, a condition in which bone mineralization is normal,
but linear growth of long bones is impaired. Chondrodystrophy causedby biotin deficiency can result in shortening of metatarsal bones and perosis. Perosis occurs when irregular bone development results in enlargement and deformity of the hock joint. Crippling in turkeys can occur as
early as 3 to 4 weeks of age and often seems to disappear at 6 to 7
weeks. Then it reappears with great severity between 13 and 16 weeks(Scott, 1981). At this stage, the birds are unable to walk and thus can be
trampled by other turkeys. Perosis can occur at any stage in the growthof the turkey from 3 to 24 weeks of age. In general, young chickens areless susceptible than poults to leg disorders, although biotin deficiency
does cause problems of the same type in chicks as in poults (Whitehead,1978). Once the deformities of perosis occur, biotin administration is
not curative.
Bain et al. (1988) report that “twisted leg” is the most common limb
disorder in broiler chickens and that biotin deficiency adversely affectedtibiotarsal bone growth. Tibiotarsal bones are frequently longitudinally
distorted in biotin-deficient poultry. Presumably, reduced biotin prevents463

CHAPTER ELEVEN

ready formation of prostaglandins from essential fatty acids, and bone
growth fails to respond to stresses during development (Watkins et al.,1989).
Low dietary fat and the necessity for fatty acid synthesis lead to anabnormal array of fatty acids that predisposes poultry to a fatty liverand kidney syndrome (FLKS) (Whitehead, 1988). This condition, which
has caused heavy economic losses in commercial broiler flocks, wasfound to be due to a suboptimal dietary biotin coupled with certain nutritional and environmental stress factors. Situations increasing themetabolic rate of biotin-dependent enzymes (pyruvate, acetyl-CoA carboxylase), such as low fat or protein levels, aggravate the condition. Although the signs of FLKS are not those of classic biotin deficiency, theycan virtually be eliminated by supplementing chick starter or breeder diets with biotin.
In turkeys, dry and brittle feathers usually accompany the othersigns of clinical biotin deficiency. Bronze poults can exhibit white barring of the feathers, usually affecting just tom turkeys. Likewise, deficient chicks have rough and broken feathering, with head and breastfeathers often having a spiky, matted appearance.
Poor egg production and hatchability will result from clinical biotin
deficiency (Robel, 1991; NRC, 1994). In breeder chickens, biotin deficiency will reduce hatchability but is less likely to affect egg production.
Clinical signs and conditions associated with biotin deficiency in chickembryos and/or newly hatched chicks include bone deformities (perosis),
impaired muscle coordination (ataxia), skeletal deformities (e.g.,crooked legs), extensive foot webbing, abnormal cartilage development(chondrodystrophy), embryonic mortality, twisted and malformed beak(“parrot beak”), and reduced size. In turkeys, Ferguson et al. (1961) reported that biotin deficiency resulted in a marked decrease in hatchability and a high rate of embryonic mortality during the first week of incubation. At the end of the second week, hatchability decreased from 83
to 14%. At the end of the third week, hatchability was zero. Embryonicmortality because of inadequate biotin occurs largely during the last 3days of incubation. Feeding the biotin-deficient diet resulted in an
abrupt decrease in egg production after 13 weeks.

HorsesBiotin is synthesized in the lower digestive tract of the horse, but information on dietary requirement is lacking (NRC, 1989b). Hoof integrity for a number of cases has been reported to improve as a result of
biotin supplementation (Fig. 11.8). Such cases were characterized by464

BIOTIN

Fig. 11.8 Hoof condition of a 7-year-old 18hands heavyweightshow hunter resulting
from biotin supplementation. The horse had ahistory of tender feet.
(A) The walls of thehooves were very weak,
crumbling at the lower
edges, with large areas
breaking out and detaching when shoes
were nailed. (B) Fivemonths after supplementation with 15 mgof biotin per day, thewalls of the hooves
were thicker and harder,
so nailing was achieved.
(Courtesy of N.
Comben, R.J. Clark,
and D.J.B. Sutherlandand permission of Veterinary Record 115[1984], 642–645.)

A

B

horn that tended to crumble at the lower edges of the hoof walls andthat was generally prone to poor conformation and damage to the walls,
soles, and the white line junction (Comben et al., 1984).
Biotin supplementation was given to 38 horses at 5 mg/150 kg bodyweight once daily for at least 9 months (Schulze and Scherf, 1989). Initially, all horses had hoof horn of poor quality. By the end of treatment
the hoof defects were regarded as cured in 24 horses and improved in
the remainder. In a 19-month experiment with Lipizzaner stallions, biotin supplementation decreased incidence and severity of horn defects,
increased tensile strength (4.5 versus 3.7 kp/mm2), and improved condition of the white line (Linden et al., 1993).

Other Animal Species
For a number of species, including guinea pigs, rats, hamsters, cats,and rabbits, no deficiency signs have been observed when animals were465

CHAPTER ELEVEN

fed purified diets and/or natural diets containing low biotin concentrations. However, with these same species, feeding a biotin-deficient dietcontaining raw egg white (Fig. 11.9) produced typical deficiency signs,
often including weight loss and alopecia.

DOGS AND CATS
Biotin-responsive disease conditions in dogs include dull coat, brittle hair, loss of hair, scaly skin pruritis, and dermatitis (NRC, 1985a;
Frigg et al., 1989). In a study with small animal practitioners, 60% of
clinical signs in dogs were cured, and another 31% improved (119 cases)in dogs with hair and skin conditions (Frigg et al., 1989).
In cats, severe, experimental biotin deficiency may be marked bybloody diarrhea, anorexia, and emaciation. Scaly dermatitis about thenose and mouth, generalized alopecia, and hypersalivation have been
noted (NRC, 1986; Pastoor et al., 1991). Most cats with the deficiency
develop signs of alopecia; dried secretion around the eyes, nose, mouth,and feet; focal dermatitis of the lips near the eyeteeth; and a brownish
appearance of the skin (Pastoor et al., 1991).

FISH
Biotin is synthesized in many species of fish by intestinal microflora,
but supplemental biotin may be required for maximum growth and toprevent clinical deficiency signs (NRC, 1993; Shiau and Chin, 1998).
Signs of biotin deficiency in trout and salmon include anorexia, slow
growth, increased mortality, and poor feed conversion. The “blue slime”
disease of trout can be prevented with dietary biotin. Biotin-deficientcatfish develop light skin pigmentation and anemia. Japanese eels fed abiotin-deficient diet exhibited reduced growth, abnormal swimming behavior, and dark coloration.
Common carp and channel catfish required from 8 to 12 weeks, andrainbow trout from 4 to 8 weeks, to show growth depression when fed
biotin-deficient diets (NRC, 1993). Signs of biotin deficiency were notdetected in rainbow trout or channel catfish fed natural ingredient diets
without supplemented biotin for 24 and 17 weeks, respectively (NRC,
1993). These studies concluded that adequate biotin was available in thenatural feed ingredients.

FOXES AND MINK
Biotin deficiencies have been produced by feeding purified diets ordiets containing egg products to growing mink. Apparently, the short intestinal tract of the mink and rapid feed passage make it impossible for466

BIOTIN

A

Fig. 11.9 (A) Egg-white
injury as a result of
feeding a rat raw egg
white, which contains abiotin antagonist,
avidin. The resulting
dermatitis progressed togeneralized alopecia. (B)After 3 months of treatment, the animal returned to normal.
(Courtesy of Alan T.
Forrester, Scope Manualon Nutrition, The Upjohn Company, Kalamazoo, Michigan.)Bthe animal to obtain adequate biotin from that synthesized by its intestinal flora. Stout et al. (1966) reported biotin deficiency in mink whendiets contained 40% or more offal from breeder hen turkeys. Presenceof raw eggs in the offal was presumed responsible for the deficiency.
Later studies in Oregon showed that inclusion of spray-dried egg in thediet can have a similar effect (Wehr et al., 1980). In both mink andfoxes, biotin deficiency resulted in graying or discoloration of hair (Fig.
11.10) and, in extreme cases, hair loss (NRC, 1982a).

LABORATORY ANIMALS
Signs of biotin deficiency (Fig. 11.9) in avidin-fed rats are weight
loss, exfoliative dermatitis, alopecia, achromotrichia, and an abnormal,
spastic gait (NRC, 1995). Heard et al. (1989) reported that biotin deficiency causes impaired auditory brainstem function in young biotin-deficient rats. Skin from biotin deficient rats contained a different pattern
467

CHAPTER ELEVEN

Fig. 11.10 The newborn
fox pup on the left isfrom a biotin-deficient
dam that received a dietcontaining raw egg
white. Thin, gray pelt
and deformed legs areapparent. A control dietcontaining cooked egg
white was fed to thedam of the newborn
pup on the right. (Courtesy of A. Helgebostad,
Veterinary College of
Norway, Heggedal.)of fatty acids than that of controls (Proud et al., 1990). Biotin deficiency
in rats and mice can result in a hind limb paralysis referred to as kangaroo gait.
Clinical signs for biotin deficiency in mice, hamsters, and guinea
pigs included alopecia, achromotrichia, and growth failure (NRC,1995). Biotin supplementation improved reproduction and lactation inmice when added to a purified diet devoid of egg white (NRC, 1995).
Biotin deficiency interfered with synthesis of specific proteins, which resulted in congenital malformations (e.g., cleft palate) (Watanabe, 1995).
On day 10 of gestation in hamsters, biotin deficiency resulted in a highincidence of resorbed and dead embryos, and by day 14, for the remaining embryos, there were growth retardation, morphological abnormalities, and skeletal defects (Watanabe, 1993).

NONHUMAN PRIMATES
Biotin deficiency was produced in adolescent rhesus monkeys withthinning of hair and loss of hair color as a result of a low-biotin diet for
468

BIOTIN

long periods (15 to 28 months) (Waisman et al., 1945). Use of diets withegg white and/or sulfonamides greatly hastened and intensified biotindeficiency signs.

RABBITS
Deficiency of biotin, characterized by loss of hair and dermatitis, occurred in rabbits that were fed raw egg white over a period of time
(Lease et al., 1937).

HumansExcept in infants, there is no evidence of spontaneous biotin deficiency in humans. This is probably due to the ubiquitous nature of thevitamin in diets plus benefits derived from microbial synthesis. Biotin
deficiency occurs in individuals consuming a large number of raw eggs
daily because of the antagonistic effect of the egg-white protein avidin(see History). Adult volunteers fed 200 g of dehydrated egg white daily
for 5 weeks displayed symptoms of mild depression, lassitude, somnolence, hallucination, anxiety with muscle pain, and hyperesthesia(Sydenstricker et al., 1942). After 8 weeks, anorexia and a striking grayish pallor with dermatitis and desquamation occurred. The experiment
was terminated because of a marked fall in food intake.
Biotin deficiency has also been demonstrated in biotinidase deficiency, with many clinical findings and biochemical abnormalities similar to those associated with biotin deficiency (Mock, 1990). Patientswith biotinidase deficiency cannot utilize biocytin, which is excreted intheir urine, and must depend solely upon free biotin from food to maintain normal biotin levels. Two conditions in infants—seborrheic dermatitis and desquamative erythroderma (Leiner’s disease)—are apparently connected with biotin deficiency, since children with these diseases
have subnormal biotin in blood and urine and respond to supplementation with the vitamin. Some children have an inborn error of metabolism resulting from lack of biotin-dependent carboxylases. Children withthis disorder can be divided into two groups, namely, those affected by
an isolated defect in only one and those with a combined deficiency of
the four biotin-dependent enzymes. The multiple carboxylase deficiency
apparently results from a deficiency in biotinidase activity.
Biotin deficiency can result when insufficient biotin is added to infant milk formulations or to total parenteral nutrition solutions. Infants
with short bowel syndrome receiving total parenteral nutrition lacking
in biotin showed an increase in serum odd-chain saturated fatty acids
and a decrease in linoleic acid compared with controls (Anonymous,469

CHAPTER ELEVEN

1989). Although the evidence is circumstantial, some researchers havesuggested that sudden infant death syndrome (SIDS) may be related to
low biotin intakes of bottle-fed infants (Bonjour, 1991). The median biotin levels in the livers of infants who died from SIDS between 1 and 52
weeks of life were significantly lower than those of infants who diedfrom explicable causes (Heard et al., 1983). Anticonvulsant drugs (e.g.,
carbamazepine and primidone) are competitive inhibitors of biotin
transport in the intestine and would relate to inadequate biotin status
for long-term therapy of these drugs (Said et al., 1989). Cases of low circulating plasma biotin have also been reported for alcoholics, elderly
persons, epileptics, and burn victims.

Assessment of Status
Detection of subclinical or marginal biotin deficiency, which is most
likely to occur under field conditions, is difficult. Field observations areimportant, but it is essential to realize that there may be no clinical signsin spite of deficiency. If clinical signs are seen, they may not all be present, and only a small percentage of animals may exhibit such clinicalsigns. Chemical, chick, or microbiological assays often give different results for feed biotin concentrations. Likewise, there is great variability infeed biotin concentrations and availability in various feeds (see Natural
Sources).
Whitehead et al. (1980) and Misir and Blair (1984) suggested thatplasma biotin concentration and plasma pyruvate carboxylase activity
are methods of assessing the biotin status of pigs. However, blood biotin
concentrations often lack dependability as a good indicator of biotin status of pigs (Cunha, 1984). This is particularly true with borderline deficiency, in which plasma biotin varies with dietary intake and shows considerable individual variation among pigs. In chickens, several reports
have indicated that plasma biotin concentration below 100 ng/100 ml isindicative of biotin deficiency (Scott et al., 1982). In three experiments
using chickens and turkeys, plasma pyruvic carboxylase activity was
positively related to supplemental biotin in both species (Whitehead andBannister, 1978). Robinson and Lovell (1978) reported that biotin status in fish may be assessed by measuring pyruvate carboxylase activity
in the liver. Hepatic pyruvate carboxylase activity in rainbow trout fed
a lipid-free and biotin-deficient diet decreased to 3.3% of normal (Walton et al., 1984). Biotin concentration in egg yolk is an additional
method of evaluating the status of the vitamin (see Analytical Procedures). In both poultry and swine, the ratio of the blood fatty acids is470

BIOTIN

likewise indicative of biotin status, with palmitoleic acid particularly increasing in relation to stearic acid (Edwards, 1974).
Mock and Mock (1992) reported abnormal catabolism of leucine asa basis for early detection of biotin deficiency in rats. 3-MethylcrotonylCoA originates from catabolism of leucine and is normally metabolized
to acetyl-CoA. However, in biotin deficiency, reduced hepatic activity ofthe biotin-dependent enzyme methylcrotonyl-CoA carboxylase causes
the enzyme’s substrate 3-methylcrotonyl-CoA to be shunted via an alternate pathway to 3-hydroxyisovaleric acid (3-HIA), which is excreted
at increased rates in the urine. Elevated concentrations of 3-HIA would
be an early indication of biotin inadequacy.
Evaluation of human biotin status employs blood and urinary biotin
and biotin metabolites (Mock et al., 1997). As with animals, there areconsiderable differences and variations among humans that are difficult
to explain. From the available data, a biotin level in urine of approximately 70 nmol/L and a circulating level in plasma of around 1,500
nmol/L seem to indicate an adequate supply of biotin for humans (Bonjour, 1991).

SUPPLEMENTATION
For supplementation in livestock, consideration must be given tocost, as biotin is one of the more expensive vitamins. However, based on
the requirements and recommended industry levels for biotin supplementation, it costs less than other vitamins, such as choline and vitamin
E, in monogastric diets. In view of the considerable cost of synthetically
produced biotin, the question of biotin supplementation of practical diets is economically significant.
Although clinical signs were found for poultry and swine under experimental conditions in the 1940s, for many years it was believed that
supplemental biotin was not needed in swine and poultry diets becauseof the production of biotin by the animal’s intestinal microflora. In 1967
for poultry and in the mid-1970s for swine, interest was rekindled whenmore field cases occurred than in the past. Interference with the biosynthesis of biotin by intestinal bacteria can individually or collectively leadto biotin deficiency. These interferences can be in the form of therapeutic administration of antibacterial agents and modern housing systems,
which limit animals’ access to feces. Additionally, biotin deficiency is
now more prevalent because of limited bioavailability of biotin found insome grains (e.g., wheat, barley, sorghum) and in some animal protein

471

CHAPTER ELEVEN

sources (e.g., meat meal, poultry by-product meal), biotin antagonists
including molds, feed rancidity, and improved genetic characteristics for
greater production. Cunha (1984) summarized the possible reasons formore frequent occurrence of biotin deficiency in swine (Table 11.3); themajority of these reasons are likewise applicable to the poultry industry.
Biotin deficiency in fur-bearing animals caused by feeding of egg products is economically disastrous because of interference with fur coloration, and supplementation is mandated.
Some reports conclude that foot lesions in adult sows healed in amatter of weeks (Tagwerker, 1974) when sows were supplemented withhigh levels of biotin, while Brooks et al. (1977) found that biotin supplementation resulted in a 28% reduction in lesions after 6 months. Pigs
housed on badly designed floors have little opportunity for recovery, as
traumatic injury exceeds the capacity of the hoof for growth and repair.
Brooks (1982) summarized data from Great Britain, Denmark, andSwitzerland and concluded that where foot lesions already existed, dietary supplements of 2,000 to 3,000 μg of biotin per kilogram of dietwere beneficial.
For many species receiving typical diets, supplementation is notneeded because of adequate sources in feed ingredients and/or intestinal
microflora synthesis. As an example of dietary adequacy, practical dietsfor channel catfish made from the commonly used ingredients—soybean
meal, corn, and menhaden fish meal—did not need supplemental biotin(Lovell and Buston, 1984). There may be less need for biotin supplementation for swine fed a high-quality corn-soybean diet that has no
mold (short-term storage) and for swine raised under favorable environmental and nutritional conditions (Watkins et al., 1991).
For some species, including equines, the need for supplemental biotin is controversial. If hoof problems exist, daily biotin supplementation has been suggested (Comben et al., 1984) for each animal as follows: donkeys and ponies, 5 to 10 mg; riding horses, 15 mg; and heavy
horses, up to 30 mg. The recommendation is to continue supplementation as increased hoof horn strength may be anticipated within 3 to 5
months. Using scanning electron microscope observations, Kempson(1987, 1994) concluded that biotin-related improvements in hoof quality are limited to the stratum externum and not the stratum medium or
stratum internum. He suggested that structural defects in the stratum externum of the hoof horn could be remedied by biotin supplementation
alone, but poor attachment of the horn squames required biotin plus
supplemental calcium and protein.
Young ruminants not receiving their mother’s milk and before the472BIOTIN■ Table 11.3

Possible Explanations Why Biotin Deficiencies Have Become More
Prevalent in Swine and Poultry Operations in Recent Years

Increased use of confinement, which lessens opportunity for coprophagy (feces eating);
feces containing biotin synthesized in the intestines.
Increased use of grain–soybean meal diets, with less utilization of biotin-rich feeds including whey, fermentation by-products, yeast, dehydrated alfalfa, and pasture.
Biotin antagonists in feeds such as streptavidin, certain antimicrobial drugs, and dieldrin(a pesticide). Streptomyces are molds affecting biotin availability that are found in
soil, moldy feeds, manure, and litter.
Rancidity in feeds causes biotin to be readily destroyed. Biotin in feedstuffs may be destroyed by heat curing, solvent extraction, pelleting, and improper storage conditions.
Length of storage, temperature, and humidity result in biotin loss.
Reduced intestinal synthesis and/or absorption of biotin may result from diseases andother conditions affecting the gastrointestinal tract.
Improved genetic characteristics (breed, type, and strain) and intensified production forfaster weight gains and better feed conversion. Increased reproductive performancewith more farrowings per sow yearly or increased egg production in poultry operations.
Decreased level of biotin or its availability in feeds because of new plant varieties, newfeed production practices, and processing methods.
For swine, reduced feed consumption (thus reduced biotin intake) by sows to avoid excess weight, which is detrimental to reproduction. Sows now receive 1.4–2.3 kg feed
per day during gestation whereas previously they had received 2.7–3.2 kg.
Interrelationships between biotin and other nutrients affect requirements. Dietary fats,pantothenic acid, vitamin B6, vitamin B12, folacin, thiamin, riboflavin, myo-inositol,
and ascorbic acid are related to biotin requirements and metabolism.
Lower levels and availability of biotin in feeds. Different batches of the same feed mayvary considerably in biotin content. Biotin in feeds exists in both free and boundforms. Certain feeds have a low availability (i.e., wheat, barley, sorghum). A shift
from use of corn, soybean meal, and cottonseed meal often results in diets with lessavailable biotin.
Source: Adapted from Cunha (1984).

rumen is developed are recommended to receive supplemental biotin.
Milk replacers for calves are recommended to contain 0.1 mg/kg of biotin (NRC, 1989a). In acute cases of biotin deficiency in calves, single
injections of 100 μg of biotin subcutaneously or 1 mg of biotin intravenously reversed the deficiency (Wiese et al., 1946). Dietary biotin supplementation (e.g., 20 mg/day) for adult ruminants would be recommended where field conditions suggest a need (Seymour, 1999). Ruminal
bacteria involved in the synthesis of biotin appear to be sensitive to low
pH; therefore, high grain diets reduce rumen synthesis (Seymour, 1999).
Supplementation of human diets is generally considered unnecessary
because of adequate dietary intakes as well as intestinal microbial synthesis. Exceptions would be for individuals consuming raw eggs and infants with inborn errors of metabolism. Eggs are a good source of biotin;
however, the biotin contained in egg yolk is inadequate to counteract the473

CHAPTER ELEVEN

deficiency of the vitamin caused by the avidin in egg white, so unheateddried whole egg is deficient in this vitamin (Kratzer et al., 1988). In addition, since the advent of total parenteral nutrition, biotin deficiency
has resulted in some cases in which biotin was omitted from intravenous
fluids (Bonjour, 1991). In humans, biotin supplements in the form of injections of 150 to 300 μg/day can clear up symptoms of deficiency in amatter of a few days.
Biotin is commercially available as a 100% crystalline product or as
dilutions (45.5 mg/kg and as a 1% premix). The form containing vitamin activity is d-biotin, which occurs in nature and is the commerciallyavailable form. A 2% spray-dried biotin product is also available for use
in either feed or drinking water. An example of supplemental biotin provided in water for biotin-deficient young turkeys is 0.25 mg of biotin per
gallon of drinking water for 3 to 4 weeks, and for older birds 0.50 mg
per gallon (Bauernfeind, 1969).
Losses of biotin during storage can be considerable. Biotin is readily destroyed by feed rancidity (Pavcek and Shull, 1942). Preparing fresh
feeds, storing them for only short periods, and keeping them dry and ina well-ventilated storage area will minimize rancidity problems. Hamilton and Veum (1984) have suggested that overdrying, poor storage conditions, and presence of mold may reduce the availability of biotin incorn. Also, the diet should be low in feeds that are high in prooxidants
and/or properly protected by an effective antioxidant to avoid destruction of biotin, vitamin E, selenium, and other nutrients.
Biotin is relatively stable in multivitamin premixes, unless combinedwith choline or trace minerals. Coelho (1991) reported only 57% retention of biotin when the vitamin supplement contained choline and traceminerals. Biotin is generally stable in feeds, but losses of the vitamin during processing do occur. Processing losses of biotin in extruded pet foodhave been reported at about 15% (Anonymous, 1981).

TOXICITY
For both humans and animals, biotin is apparently not toxic even inlarge doses. However, Paul et al. (1973) reported that an acute dose of
biotin (5 mg/100 g of body weight) caused irregularities of the estrus cycle, with heavy infiltration of leukocytes in the vagina of the rat up to
14 days after treatment. In additional rat studies, a dose of biotin at least
5,000 or 10,000 times the daily requirement had no deleterious effects(Mittelholzer, 1976). Studies with poultry and swine indicate a safety
tolerance of 4 to 10 times their requirement, with maximum tolerable
474

BIOTINlevel probably much higher (NRC, 1987). Likewise with humans, no adverse effects have resulted from oral or intravenous administration ofhigh doses over prolonged periods.

■ REFERENCES
Achuta Murthy, P.N., and Mistry, S.P. (1977). Prog. Food Nutr. Sci. 2, 450.
Anonymous (1981). Rationale for Roche Recommended Vitamin Fortification,
RCD 5963/1280. Hoffmann-La Roche, Nutley, New Jersey.
Anonymous (1989). Nutr. Rev. 47, 121.
Bain, S.D., Newbrey, J.W., and Watkins, B.A. (1988). Poult. Sci. 67, 590.
Barth, C.A., Frigg, M., and Hagemeister, H. (1986). J. Anim. Phys. Anim. Nutr.
55, 128.
Bauernfeind, J.C. (1969). World Rev. Anim. Prod. 5, 20.
Bender, D.A. (1992). Nutritional biochemistry of the vitamins, Cambridge University Press, Cambridge, England.
Bonjour, J.P. (1991). In Handbook of Vitamins 2nd Ed. (L.J. Machlin, ed.), p.
403. Marcel Dekker, New York.
Brooks, P.H. (1982). Pig News Inf. 3, 1.
Brooks, P.H., Smith, D.A., and Irwin, V.C.R. (1977). Vet. Res. 101, 46.
Bryant, K.L., Kornegay, E.T., Knight, J.W., Webb, K.E. Jr., and Notter, D.R.
(1985). J. Anim. Sci. 60, 145.
Bryden, W.L. (1989). Br. J. Nutr. 62, 389.
Budras, K.D., Hochstetter, T., Muelling, Ch., and Natterman, W. (1997). J.
Dairy Sci. 80(Suppl. 1), 192.
Buenrostro, J.L., and Kratzer, F.H. (1984). Poult. Sci. 63, 1563.
Campbell, J., Greenough, P.R., and Petrice, L. (1995). Western College of Vet.
Med. University of Saskatchewan, Saskatoon, Canada.
Coelho, M.B. (1991). Vitamin Stability in Premixes and Feeds: A Practical Approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Comben, N., Clark, R.J., and Sutherland, D.J.B. (1984). Vet. Rec. 115, 642.
Cunha, T.J. (1977). Swine Feeding and Nutrition, Academic Press, New York.
Cunha, T.J. (1984). Feed Manage. 35, 14.
Cunha, T.J., Lindley, D.C., and Ensminger, M.E. (1946). J. Anim. Sci. 5, 219.
de Jong, M.F., and Sytsema, J.R. (1983). Vet. Q. 5, 58.
Distl, O., and Schmid, D. (1994). Tierarzliche Umschauy 49, 581.
Drouchner, W., and Völker, L. (1984). Proc. TAD Symposium, Cattle and Pig
Diseases. Cuxhaveny 1, 105.
Edwards, H.M. (1974). In Proc. Nutr. Conf., p. 1. Atlanta.
Ferguson, T.M., Whiteside, C.H., Creger, C.R., Jones, M.L., Atkinson, R.L., andCouch, J.R. (1961). Poult. Sci. 40, 1151.
Flipse, R.J., Huffman, C.F., Duncan, G.W., and Thorp, F. (1948). J. Anim. Sci.
7, 525.
Fonge, J. (1977). Pig Farm. 25(6), 61.
Frigg, M. (1976). Poult. Sci. 55, 2310.
Frigg, M. (1984). Poult. Sci. 63, 750.
475

CHAPTER ELEVEN

Frigg, M., and Volker, L. (1994). Feedstuffs 66(1), 12.
Frigg, M., Schulze, J., and Volker, L. (1989). Schweizer-Archiv-fur-Tierheilkunde131, 621.
Frigg, M., Straub, O.C., and Harmann, D. (1993). Int. J. Vit. Nutr. Res. 63, 122.
Glättli, H.R. (1975). Zentralbl. Veterinaermed. A 22, 102.
Greenough, P.R., Gay, J.M., Dobson, R.C., and Gay, C.C. (1999). J. Dairy Sci.
82(Suppl. 1.), 34 (Abstr.)
Greer, E.B., Leibholz, J.M., Pickering, D.I., Macoun, R.E., and Bryden, W.L.
(1991). Aust. J. Agr. Res. 42, 1013.
Hamilton, C.R., and Veum, T.L. (1984). J. Anim. Sci. 59, 151.
Heard, G.S., Hood, R.L., and Johnson, A.R. (1983). Med. J. Aust. 2, 305.
Heard, G.S., Lenhardt, M.I., Bowie, R.M., Clarke, A.M., Harkins, S.W., andWolf, B. (1989). FASEB J. 3, A1242.
Karlin, R., and Foisy, C. (1972). Int. J. Vit. Nutr. Res. 42, 545.
Kempson, S.A. (1987). Vet. Rec. 120, 568.
Kempson, S.A. (1994). In Ninth Annual Bluegrass Laminitis Symposium.
Louisville, Kentucky.
Kempson, S.A., Currie, R.J.W., and Johnston, A.M. (1989). Vet. Rec. 124, 37.
Kopinski, J.S., and Leibholz, J. (1985). Proc. Nutr. Soc. Aust. 10, 170.
Kopinski, J.S., Leibholz, J., and Love, R.J. (1989a). Brit. J. Nutr. 62, 767.
Kopinski, J.S., Leibholz, J., and Love, R.J. (1989b). Brit. J. Nutr. 62, 781.
Kopinski, J.S., Leibholz, J., Bryden, W.L., and Fogarty, A.C. (1989c). Brit. J.
Nutr. 62, 751.
Kornegay, E.T. (1986). Livestock Prod. 14, 65.
Kramer, T.R., Briske-Anderson, M., Johnson, S.B., and Holman, R.T. (1984). J.
Nutr. 114, 2047.
Kratzer, F.H., Knollman, K., Earl, L., and Buenrostro, J.L. (1988). J. Nutr. 118,604.
Kroneman, A., Vellenga, L., Vermeer, H.M., and Van-der Wilt, F.J. (1992). Proefverslag-Proefstation-Voor-de-Varkenshouderij, pp. 1, 78.
Lease, J.G., Parsons, H.T., and Kelly, E. (1937). Biochem. J. 31, 433.
Lee, H.M., McCall, N.E., Wright, L.D., and McCormick, D.B. (1973). Proc.
Soc. Exp. Biol. Med. 142, 642.
Li-Hsieh, Y.T., and Mistry, S.P. (1992). Nutr. Res. 12(6), 787.
Linden, J., Josseck, H., Zenker, W., Geyer, H., and Schulze, J. (1993). In Proc.
13th Equine Nutrition and Physiology Symposium, p. 58. University of
Florida, Gainesville.
Lindley, D.C., and Cunha, T.J. (1946). J. Nutr. 32, 47.
Loosli, J.K. (1991). In Animal Science Handbook (P.A. Putnam, ed.). Academic
Press, San Diego.
Lovell, R.T., and Buston, J.C. (1984). J. Nutr. 114, 1092.
Maeda, Y., Kawata, S., Inui, Y., Fukuda, K., Igura, T., and Matsuzawa, Y.
(1996). J. Nutr. 126, 61.
McGinnis, C.H. (1986a). Water Soluble Vitamins. Rhone Poulenc, Atlanta.
McGinnis, C.H. (1986b). Bioavailability of Nutrients in Feed Ingredients, p. 1.
National Feed Ingredient Association (NFIA), Des Moines, Iowa.
Misir, R., and Blair, R. (1984). J. Anim. Sci. 59(Suppl. 1), 254.
Mittelholzer, E. (1976). Int. J. Vit. Res. 46, 33.
476

BIOTIN

Mock, D.M. (1990). In Nutrition Reviews, Present Knowledge in Nutrition(R.E. Olson, ed.), p. 189. Nutritional Foundation, Washington, D.C.
Mock, D.M., and Malik, M.I. (1992). Am. J. Clin. Nutr. 56, 427.
Mock, D.M., Stadler, D.D., Stratton, S.L., and Mock, N.I. (1997). J. Nutr. 127,
710.
Mock, N.I., and Mock, D.M. (1992). J. Nutr. 122, 1493.
Moretti, P., Petrelli, C., Petrelli, F., and Sciarresi, P. (1990). Med. Sci. Res. 18,
73.
Nietlis-Bash, C., and Triebel, D.F. (1988). Die Gruene 116, 28.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Pastoor, F.J.H., Van Herick, H., Van’t Klooster, A., and Beynen, A.C. (1991). J.
Nutr. 121, 573.
Paul, P,K., Duttagupta, P.N., and Agarwal, H.C. (1973). Curr. Sci. 42, 613.
Pavcek, P.L., and Shull, G.M. (1942). J. Biol. Chem. 146, 351.
Petrelli, F., Moretti, P., and Paparelli, M. (1979). Mol. Biol. Rep. 4, 247.
Proud, V.K., Rizzo, W.B., Patterson, J.W., Heard, G.S., and Wolf, B. (1990). Am.
J. Clin. Nutr. 51, 853.
RDA. (1989). Recommended Dietary Allowances, 10th Ed. National Academyof Sciences-National Research Council, Washington, D.C.
Robel, E.J. (1991). Poult. Sci. 70, 1716.
Robinson, E.H., and Lovell, R.T. (1978). J. Nutr. 108, 1600.
Roland, D.A., and Edwards, H.M. (1971). J. Nutr. 101, 811.
Said, H.M., and Derweesh, I. (1991). Am. J. Phys. 261, R94.
Said, H.M., Redha, R., and Nylander, W. (1988). Gastroenterology 95, 1312.
Said, H.M., Redha, R., and Nylander, W. (1989). Am. J. Clin. Nutr. 49, 127.
Said, H.M., Hoefs, J., Mohammadkhani, R., and Horne, D.W. (1992). Gastroenterology 102, 2120.
Sauer, W.C., Mosenthin, R., and Ozimek, L. (1988). J. Anim. Sci. 66, 2583.
Scholtissek, J., Barth, C.A., Hagemeister, H., and Frigg, M. (1990). Br. J. Nutr.
64, 715.
477

CHAPTER ELEVEN

Schulze, J., and Scherf, H. (1989). Tierarztliche-Umschau 44, 187.
Scott, M.L. (1981). Feedstuffs 53(8), 59.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Seymour, W.M. (1999). In Proc. “Tri-state Dairy Nutrition Conference,” p. 43,
Grand Wayne Center, Fort Wayne, Indiana.
Shiau, S., and Chin, Y. (1998). J. Nutr. 128, 2494.
Simmins, P.H., and Brooks, P.H. (1983). Vet. Rec. 112, 425.
Simmins, P.H., and Brooks, P.H. (1988). Vet. Rec. 122, 431.
Stout, F.M., Adair, J., and Oldfield, J.E. (1966). Nutr. Fur. News 38, 13.
Sunde, M.L., Cravens, W.W., Elvehjem, C.A., and Halpin, J.G. (1950). Poult.
Sci. 29, 10.
Sydenstricker, V.P., Singal, S.A., Briggs, A.P., DeVaughn, N.M., and Isbell, H.
(1942). JAMA 118, 1199.
Tagwerker, F. (1974). Roche Information Service Bulletin No. 1494. HoffmannLa Roche & Co., Ltd., Basel, Switzerland.
Tagwerker, R.J. (1983). Feed Int. 4, 22.
Terroine, T. (1960). Vitam. Horm. 18, 1.
Waisman, H.A., McCall, K.B., and Elvehjem, C.A. (1945). J. Nutr. 29, 1.
Walton, M.J., Cowey, C.B., and Andron, J.W. (1984). Aquaculture 37, 21.
Watanabe, T. (1993). J. Nutr. 123, 2101.
Watanabe, T., Dakshinamurti, K., and Persaud, T.V.N. (1995). J. Nutr. 125,
2114.
Watkins, B.A. (1989). Brit. J. Nutr. 61, 99.
Watkins, B.A., and Kratzer, F.H. (1987a). Poult. Sci. 66, 1818.
Watkins, B.A., and Kratzer, F.H. (1987b). Poult. Sci. 66, 306.
Watkins, B.A., Bain, S.D., and Newbrey, J.W. (1989). Calcif. Tissue Int. 46, 41.
Watkins, K.L., Southern, L.L., and Miller, J.E. (1991). J. Anim. Sci. 69, 201.
Wehr, N.B., Adair, J., and Oldfield, J.E. (1980). J. Anim. Sci. 50, 877.
Wellenberg, G.J., and Banks, J.N. (1993). J. Sci. Food Agric. 63, 1.
Whitehead, C.C. (1978). In Handbook Series in Nutrition and Food, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.),Vol. 2, p. 65. CRC Press, Boca
Raton, Florida.
Whitehead, C.C. (1988). Broiler Industry 51(9), 60.
Whitehead, C.C., and Bannister, D.W. (1978). Br. J. Nutr. 39, 547.
Whitehead, C.C., Armstrong, J.A., and Waddington, D. (1982). Br. J. Nutr. 48,
81.
Whitehead, C.C., Bannister, D.W., and D’Mello, J.P.F. (1980). Res. Vet. Sci. 29,
126.
Wiese, A.C., Johnson, B.C., and Nevens, W.B. (1946). Proc. Soc. Exp. Biol.
Med. 63, 521.
Zempleni, J., Green, G.M., Spannagel, A.W., and Mock, D.M. (1997). J. Nutr.
127, 1496.

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chapter twelve

FOLACIN

INTRODUCTION
Folacin and folate are generic terms used to describe folic acid andrelated compounds that exhibit the biological activity of folic acid. The
terms folacin, folate, and folic acid will be used interchangeably. A number of researchers in both developed and developing countries have reported a high incidence of folacin deficiency in pregnant women. It has
been estimated that up to one-third of all pregnant women in the world
may experience folacin deficiency of varying severity (Rothman, 1970).
Because of their rapid growth rate, cancer cells have an exceptionallyhigh folacin requirement. Therefore, drugs that inhibit folacin-requiring
enzymes are widely used in medicine for cancer chemotherapy. Megaloblastic anemia of pregnancy, resulting from low folacin intakes, is associated with poverty and poor diet selection. While folacin deficiency is
extremely common in women 16 to 40 years of age because of the effects of pregnancy and lactation, it is rare in men younger than 60 yearsof age. After age 60, folacin deficiency is equally high in both men andwomen.
Recent research has shown that adequate serum folacin not only
corrects megaloblastic anemia but prevents a number of life threatening
diseases, including cardiovascular disease, neural tube defects (e.g.,spina bifida and anencephaly), and risk of colorectal and other forms ofcancer (Glynn and Albanes, 1994; Bower, 1995; Brattström, 1996). In
relation to cardiovascular disease, moderate hyperhomocysteinemia is arisk factor. Supplementation with folacin and vitamin B12 reduces elevated plasma homocysteine (Brattström, 1996).
For animals, folacin needs are met principally by dietary sources andto some extent by intestinal bacterial synthesis. Different species vary in
479

12

C H A P T E R T W E LV E

ability to utilize microbial intestinal synthesis as a source of folacin.
Poultry and swine need supplemental folacin under certain conditions.
Folacin supplementation is most important when poultry and swine receive diets containing sulfa drugs and grains contaminated with toxinproducing molds. To maximize reproductive efficiency, gestating swine
often receive supplemental folacin.

HISTORY
In the 1930s and early 1940s, a number of active substances were
described that were effective against nutritional deficiencies in humans,certain animals, and bacteria. Factors that later were found to be folacin-related include vitamin M, factor U, vitamin Bc, Bc conjugate, Lactobacillus casei factor, Streptococcus lactis R. (SLR) factor, folic acid,citrovorum factor, and others. The early history of folacin is discussed
by Scott et al. (1982), Blakely and Benkovic (1984), and Loosli (1991).
Willis in 1931 demonstrated a factor from yeast that was active in treating a tropical macrocytic anemia seen in women of India. An anemiapreventive factor for monkeys was found in yeast or liver extracts anddesignated vitamin M in 1935 by Day and associates. In 1939 Hogan
and Parrot prevented anemia in chicks with a factor in liver called Bc. In
the late 1930s, factors needed for growth and anemia prevention inpoultry were referred to as Bc, factor U, or factor R. In 1940 Snell andcoworkers found a growth factor for L. casei, and in the same year, agrowth factor for S. lacti was found in spinach. The growth factor had
been isolated from 4 tons of spinach leaves. The isolated factor wascalled folic acid from folium, the Latin word for leaf.
Folic acid was isolated from the liver with the structure and synthesis accomplished by the Lederly group, who named it pteroylglutamic
acid on the basis of chemical structure. Active forms were found to contain a formyl group or a methyl group attached to the number 5 nitrogen of the pteridine nucleus.
Confusion existed in the 1940s concerning the identity of these various factors because both L. casei factor and folic acid were active forboth microorganisms and animals, whereas vitamin M, factor R, and vitamin Bc were active for monkeys and chicks but not microorganisms.
This was resolved when studies showed that folacin exists in nature inboth free and bound (additional glutamic acid molecules) forms. Incubating vitamin M with rat or chick liver enzymes markedly increased activity for L. casei and S. faecalis. Folacin conjugases (enzymes that hydrolyze polyglutamates to monoglytamyl forms) were also shown to480

FOLACIN

occur in hog kidney and chick pancreas. These studies, therefore,
showed that most species except microorganisms are able to utilize
bound forms.
Taking advantage of the high folacin requirement of cancer cells,
during the 1950s, two drugs, methotrexate and 5-fluorouracil, were
found to be useful in cancer chemotherapy. Both of these drugs are powerful inhibitors of folacin-requiring enzymes and are widely used in medicine. In the 1960s Herbert, of Boston College, clarified biochemical andhematological changes associated with a low folacin diet on himself andestablished progressive abnormalities as the deficiency intensified. In the
1970s the widespread world incidence of megaloblastic anemia was established, and in the 1980s and 1990s, the importance of folacin for reducing the incidence of neural tube defects, cardiovascular disease, andcancer began to emerge.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Folacin is the group name used to distinguish naturally occurring
compounds of this class; the pure substance is designated pteroylmonoglutamic acid. The chemical structure of folacin (pteroylglutamicacid) is shown in Fig. 12.1. Its chemical structure contains three distinct
parts. Reading from right to left, this compound consists of glutamic
acid, ρ-aminobenzoic acid (PABA), and a pteridine nucleus, the last two
making up pteroic acid. Thus the name pteroylglutamic acid was suggested. The PABA portion of the vitamin structure was once thought tobe a vitamin. If the folacin requirement is met, there is no need to add
PABA to the diet (see Chapter 17).
Much of the folacin in natural feedstuffs is conjugated with a varying number of extra glutamic acid molecules. Folacin as pteroyloligo-γL-glutamates (PteGlun) is generally from one to nine glutamates long,with n indicating the number of glutamyl residues. Polyglutamate
forms—usually of 3 to 7 glutamyl residues linked by peptide bonds—of
folacin are the natural coenzymes that are most abundant in every tissue
examined (Wagner, 1984). The conjugated forms with two or more glutamic acid residues are joined by γ-glutamyl linkages to the single glutamic acid moiety of the vitamin. Synthetic folacin, however, is in themonoglutamate form.
It has been concluded that there are more biologically active formsof folacin than any other known vitamin. Naturally occurringpteroylpolyglutamates constitute a large family of closely related com481

C H A P T E R T W E LV E

Fig. 12.1 Structures of folacin compounds. (R is one or more glutamic acidmolecules.)pounds arising from modifications of the three parts of the parent compound pteroylglutamic acid. Changes in the state of reduction of thepteridine moiety, addition of various kinds of one-carbon substituents,
and addition of glutamic acid residues lead to a wide array of compounds. Baugh and Krumdieck (1971), on the basis of the three known
states of reduction of the pyrazine ring, the six different one-carbon substituents that may occur at N-5 and/or N-10, and assuming that thepolyglutamyl chain would have no more than seven glutamyl residues,

482

FOLACIN

calculated that the theoretical number of folacins approached 150.
However, this figure includes compounds that have never been identified
in natural materials. Since it is clear now that the polyglutamyl chain
reaches at least 8 or 9 residues in animal tissues (and as many as 12 inbacterial cells), the number of folacin compounds that might be expected to occur in animal tissues still approaches 100 compounds. The
active forms of folacin contain a formyl group or a methyl group attached to the number 5 or number 10 nitrogens of the compound, or amethylene group between nitrogens 5 and 10. Tetrahydrofolic acid is theprincipal coenzyme form, while the main storage form is 5-methyltetrahydrofolic acid (Fig. 12.1).
Folacin is a yellowish-orange crystalline powder, tasteless and odorless, and insoluble in alcohol, ether, and other organic solvents. It isslightly soluble in hot water in the acid form but quite soluble in the salt
form. It is fairly stable to air and heat in neutral and alkaline solution,
but unstable in acid solution. From 70 to 100% of folacin activity is destroyed on autoclaving at pH 1 (O’Dell and Hogan, 1943). Large losses
in food folate can occur during food preparation such as heating, particularly under oxidative conditions (Gregory, 1989). Oxidation of reduced folates usually results in cleavage products lacking vitamin activity. Folacin is readily degraded by light and ultraviolet radiation.
Cooking can considerably reduce folacin food content (see Natural
Sources).
A great variety of folate analogs have been prepared, mainly for the
purposes of anticancer (e.g., methotrexate and 5-fluorouracil) and antitumoral therapy (Brody, 1991). Since folacin deficiency is more detrimental to cells that are rapidly growing, these antagonists are used as
potent antibacterial and antitumoral agents. Folacin antagonists can actby (1) blocking conversion of pteroylmonoglutamic acid to tetrahydrofolic acid by binding to dihydrofolic acid reductase or (2) blocking thetransfer of single-carbon units from tetrahydrofolic acid to acceptors,
such as in synthesis of methionine or purines (Scott et al., 1982). The folacin enzyme dihydrofolate reductase is the usual target of these antifolacin drugs. Methotrexate binds extremely tightly to dihydrofolate reductase. This drug is used both as an anticancer agent and (in low doses)
as a treatment for a number of nonmalignant disorders, such as psoriasis and rheumatoid arthritis.
Sulfonamides, though not folacin analogs, are analogs of the folacin
biosynthetic intermediate PABA and are widely used as antibacterial
agents (Brown, 1962). By competing with PABA, sulfonamides prevent

483

C H A P T E R T W E LV E

folacin synthesis so the microorganisms cannot multiply, with the resultthat an important source of folacin to the animal is reduced or eliminated. The anticonvulsant phenytoin is antagonistic to folacin, inhibiting the formation of polyglutamyl folates in rat liver (Carl et al., 1997).

ANALYTICAL PROCEDURES
Determination of folacin in biological materials is a difficult analytical problem because of the existence of a number of folacin complexes
that exhibit the vitamin activity. High sensitivity is essential in methodsof analysis because of the low concentration of folacin in foods and feedstuffs. Both microbiological and chemical methods have been employed
to identify folacin and its derivatives. Because folacin in natural feedstuffs is conjugated with varying numbers of extra glutamic acid molecules, the bound forms must be freed to be active for the assay microorganisms. Conjugase enzymes, widely distributed in animal tissues, arecapable of releasing free folacin. Typically, conjugases from the pancreas, liver, or kidney are used to reduce the conjugate to the monoglutamate form.
Folacin can be detected by microbiological assays (O’Broin and
Kelleher, 1992). The microbiological assay can use L. casei, which responds to all the common monoglutamate forms of folacin, and a differential assay using Enterococcus hirae (formerly known as S. faecalis)and L. citrovorum. A radiometric microbiological method has also beenreported, as well as a fluorometric procedure (Cruces-Blanco et al.,1994). For a biological assay of feeds, the chick is the preferred animal.
Although growth may not be a specific indicator of available dietary folacin, plasma and liver folacin are highly sensitive criteria.
Various methods have been developed for separation of folacins by
high-pressure liquid chromatography (HPLC). Separations of monoglutamyl folacins have been accomplished by anion-exchange, paired-ion
reverse-phase, and conventional reverse-phase HPLC methods (Gregory
et al., 1984; Selhub, 1989). Quantitation of the major folacin compounds in biological materials has been reported by Gregory et al.
(1984) to combine fluorometric determination with HPLC. Methods
have been developed for simultaneous measurement of one-carbon andpolyglutamate derivatives of folacin using enzymatic interconversions offolates and ternary complexes separated by isoelectric focusing (Carland Smith, 1995).

484

FOLACIN

METABOLISM
Digestion, Absorption, and Transport
Polyglutamate forms are digested via hydrolysis to pteroylmonoglutamate prior to transport across the intestinal mucosa. Intraluminal
polyglutamate hydrolysis is catalyzed by a conjugase intestinal enzyme
found in the brush border. This brush border pteroylpolyglutamate hydrolase (γ-carboxy peptidase) is an exopeptidase that cleaves the polyglutamyl chain one residue at a time starting from the carboxyl end. It has
a pH optimum near neutrality and is activated by zinc (Chandler et al.,1986). In humans, a zinc deficiency resulted in a decreased intestinal hydrolysis of pteroylpolyglutamate (Tamura et al., 1978). Conjugase activity is widely distributed in the mucosa of the proximal small intestine,
both intracellularly and in association with the brush border. Conjugase
activities have also been found in bile, pancreatic juice, kidney, and liver.
Conjugase activity is reduced by nutritional zinc deficiency, chronic consumption of alcohol, and exposure to naturally occurring inhibitors infoods.
Pteroylmonoglutamate is absorbed predominantly in the jejunum,with lesser amounts in the duodenum, by a Na+-coupled carrier-mediated process. Folacin is also absorbed passively, presumably by diffusion; this mechanism accounts for 20 to 30% of folacin absorption, regardless of folate concentration.
Dietary folates, after hydrolysis and absorption from the intestine,
are transported in plasma as monoglutamate derivatives, with only limited methylation (5-methyltetrahydrofolate). Folacin taken up by theliver is converted primarily to 5-methyltetrahydrofolate and 10-formyltetrahydrofolate and then transported to the peripheral tissues. The
monoglutamate derivatives are then taken up by cells in tissues by specific transport systems. There, the pteroylpolyglutamates—the major folacin form in cells—are built up again in stepwise fashion by the enzyme
folate polyglutamate synthetase. Polyglutamation traps folates inside
cells at concentrations one to two orders of magnitude greater thanthose of extracellular fluids. Polyglutamates serve to keep folacin within
the cells since only the monoglutamate forms are transported acrossmembranes, and only the monoglutamates are found in plasma andurine (Wagner, 1995). Folacin enzymes are compartmentalized between
the cytosol and the mitochondria. Almost all the folate in the cell is distributed equally between the two compartments. There are also mito485

C H A P T E R T W E LV E

chondrial and cytosolic isoforms of the same enzymes (Wagner, 1996).
Studies showed that about 79 to 88% of labeled folacin is absorbed,
and that absorption is rapid since serum concentrations usually peak
about 2 hours after ingestion. Using in vitro studies, Wagonfeld et al.
(1975) showed that the rate-limiting step in folacin absorption is transport of the monoglutamyl folacin into the mesenteric circulation rather
than hydrolysis of polyglutamyl folacins, which is very rapid. However,in cases where the conjugase is strongly inhibited, absorption of polyglutamyl folacins may be reduced. Low availability in orange juice is dueto the inhibition of conjugase activity by the low pH. Therefore, drugs
that drastically alter intraluminal pH and/or inhibit conjugase activity
will also decrease folacin absorption.
Specific folate-binding proteins (FBPs) that bind folacin monoglutamates and polyglutamates are known to exist in many tissues and body
fluids, including liver, kidney, small intestinal brush border membranes,
leukemic granulocytes, blood serum, allantoic fluid, and milk (Tani andIwai, 1984; Shoda et al., 1990; Muldoon et al., 1996; Vallet et al.,1999). The physiological roles of these FBPs are not completely known,
but it has been suggested that they play a role in folacin transport analogous to the intrinsic factor in the absorption of vitamin B12. The FBPs
may play a role in tissue storage of folacin by protecting the polyglutamate derivatives from the action of degradative or hydrolytic enzymes(Brody, 1991). The FBPs (also referred to as folate receptors) are abundant in the kidney proximal tubule and are involved with reabsorptionof 5-methyl THF4 (Muldoon et al., 1996). Tani and Iwai (1984) reported that FBPs of bovine milk affected bioavailability of folacin invivo by resulting in a more uniform absorption throughout the small intestine and reduced loss of 5-methyltetrahydrofolacin in the urine.

Storage
Folacin is widely distributed in tissues largely in the conjugatedpolyglutamate forms. Normal body stores in humans have been estimated at 5 to 10 mg, with approximately half in the liver (Brody, 1991).
Well-nourished adults may have stores that can meet normal body requirements for up to 4 or 5 months (Baker and DeMaeyer, 1979). Body
folacin stores are small at birth and are rapidly depleted, particularly insmall premature infants. In vitamin B12 deficiency, there are defects inthe conversion of pteroylmonoglutamates to polyglutamate forms thatlead to a decreased tissue ability to retain intracellular folacin. Vitamin
B12 deficiency has been shown to lead to functional folacin deficiency
even when folacin intake and absorption are normal. Giugliani et al.
486

FOLACIN(1985) reported a significantly higher serum folacin concentration forpatients receiving vitamin B12 supplementation during pregnancy.

Excretion
Urinary excretion of folacin represents a small fraction of total excretion (e.g., < 1% of total body stores). Fecal folacin concentrations arequite high, often higher than intake, representing not only undigested folacin but, more important, the considerable bacterial synthesis of the vitamin in the intestine. In tracer studies for the first 24 hours after thedose, much of the dose in urine is as intact folacins. After longer time
periods, all the excreted label was in degradative products, primarily
pteridines and acetaminobenzoylglutamate (Murphy et al., 1976). Over
half of labeled dietary folacin is excreted in feces. This is expected, asbile contains high levels of folacin due to enterohepatic circulation, withmost of bile folacin reabsorbed in the intestine.

FUNCTIONS
Folacin, in the form 5,6,7,8-tetrahydrofolic acid, is indispensable intransfer of single-carbon units in various reactions, such as those occurring in the biosynthesis of lipids, proteins, nucleic acid derivatives, hormones, and neurotransmitters—a role analogous to that of pantothenic
acid in the transfer of two-carbon units. The one-carbon units can beformyl, formimino, methylene, or methyl groups. The major in vivo
pathway providing methyl groups involves transfer of a one-carbon unit
from serine to tetrahydrofolate to form 5,10-methylenetetrahydrofolate,which is subsequently reduced to 5-methyltetrahydrofolate. Methyltetrahydrofolate then supplies methyl groups to remethylate homocysteinein the activated methyl cycle, providing methionine for synthesis of theimportant methyl donor agent S-adenosylmethionine (Krumdieck, 1990;
Jacob et al., 1994). Some biosynthetic relationships of one-carbon units
are shown in Fig. 12.2. These one-carbon units are generated primarily
during amino acid metabolism and are used in the metabolic interconversions of amino acids and in the biosynthesis of the purine and pyrimidine components of nucleic acids that are needed for cell division. The
important physiological function of tetrahydrofolate (THF) consists ofbinding the C1 units to the vitamin molecule and thus transforming them
to “active formic acid” or “active formaldehyde,” so that these are interconvertible by reduction or oxidation and transferable to appropriate
acceptors.
Ability of the vitamin as a coenzyme is governed by three factors: (1)487

C H A P T E R T W E LV E

Fig. 12.2 Folacin metabolism requiring one-carbon units. (Modified from Scottet al., 1982.)

the nature of the carbon-containing substituent attached at either the N5 or the N-10 position, or bridging them; (2) the state of oxidation or
reduction of the pyrazine ring; and (3) the number of glutamic acid units
attached in gamma linkage to the glutamate of pteroylglutamic acid.
Certain polyglutamates may serve as cofactors for one enzyme while inhibiting another (White et al., 1976).
Folacin polyglutamates work at least as well as or better than thecorresponding monoglutamate forms in every enzyme system examined(Wagner, 1995). It is now accepted that the pteroylpolyglutamates are
the acceptors and donors of one-carbon units in amino acid and nucleotide metabolism, while the monoglutamate is merely a transport
form.
Glutamate chain length of folacin polyglutamate may affect metabolism of one-carbon units (Foo and Shane, 1982). In hamster ovary cellswith normal extracellular methionine, polyglutamate chain lengths werelonger, with high levels of octaglutamates, nonaglutamates, and evendecaglutamates occurring, while with suboptimal levels of methionine,
shorter chain lengths were found. This observed phenomenon of a profound effect of extracellular methionine concentration on glutamate
488

FOLACIN

chain elongation may be interpreted to mean that there is a regulatory
action of polyglutamate chain length on one-carbon metabolism.
Specific reactions involving single-carbon transfer by folacin compounds are (1) purine and pyrimidine synthesis, (2) interconversion ofserine and glycine, (3) glycine-α-carbon as a source of C1 units for many
syntheses, (4) histidine degradation, and (5) synthesis of methyl groups
for such compounds as methionine, choline, and thymine.
Purine bases (adenine and guanine) as well as thymine are constituents of nucleic acids, and with folacin deficiency, there is a reduction in the biosynthesis of nucleic acids essential for cell formation andfunction. Hence, deficiency of the vitamin leads to impaired cell division
and alterations of protein synthesis; these effects are most noticeable inrapidly growing tissues. In absence of adequate nucleoproteins, normal
maturation of primordial red blood cells does not take place, andhematopoiesis is inhibited at the megaloblast stage. As a result of thismegaloblastic arrest of normal red blood cell maturation in bone marrow, a typical peripheral blood picture results that is characterized bymacrocytic anemia. White blood cell formation is also affected, resulting in thrombopenia, leukopenia, and old, multilobed neutrophils.
In folacin deficiency, formiminoglutamic acid (FIGLU), formed as
an intermediate in degradation of histidine, can no longer be transformed completely into glutamate and formiminotetrahydrofolic acid,and is therefore excreted in urine. This excretion is suitable as a biochemical criterion for diagnosis of folacin deficiency, appearing at an
early stage of deficiency.
Vitamin B12 is also closely associated with the progress of the folacin-dependent reactions of intermediary metabolism (Savage and Lindenbaum, 1995). Vitamin B12 has two main effects in facilitating folacin:(1) Vitamin B12 regulates the proportion of methyl to nonmethyl
tetrahydrofolates according to the methyl trap theory, and (2) vitamin
B12 is necessary for transport of methyl-THF across the cell membraneand promotes folacin retention by tissues. According to the methyl trap
concept (Herbert and Zalusky, 1962), vitamin B12 deficiency decreases
the formation of methionine from homocysteine and methyl-THF by theB12-dependent methionine synthetase (see Chapter 13, Functions). This
results in an increase in methyl-THF and a decrease in THF, which is theactive coenzyme form that functions in the degradation of FIGLU andformate.
Vitamin B12 is necessary in the reduction of one-carbon compoundsof the oxidation stage of formate and formaldehyde, and in this way, it
participates, with folacin, in biosynthesis of labile methyl groups. Fo489

C H A P T E R T W E LV E

lacin is also essentially involved in all these reactions of labile methylgroups. The metabolism of labile methyl groups plays an important role
for the body in the biosynthesis of methionine from homocysteine andof choline from ethanolamine. Folacin has a sparing effect on requirements of choline (the importance of both folacin and vitamin B12 on synthesis of choline is discussed in Chapter 14).
Folacin is needed to maintain the immune system; the blastogenic
response of T lymphocytes to certain mitogens is decreased in folacindeficient humans and animals, and the thymus is preferentially altered(Dhur et al., 1991). The effects of folacin deficiency upon humoral immunity have been more thoroughly investigated in animals than in humans, and the antibody responses to several antigens have been shownto decrease. As de novo synthesis of methyl groups requires the participation of folacin coenzymes, the effect of folacin deficiency on pancreatic exocrine function was examined in rats (Balaghi and Wagner, 1992;
Balaghi et al., 1993). Pancreatic secretion was significantly reduced in
the deficient group compared with the pair-fed control groups after 5
weeks. The results indicate that severe folacin deficiency impairs pancreatic exocrine function. The ratio of S-adenosylmethionine to S-adenosylhomocysteine was rapidly reduced in the deficient pancreas. The pancreas of deficient rats had more immature secretory granules, and theducts were devoid of secreted material.

REQUIREMENTS
Various animal species differ markedly in their requirements for folacin. Because of microbial synthesis in their digestive tracts, ruminants
have no dietary requirement for folacin. Only young ruminants that do
not have a fully developed rumen would be expected to require a dietary
source. Folacin requirements for monogastric species would be dependent on degree of intestinal folacin synthesis and utilization by the animal(Rong et al., 1991). Animals that practice coprophagy would also havea lower dietary need for folacin, as feces is a rich source of the vitamin(Abad and Gregory, 1987). Many species apparently do not require dietary folacin because of their ability to utilize microbial intestinal synthesis (Rong et al., 1991).
Poultry, swine, guinea pigs, and primates (including humans) do develop deficiencies on low dietary folacin. Even though deficiencies canbe produced with special diets, corn, soybean meal, and other common
feedstuffs in a practical poultry diet usually provide ample folacin undermost conditions (Scott et al., 1982). However, marginal inadequacies
490

FOLACIN

have been produced in chicks fed corn-soybean meal diets (Pesti et al.,1991).
Self-synthesis of folacin is dependent on dietary composition. For
poultry, some research has indicated higher folacin requirements forvery high protein diets, or when sucrose was the only source of carbohydrates (Scott et al., 1982). Keagy and Oace (1984) reported that dietary fiber had an effect on folacin utilization; xylan, wheat bran, andbeans stimulated folacin synthesis in the rat, reflected as higher fecal andliver folacin. For humans it was concluded that milk type differentiallyaffects intestinal folacin biosynthesis and the superior folacin availability from human (versus cow and goat) milk-containing diets is due inpart to enhanced intestinal biosynthesis of folacin (Semchuk et al.,1994).
The levels of antibacterials added to the feed will affect microbial
synthesis of folacin. Sulfa drugs, which are commonly added to livestock
diets, are folacin antagonists (see Deficiency). In the chicken, sulfa drugs
have been shown to increase the requirement (Scott et al., 1982). Moldy
feeds (e.g., aflatoxins) have also been shown to contain antagonists that
inhibit microbial intestinal synthesis in swine (Purser, 1981).
Folacin requirements are dependent on the form in which it is fed
and concentrations and interrelationships of other nutrients. Deficiencies of choline, vitamin B12, iron, and vitamin C all have an effect on folacin needs. Although most folacin in poultry feedstuffs is present inconjugated form, the young chick is fully capable of utilizing it. On thecontrary, Baker et al. (1978) reported that human patients over 60 yearsof age utilized conjugated forms of folacin much less efficiently thanmonoglutamates.
Folacin requirements are related to type and level of production.
Growth rate, age, and pregnancy influence folacin requirements. The requirement decreases with age because diminished growth rate reduces
the need for DNA synthesis. Increased catabolism of folacin is a featureof pregnancy. Studies with both rats (McNulty et al., 1993) and humans(McPartlin et al., 1993) demonstrated an enhanced folacin catabolismthat was a feature of pregnancy per se and not simply due to increased
weight. In poultry the folacin requirement for egg hatchability is higher
than that for production (NRC, 1994). Taylor (1947) reported that 0.12
mg of folacin per kilogram of diet was satisfactory for egg production,
but higher levels were required for good hatchability. Table 12.1 summarizes the folacin requirements for various livestock species and humans; a more complete listing is given in the appendix, Table A1.
The current Recommended Dietary Allowances (RDAs) for folates491

C H A P T E R T W E LV E
■ Table 12.1

Folacin Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle

Adult
Calf
Adult
Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100-g intake)
Broilers, 0–6 weeks
Broilers, 6–8 weeks
All classes
Growing, 0–8 weeks
Growing, 8–16 weeks
Breeding hens
Adult
Growing-finishing
Gestating-lactating
Adult
Adult
Adult
Growing
Growing
Growing
All classes
CatfishPacific salmon
Rainbow trout
All classes
Infants
Children
AdultsMicrobial synthesis
0.1 ppm milk replacer
Microbial synthesis
0.55 mg/kg
0.25 mg/kg
0.25 mg/kg
0.55 mg/kg
0.50 mg/kg
1.0 mg/kg
1.0 mg/kg
0.8 mg/kg
1.0 mg/kg
Microbial synthesis
0.3 mg/kg
1.30 mg/kg
Microbial synthesis
Microbial synthesis
0.8 mg/kg
0.5 mg/kg
0.2 mg/kg
0.2 mg/kg
0.6 mg/kg
1.5 mg/kg
2–10 mg/kg
1 mg/kg
1.0 mg/kg
25–30 μg/day
50–100 μg/day
150–400 μg/day

NRC (1984)
NRC (1984a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1981b)
NRC (1986)
NRC (1982a)
NRC (1982a)
NRC (1985a)
NRC (1978a)
NRC (1993)
NRC (1993)
NRC (1993)
NRC (1978a)
RDA (1989)
RDA (1989)
RDA (1989)

Dairy cattle
ChickenJapanese quail
Turkey
Sheep
Swine
Horse
Goat
Cat
Mink
Fox
Dog
HamsterFish
Rat
HumanbaExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Requirements established for some species,
while only suggested for others. Human data are expressed as μg/day.
bA number of nutritionists believe the RDA (1989) was ultraconservative in recommending folacin allowances. Research findings would suggest the 1980 RDA to havemore realistic recommendations as follows: Infants (30–45 μg/day), children (100–300
μg/day), and adults (400–800 μg/day).

are set at 200 μg/day for adult males and 180 μg/day for adult females(RDA, 1989). This represents a significant reduction from the previously
recommended (RDA, 1980) amount of 400 μg/day for adult males andfemales (Bailey, 1995). The later RDA also reduced the recommended
requirement for children and infants (Table 12.1). Many nutritionists
feel that the 1989 RDA committee was ultraconservative in suggesting
folacin recommendations and prefer the 1980 RDA values. Also, research associating folacin deficiency with hyperhomocysteinemia and increased heart disease risk (Kang et al., 1992), cervical dysplasia (Butterworth et al., 1992), bronchial metaplasia (Heimburger et al., 1988), and492

FOLACIN

neural tube defects (Centers for Disease Control, 1992), and the wideuse of folate-antagonist medications, clearly demonstrate an increased
need for higher levels of dietary folacin (see Supplementation). As an example, the U.S. Public Health Service issued a recommendation that all
U.S. women of childbearing age capable of becoming pregnant should
consume 0.4 mg (400 μg) of folacin per day for the purpose of reducing
their risk of a pregnancy affected with spina bifida or other neural tube
defects (Centers for Disease Control, 1992).
In pregnancy and lactation, folate requirements clearly increase. The
burden of lactation on maternal folacin reserves is estimated to be 20 to50 μg/day, varying with the folacin content and volume of milk (Matothet al., 1965). This estimate, based on production of 850 cc of average
milk folacin content, should be doubled to meet the needs of mothers
producing milk with high folate content. An additional 100 μg of dietary folacin should provide for the absorption of the excess need. The
recommended daily allowance (RDA, 1980) for folacin is necessarily inexcess of the minimum daily requirement to provide a margin of safety
and allow for losses in preparation of foods as well as the decreased
availability of food folacin as compared to folacin in the monoglutamate
form. The newer RDA (1989) has set 400 μg/day during pregnancy and280 μg/day during the first 6 months of lactation as a folacin recommendation. This likewise is low compared with the earlier RDA (1980)
recommendation. It should be noted that a study of 10 nonpregnant
women followed for 92 days in a metabolic unit arrived at a higher recommended intake of 300 μg/day to meet requirements and provide anallowance for storage (Sauberlich et al., 1987).
Smoking, alcohol consumption, antifolacin drugs (e.g., methotrexate), anticonvulsant drugs, oral contraceptives, and certain antibiotics(e.g., sulfasalazine) interfere with folacin activity and therefore increase
the body’s folacin requirements (Brody, 1991). Tobacco smoking is believed to represent a second model of localized folacin deficiency often
associated with preneoplastic changes of the affected tissues(Krumdieck, 1990). Methotrexate lowers folacin status, whether used as
an anticancer agent or in low doses as a treatment for psoriasis andrheumatoid arthritis (Green and Jacobsen, 1995). Anticonvulsant drugs
have been found to reduce blood folacin and lead to problems such as
megaloblastic anemia (Porras-Tejero and Lluch-Fernandez, 1993). As
many as 19% of the users of oral contraceptive agents showed cytological abnormalities that, although not associated with systemic evidenceof folacin or vitamin B12 deficiency, were corrected by the administration of oral folic acid (Krumdieck, 1990).
493

C H A P T E R T W E LV E

NATURAL SOURCES
Folacin is widely distributed in nature, almost exclusively as THF
acid derivatives; the stable ones have a methyl or formyl group in the 5γ position and generally possess three or more glutamic acid residues inglutamyl linkages. Only limited amounts of free folacin occur in natural
products, with most feed sources containing predominantly polyglutamyl folacin. However, in seeds or fruit, which presumably store the vitamin, a considerable amount is present as a monoglutamate. More thana third of the folates in orange juice are present as monoglutamates andnearly half as pentaglutamates. A high proportion of monoglutamate
forms of folacin are found in milk and soybeans. Much of the folacin in
milk is available in the monoglutamate form, which is necessary for absorption by the newborn (Wagner, 1984). The predominant form of folacin in vegetables such as spinach, asparagus, broccoli, lettuce, yeast,
rice, and peas is the N-10 formyl derivative. Folates in tissues such as
liver, kidney, and red blood cells are predominantly pentaglutamates.
Folacin is abundant in green leafy materials and organ meats. Soybeans, other beans, nuts, some animal products, and citrus fruits aregood sources. Cereal grains, milk, and eggs are generally poor sourcesof the vitamin. Any animal or human diet without green leafy materials
or animal protein, especially organ meats, is likely to be low in folacin.
Human diets may be particularly low during the winter season, whengreen vegetables and citrus fruits are less plentiful. The folacin content
of typical foods and feedstuffs is shown in Table 12.2.
Folacin bioavailability of beef liver, lima beans, peas, spinach,mushrooms, collards, and wheat germ was found to generally exceed
70% (Clifford et al., 1990). Bioavailability of monoglutamate folacin
was substantially greater than polyglutamyl forms (Clifford et al., 1991
Gregory et al., 1991). The availability of food folacins may range from30 to 80% that of the monoglutamate form, being generally less well
utilized from plant-derived foods than from animal products. Deficiencies of both iron and vitamin C in humans is associated with impaired
utilization of dietary folacin (see Deficiency, Assessment of Status).
Bioavailability of orally administered 5-methyl folacin and 5-formyl folacin compared to folacin were found to be equal for rats (Bhandari and
Gregory, 1992).
A considerable loss of folacin (50 to 90%) occurs during cooking orprocessing of foods. Folacin is sensitive to light and heating, particularlyin acid solution. Under aerobic conditions, destruction of most folacin
494

FOLACIN■ Table 12.2

Typical Folacin Concentrations in Foods and Feedstuffs (ppm, dry basis)

Alfalfa meal, dehydrated
Asparagus, fresh
Bananas, fresh
Barley, grain
Blood meal, dehydrated
Brewer's grains
Broccoli, fresh
Broccoli (boiled), fresh
Cabbage, fresh
Cabbage (boiled), fresh
Carrot, roots
Coconut meal
Corn (maize), grain
Corn (maize), gluten meal
Cottonseed meal, solvent extracted
Fish meal, anchovy
Fish meal, menhaden
Linseed meal, solvent extracted
Liver, cattle

5.5
1.4
0.28
0.6
0.1
7.7
1.69
0.65
0.3
0.16
1.2
1.5
0.3
0.3
2.8
0.2
0.2
1.4
8.4

Milk, skim, cow's
Orange juice, fresh
Peanut meal, solvent extracted
Rice, branRice, grain
Rice, polished
Rye, grain
Sorghum, grain
Soybean meal
Soybean, seeds
Spinach, fresh
Spinach (boiled), fresh
Sugarcane molasses
Timothy hay, sun cured
Wheat, bran
Wheat, grain
Whey, cattle
Yeast, brewer's

0.7
1.4
0.7
2.4
0.4
0.2
0.7
0.2
0.7
3.9
1.93
0.91
0.1
2.3
1.6
0.5
0.9
10.3

Sources: Concentrations from NRC (1982a) and Brody (1991).

forms is significant with heating, with reduced folacins more stable infoods due to relatively anaerobic conditions and because folacin is protected from light (Brody 1991). Fresh cabbage and broccoli are goodsources of folacin and contain (wet-weight basis) 0.30 and 1.69 ppm, respectively. However, when these vegetables are boiled and the water isdiscarded, losses are considerable, reducing concentrations to 0.16 and0.65 ppm, respectively (Leichter et al., 1978). Although the cooking
losses of folacin in vegetables may be great, the bioavailability of the remaining folacin is quite high (Clifford et al., 1990): broccoli 80 to 90%
and cabbage 79%.

DEFICIENCY
Folacin deficiency has been produced experimentally in many animal species, with macrocytic anemia (megaloblastic anemia) andleukopenia (reduced number of white cells) being consistent findings.
Tissues that have a rapid rate of cell growth or tissue regeneration, suchas epithelial lining of the gastrointestinal tract, the epidermis, and bonemarrow, are principally affected (Hoffbrand, 1978).
For some animals, such as the chick, guinea pig, monkey, and pig,
the presence of adequate amounts of folacin in the diet is essential, anddeficiency signs can readily be induced by feeding a diet deficient in the495

C H A P T E R T W E LV E

vitamin. In other animals, such as the rat and dog, folacin produced by
the intestinal microflora is usually adequate to meet requirements. Consequently, deficiency signs do not develop unless an intestinal antiseptic
is also included in the diet to depress bacterial growth.

Effects of Deficiency
Ruminants
Folacin synthesis occurs in the rumen. However, young animals that
do not have a fully developed rumen would be expected to be folacin deficient. Draper and Johnson (1952) reported folacin deficiency in lambs
fed synthetic diets. The disease was characterized by leukopenia followed by diarrhea, pneumonia, and death. Folacin therapy promoted regeneration of white blood cells, and 0.39 mg/L of milk in control animal
diets prevented the deficiency. There was no indication of folacin deficiency in calves fed synthetic milk containing 52 mg of folacin per kilogram of liquid feed fed at 10% of live weight (Wiese et al., 1947). In
later studies, dairy veal calves were fed daily 0, 4, or 16 mg of folacin in
milk (Lévesque et al., 1993). Supplemental folacin increased serum andred blood cell folacin concentrations and reduced the rearing period toreach 180 kg without increasing feed intake.
Girard et al. (1994) studied whether ruminal folacin is affected bydietary folacin and if this response is modified by the diet. Concentrations of folacin in rumen contents were increased by supplemental folacin and by ingestion of concentrate compared with hay-based diets.
The efficiency of folacin and synthesis by rumen microflora andwhether this is adequate at weaning and later is not yet established. For
example, in an experiment in which supplemental folacin was administered to dairy heifers intramuscularly weekly during the first 4 monthsof life, average daily gain increased by 8% during the 5 weeks following
weaning (Dumoulin et al., 1991). This supplementation also increased
serum and hepatic folates, as well as blood hemoglobin and packed cellvolume. Girard et al. (1989a) observed that concentration of serum folacin of calves at 2 weeks of age is half that of 4-month-old heifers. Girard et al. (1992) fed supplemental folacin to both preruminant and ruminant calves in an attempt to maintain high serum folacin
concentrations. Serum folacin was increased in both groups receiving dietary folacin, but the amount needed to reach similar concentrations
was higher in ruminant than in preruminant calves. These observations
seem to confirm that during the weeks before and after weaning, thesupply of folates by the diet and the rumen microorganisms might not
496

FOLACIN

be optimum for dairy heifers and possibly other preruminants and ruminant livestock.
Girard et al. (1989b) evaluated serum folates in gestating and lactating dairy cows and found that serum folacin decreased by 40% from
2 months postpartum (around mating) to parturition. The synthesis offolacin by rumen microorganisms was not sufficient to prevent fluctuations of serum folacin during gestation and lactation in dairy cows.
Moreover, serum folacin can be increased by an intramuscular injectionof folacin at the end of gestation, but in cows during early lactation, thisdid not markedly affect serum and milk folacin concentrations. Girard
et al. (1995) injected folacin (160 mg) weekly to dairy cows 45 days after mating to 6 weeks after parturition. The supply of folates by the dietand the synthesis by ruminal microflora were found to be sufficient toprevent folacin deficiency in dairy cows and to maintain normal gestation and lactation, but not to achieve maximal production of milk andprotein in multiparous, but not primiparous, dairy cows during gestation and lactation (Girard et al., 1995; Girard and Matte, 1998).

Swine
Earlier, folacin deficiency in swine had only been produced by the simultaneous feeding of sulfa drugs, indicating that intestinal synthesis
was adequate to meet needs. Deficiencies were not observed when young
pigs were fed only purified diets or low natural diets alone (Johnson etal., 1948), indicating that intestinal synthesis was adequate to meetneeds. Feeding a purified diet containing 2% sulfasuxidine to weanlingpigs resulted in reduced gains and alopecia (Cartwright and Wintrobe,
1949). The pigs also developed a milk normochromic, normocytic anemia; in bone marrow there was a decrease in the ratio of leukocytes toerythrocytes and an increase in the number of immature nucleated red
blood cells. Positive response was obtained after supplementation with
folacin. Cunha et al. (1948) found that folacin was needed for normal
hematopoiesis with 8-week-old pigs fed a purified diet with sulfasuxidine for 21 weeks. A normocytic anemia resulted that was prevented by
folacin, whereas a more severe anemia was produced by using a crude
folacin antagonist. A combination of folacin and biotin was more effective than folacin alone in counteracting the anemia. Lindemann andKornegay (1986) reported that combination of the antibiotic mixture
ASP 250 (which includes chlortetracycline, sulfamethazine, and penicillin) and folacin to a corn-soybean meal diet increased gains and feed
consumption, with no effect of either used alone.
More severe deficiency signs that responded to folacin supplemen497

C H A P T E R T W E LV E

tation were induced by feeding diets containing a sulfonamide and a folacin antagonist (Welch et al., 1947). Under such circumstances, pigs became listless, had a reduced growth rate, and developed diarrhea. Hematological manifestations were severe macrocytic anemia, leukopenia
with a more marked reduction in the number of polymorphonucleocytes, and mild thrombocytopenia. Cartwright et al. (1952) reported a
combined folacin and vitamin B12 deficiency in pigs receiving a purified
soybean protein diet that included a folacin antagonist. Growth rate was
reduced, and macrocytic anemia, leukopenia, and neutropenia developed, with erythroid hyperplasia of the bone marrow. Folacin supplementation immediately resulted in a normal blood and bone marrow
picture, but growth was decreased, and the blood picture subsequently
relapsed.
In addition to sulfa drugs and other folacin antagonists, moldy feeds
can increase the need for the vitamin. In seven swine-feeding trials involving more than 1,000 pigs fed corn with mold infestation, additional
folacin increased growth rate up to 15% and improved feed efficiency
up to 9% (Purser, 1981). Folacin supplementation was of no value whennormal corn was fed. Improvements in growth rate for pigs receiving
840 ppb aflatoxin were obtained with addition of 2 ppm folacin (Lindemann et al., 1993).
Inadequate folacin has been associated with suboptimal reproductive performance of sows in some studies (Matte et al., 1984b; Lindemann and Kornegay, 1989; Tremblay et al., 1989; Lindemann, 1993;
Matte and Girard, 1999) but without affect in others (Matte et al.,
1993; Harper et al., 1994). In sows a dramatic decrease in serum folacin
concentrations was observed during early and mid-gestation (Matte etal., 1984a; Harper et al., 1994). Low folacin during gestation may be associated in part with embryonic mortality. In one trial, folacin was administered intramuscularly according to a schedule that maintained
serum folacin concentrations at approximately the same level between
weaning and 60 days of gestation (Matte et al., 1984b). Average live litter size was 12 piglets per litter for sows receiving folacin and flushingtreatments compared with 10.5 for sows without any treatment. In another study, addition of 5.0 mg of folacin per kilogram of diet improved
survival rate of fetuses during early gestation (62.2 versus 55.1%) compared with those not receiving folacin (Tremblay et al., 1989). Lindemann and Kornegay (1989) found the number of matings required per
female farrowing was lower with folacin supplements (1.07 versus 1.16
for controls). Matte et al. (1992) reported that growth of piglets and total litter weight from birth to 8 weeks of age increased linearly from the498

FOLACIN

level of folacin that had been provided to sows during the gestation diet.
There seems to be no major benefits to lactational supplementation,although it is effective in elevating sow serum folacin, milk folacin, andnursing pig serum folacin. The response in gestation of increased litter
size seems to be a result of improved embryo or fetal survival rather thanincreased ovulation, although the mechanism whereby survival rate isimproved is not yet understood (Lindemann, 1993). Gilts fed 4 mg of folacin per kilogram of feed 17 days before and 7 days after insemination
had an average embryo survival rate (based on ovulation rate) of 88.5
versus 62.9% for controls and 4.4 versus 22.4% degenerated embryos
for the treatments, respectively (Stancic et al., 1993). Additional research is required to determine if supplemental dietary folacin will reduce embryonic death loss under differing management systems to improve overall efficiency of production (i.e., to optimize pig survival andlitter size).

Poultry
Poultry are more susceptible to lack of folacin than other farm livestock, as deficiency can readily be produced by feeding a folacin-deficient diet. Folacin deficiency, as indicated by retarded growth and feed
efficiency, was produced in 15-day-old chicks fed corn-soybean meal diets (Pesti et al., 1991).
In folacin deficiency, megaloblastic arrest of erythrocyte formationin bone marrow causes severe macrocytic anemia as one of the firstsigns. Folacin deficiency in chicks is also characterized by poor growth,very poor feathering, anemic appearance, and perosis (Figs. 12.3 and12.4). The chicks become lethargic and feed intake declines. As anemia
develops, the comb becomes waxy white, and mucous membrane of themouth becomes pale (Siddons, 1978). Turkey poults fed a folacin-deficient diet show reduced growth rate and increased mortality (Fig. 12.5).
The birds develop a spastic type of cervical paralysis in which the neckis stiff and extended, but with only a moderate degree of anemia. Poultswith cervical paralysis will die within 2 days after the onset of thesesigns unless folacin is administered immediately (Scott et al., 1982). Erythrocytes of deficient birds tend to be larger in diameter, and their nuclei
are less dense than those of birds receiving supplementary folacin(Schweigert et al., 1948).
In chicks and turkeys, folacin deficiency also results in poor feather
development, with the shafts weak and brittle. Folacin, along with lysine
and iron, is required for feather pigmentation, as depigmentation occurs
in colored feathers during folacin deficiency.
499

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Fig. 12.3 Folacin-deficient bird on left with depigmentation and reduction in
growth (compare with normal bird on the right). (Courtesy of G.F. Combs,
Department of Poultry Science, University of Maryland.)Fig. 12.4 Folacin-deficient chick, with cervical paralysis, at 5
weeks of age. Note theweakened condition oflegs and the way thebird holds the left
wing. Folacin-deficientbird will shake the endof the wing, and thewhole bird will quiverat times. (Courtesy of
M.L. Sunde, Universityof Wisconsin.)

It appears that egg production is less affected by folacin deficiency
than the development of the chick or poult. Egg and poult weights weresignificantly increased when turkey hens received higher dietary folacin
and when eggs were injected with folacin (Robel, 1993). Inadequate folacin provided to the hen impairs the oviduct’s response to estrogen andthe ability to form albumen (NRC, 1994). Inadequate intake of folacin
by breeding hens results in poor hatchability and a marked increase in500

FOLACIN

Fig. 12.5 Folacin-deficient poult was hatched from a hen fed a diet low in folacin. (Courtesy of M.L. Sunde, University of Wisconsin.)embryonic mortality (Fig. 12.6), which occurs during the last days of incubation. A deformed beak and bending of the tibiotarsus are signs of
the embryonic deficiency. Chicks that successfully emerge are stunted
and have feathers that are poorly developed and abnormally pigmented(NRC, 1994).
Folacin deficiency has sometimes been associated with perosis, orslipped tendon. Pollard and Creek (1964) demonstrated histologicallythat the lesions of folacin-deficient bones and cartilage are different
from those produced by choline or manganese deficiencies. Abnormal
structure of the hyaline cartilage is found in folacin-deficient chicks, andossification is retarded. These disorders are not found in chicks deficientin choline or manganese, although bone deformities and slipped tendons
are found in both types of disorders. However, Bechtel (1964) claimed
that choline is effective in preventing perosis only when sufficient folacin
is present in the diet. Dietary choline content has been shown to affectthe chick’s requirement for folacin. When the diet contained adequatecholine, the folacin requirement was 0.46 mg/kg diet, but this increased
to 0.96 mg/kg diet when the diet was choline deficient (Young et al.,1955). Increasing the protein content of the diet has also been shown toincrease the incidence and severity of perosis in chicks receiving low levels of dietary folacin. It is suggested that this increased requirement forfolacin in high protein diets for poultry is a consequence of greater de501

C H A P T E R T W E LV E

Fig. 12.6 Folic acid deficiency. Abnormal embryo from an egg laidby a hen on a low-folacin diet. (Courtesy of
M.L. Sunde, Universityof Wisconsin.)mand for folacin in uric acid formation (Creek and Vasaitis, 1963).
Folacin appears to be necessary for cell mitosis. In the absence of folacin, oviduct growth is not increased in estrogen-treated chicks. The
production of water-soluble proteins (particularly the albumin fraction)in the hormone-stimulated oviduct is also greatly reduced, and theamino acid composition of these proteins is altered. The percentages ofarginine, leucine, serine, and tryptophan are decreased and those ofglycine and methionine increased (Siddons, 1978).

Horses
It has been shown that the horse synthesizes folacin in the intestinal
tract. Carroll et al. (1949) fed a diet containing less than 0.1 mg of folacin per kilogram of dry matter and found the following folacin concentrations in ingesta (mg/kg dry matter): duodenum, 0.9; ileum, 0.5; cecum, 3.0; anterior large colon, 4.7; and anterior small colon, 2.7.
Folacin synthesis may not be sufficient; Seckington et al. (1967) reported
a case of folacin deficiency in a 7-year-old gelding that had been receiving a diet lacking in fresh grass for many months. The folacin-deficientanimal had poor performance associated with low serum folacin. Administration of 20 mg of folacin dramatically increased performanceand elevated blood folacin. Roberts (1983) examined serum and erythrocyte folacin levels in Australian horses and found higher concentrations in those on pasture than in permanently stabled horses.

Other Animal Species
Folacin deficiency has been reported using only purified feed ingredients or only diets naturally low in the vitamin for a number of species,502

FOLACIN

including the dog, fox, guinea pig, hamster, mink, monkey, salmon, andtrout. For other species, including the rat, cat, carp, and catfish, folate
deficiencies are difficult to establish unless use is made of an intestinal
antiseptic (sulfa drug) or folate antagonist. For mice and certain otherspecies, care must be taken to prevent coprophagy if folacin deficiency
is to be produced on a diet without the aid of an antagonist or sulfa
drug.

DOGS AND CATS
Folacin deficiency was produced in cats by adding sulfonamides tosemipurified diets; the deficiency was characterized by weight loss, anemia (macrocytic tendencies), and leukopenia. Blood-clotting time wasincreased, and plasma iron concentrations were elevated (Carvalho da
Silva et al., 1955).
Folacin deficiency resulted in erratic appetite, decreased gain, water
exudate from eyes, glossitis, leukopenia, hypochromic anemia, and decreased antibody response to infectious canine hepatitis and canine distemper virus (NRC, 1985a). A positive response was obtained by subcutaneous injections of folacin. Serum folacin was increased in various
breeds of dogs that received dietary folacin (Davenport et al., 1994). A
fox terrier bitch with chronic ehrlichiosis exhibited a regenerative anemia and thrombopenia associated with marrow hypercellularity.
Dyserythropoiesis and dysthrombopoiesis were attributed to folacin andvitamin B12 deficiency, which was thought to have been the result of
medullary hyperconsumption during the subclinical phase of the disease(Caprelli et al., 1994). Sheffy (1964) illustrated the importance of folacin in the immune response of folacin-deficient puppies inoculatedwith distemper and hepatitis antigens. Half the dogs were also given
27.5 μg of folacin per kilogram of body weight. Depleted dogs had delayed antibody production responses against both distemper and infectious hepatitis antigens. Antibodies were detected in depleted dogs supplemented with folacin 8 days after challenge with antigen, whereas
depleted dogs without folacin did not show antibodies until 17 days.

FISH
Signs of folacin deficiency in trout and salmon include anorexia, reduced growth, poor feed conversion, and macrocytic, normochromic,
megaloblastic anemia characterized by pale gills and poikilocytosis (increased numbers of hemocytoblasts) (Cowey and Woodward, 1993;
NRC, 1993). Anorexia, poor growth, and dark skin coloration werenoted in Japanese eels fed a folacin-deficient diet for 10 weeks (Arai et503

C H A P T E R T W E LV E

al., 1972). Folacin-deficient yellowtail fingerlings also showed congestion in fins and bronchial mantle, dark skin coloration, and anemia(Hosokawa, 1989). Folacin deficiency signs in channel catfish included
reduced growth, anemia, and increased sensitivity to bacterial infection(Duncan and Lovell, 1991). Anemia in folate-deficient channel catfish
was characterized by pale livers, spleens, gills, and kidneys and by poikilocytosis (Duncan et al., 1993).
An anemic disease in catfish known as “no blood disease” or “white
lip disease” was reported to result in catfish fed sulfonamide-degrading
Pseudomonas bacteria (Plumb et al., 1991). The researchers were able
to isolate two strains of folacin-degrading enzymes from feed, environment, and catfish tissue samples. Catfish fed sulfonamide showed higher
mortality, lower weight gain, lower thrombocyte counts, higher hemocytoblast and neutrophil counts, and lower liver folacin values compared with controls (no folacin), indicating that significant intestinalbacterial synthesis of folacin occurs in channel catfish (Duncan et al.,1993).

FOXES AND MINK
Both adult and growing foxes receiving a folacin-deficient diet exhibit anorexia; body weight loss; and a decrease in hemoglobin, erythrocyte, and leukocyte concentrations (NRC, 1982a; Pölönen et al.,1997). Folacin deficiency in mink results in anorexia, growth depression, diarrhea, and ulcerative hemorrhagic gastritis (NRC, 1982a; Siddons, 1978).

LABORATORY ANIMALS
Rodent species have generally similar folacin-deficiency signs, including reduced growth, anemia, leukopenia, evidence of reduced protein synthesis, reduced folacin tissue levels, and impaired antibody response (Siddons, 1978; NRC, 1995; Achón et al., 1999). An increase inurinary excretion of FIGLU occurs with folacin deficiency in the rat andmouse. In rats, severe folacin deficiency caused secondary hepaticcholine deficiency (Kim et al., 1994). Folacin deficiency has been shownto have a teratogenic effect on the rat, with congenital abnormalities inoffspring, including hydrocephalus. Hamsters resemble guinea pigs
rather than rats in their ability to develop folacin deficiency without theuse of intestinal antiseptic or folacin antagonist (Cohen et al., 1971). For
the rat, folacin biosynthesis by intestinal bacteria may meet much of therequirement for the vitamin (Rong et al., 1991).
Deficiency of folacin in the pregnant mouse has an adverse effect on504

FOLACIN

reproduction and lactation. Shaw et al. (1973) observed decreased
growth, especially of brain and liver, in young mice deprived of folate
both prenatally and postnatally. Cerecedo and Mirone (1947) reported
that folacin increased survival to weaning of suckling mice from 34%(basal diet) to 69%.

NONHUMAN PRIMATES
After 3 to 9 months, monkeys on a low-folacin diet lost weight, became inactive, and between 3 and 18 months, developed a megaloblastic macrocytic anemia (NRC, 1978; Siddons, 1978; Thenen et al., 1991).
In rhesus monkeys a variety of clinical and hematological signs arise asa result of the deficiency, including weight loss, anorexia, listlessness,
mucoid to bloody diarrhea, gingivitis and oral ulceration, increased susceptibility to infection (especially dysentery), macrocytic anemia,
leukopenia, and increased excretion of FIGLU. Leukopenia is the mostcharacteristic hematological sign, and in some monkeys, it may prove fatal without the development of anemia.

HumansFor humans, folacin deficiency is probably the most common vitamin deficiency in the world. Infants, adolescents, elderly persons, andpregnant women seem particularly vulnerable. Studies involving theWorld Health Organization in various countries suggest that up to onethird of all pregnant women in the world have folacin deficiency (Herbert, 1981). In a clinic in New York City, 16% of pregnant women were
deficient, with erythrocyte folacin below 150 ng/ml. A further 14% had
erythrocyte folacin concentrations suggestive of a deficiency. In a study
in Paris, 18% of immigrant pregnant women were deficient in folacin(Hercberg et al., 1987). Black South African women had a high incidence of megaloblastic anemia of pregnancy, and 40% of apparently
otherwise healthy pregnant women had morphological and biochemical
evidence of folate deficiency (Colman, 1982).
Adolescent girls have a greater nutritional requirement for folacin in
relation to body size than do adult women (Heald, 1975). Additional folacin demand of pregnancy and poor dietary habits may compromise
their growth potential and increase risk in pregnancy. Folacin deficiency
in the older population is due in part to an impairment of dietary folate
utilization (Baker et al., 1978). Elderly subjects (older than 60 years)
were unable to utilize polyglutamate forms of folacin effectively compared to younger subjects, indicating a lack of conjugase enzymes. In
Italy (Maiani et al., 1992), 15% of free living elderly were deficient in
505

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folacin, while in Holland (Lowik et al., 1992), 28% of elderly women
living in a nursing home were deficient in the vitamin. In Spain 80% of
free living elderly subjects were deficient in folacin on the basis of low
serum and red blood cell folacin values (Ortega et al., 1993).
Infants have a high requirement for folacin, with preterm infants reported to have significantly lower erythrocyte folacin than those born atterm (Ek, 1980). These data would suggest a heightened maternofetaltransfer of folacin during the last few weeks of pregnancy. One study
evaluated the folacin status of 31- to 210-day-old Brazilian infants from
low-income families (Trugo et al., 1991). Some of the babies were exclusively breast-fed, and some were partially weaned. Low serum folacin
was found in 33% of exclusively breast-fed infants and in 27% of partially weaned infants.
Typical reaction to folacin deficiency in the human is megaloblastic
red cell maturation in the bone marrow with resulting macrocytic anemia (Fig. 12.7), which is accompanied by leukopenia. The macrocytic
anemia that occurs resembles pernicious anemia without the nervous
system involvement. Glossitis, gastrointestinal lesions, diarrhea, and intestinal malabsorption may accompany macrocytic anemia. Likewise,
clinical findings of folacin deficiency include pallor, weakness, forgetfulness, sleeplessness, and bouts of euphoria (Herbert, 1962). The most
rapidly proliferating tissues of the body, such as the bone marrow forblood cell production, have the greatest requirement for DNA synthesis
and thus are principally affected when severe deficiency occurs.
The sequence of signs in the development of human folacin deficiency was reported by Herbert (1967), who examined biochemical andhematological changes in himself as he consumed a low-folacin diet overtime (Table 12.3). After only 3 weeks of dietary folacin deprivation, hisserum folacin dropped from 7 to less than 3 ng/ml. However, low erythrocyte folacin did not appear until 4 months after initiation of folacin
deprivation. At about 4.5 months, his bone marrow became megaloblastic and anemia developed.
Apart from features due to megaloblastic anemia or to abnormalities of the epithelial cell surfaces, sterility and gastrointestinal abnormalities are established effects of folacin deficiency (Hoffbrand, 1978).
Sterility in both men and women and, in a small percentage of cases,
widespread reversible melanin pigmentation of the skin, mainly affecting the skin creases and nail-beds, have been reported (Fleming andDawson, 1972). Folacin deficiency may be related to abnormalities ofpregnancy, including postpartum hemorrhage, antepartum hemorrhage,
congenital malformation, and prematurity. Clinical signs related to the506

FOLACIN

A

B

Fig. 12.7 Pernicious anemia in a human, characterized by (A) marked hyperplasia in bone marrow compared to (B) normal. Deficiency of either folacin or
vitamin B12 results in ineffective erythropoiesis. (Courtesy of R.R. Streiff, Veterans Administration, University of Florida.)

gastrointestinal tract include sore tongue, angular cheilosis, loss of appetite, and diarrhea (Rose, 1971). The tongue may appear red and shiny
or smooth, pale, and atrophic. Some researchers have shown structuraland functional jejunal changes in nutritional folacin deficiency that canbe reversed by supplemental folacin (Hoffbrand, 1978). There is a complicated relationship between tropical sprue and folacin deficiency. Nutritional folacin deficiency may predispose to the disease, and that deficiency, due to reduced diet and to malabsorption, may aggravate thesmall intestinal lesion.
Numerous studies have shown that supplemental folacin (0.4 to 4507

C H A P T E R T W E LV E
■ Table 12.3

Progressive Folacin Deficiency Sequence of Events in a 35Year-Old Human MaleTime (weeks)Biochemical and Hematological Events

3
7
13
17
19–20

Low serum folacin (< 3 ng/ml)
Hypersegmentation of neutrophils (leukocytes)
High urine formiminoglutamic acid (FIGLU)
Low RBC folacin (< 20 ng/ml)
Megaloblastic anemia

Source: Modified from Herbert (1967).

mg) can prevent serious birth defects, known as neural tube defects, indeveloping fetuses (Rush, 1992; Bower, 1995; Scott et al., 1995). In the
United States and around the world, neural tube defects—anencephaly,
spina bifida, and encephalocoele—contribute significantly to infantmortality and morbidity. According to the Centers for Disease Control
report (1992), 4,000 infants with neural tube defects are born annuallyin the United States; worldwide, nearly 400,000 are born with these conditions (Rush, 1992). Neural defects occur when the neural tube, which
will develop into the spinal cord, fails to close 18 to 26 days after actual
conception. If the neural tube defect occurs toward the top of the tube,
the child is born with anencephaly (without a brain) and will die soonafter birth. If the defect occurs further down the tube, the child will beborn with spina bifida (an open spine) and will be crippled for life
(Bower, 1995).
The exact mechanism by which folacin prevents these birth defects
is not known. There is substantial geographical and ethnic variation in
the prevalence of neural tube defects; the etiology is unknown but is believed to be multifactorial, involving both genetic and environmental determinants. Evidence indicates that vitamin B12 is also an independentrisk factor, which makes it very probable that, either directly or indirectly, the primary cause of neural tube defects is a defective enzyme.
This enzyme is most probably in the affected fetus as well as in themother, namely methionine synthase, since this is the only point in metabolism at which both folacin and vitamin B12 interact (Scott et al.,
1995; see also Chapter 13, Deficiency, Humans).
Data have accumulated indicating that low folacin status may promote cancer. Low folacin levels are associated with cytogenetic abnormalities in vivo and in vitro. Epidemiological studies examined the association between folacin and cancer of the cervix, colorectum, lung,esophagus, and brain and suggested that low folacin status may play an
508

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important role early in the neoplastic process (Glynn and Albanes,1994). Butterworth (1993) suggested that a significant reduction in earlycervical cancer could be achieved in up to two-thirds of high-risk populations through improved nutritional intake of folacin before exposure
to an oncogenic strain of papilloma virus.
In a prospective analysis of over 25,000 men and women, Giovannucci et al. (1993) found an inverse association between dietary folacin
and the incidence of adenomatous polyps in the distal colon and rectum.
They also found an inverse association with another dietary source of
methyl groups (methionine), as well as a direct association with moderate alcohol consumption, which is known to adversely affect folacin metabolism. This suggests that diminished folacin status and excessive alcohol intake, which may decrease S-adenosyl-methionine levels, may
induce hypomethylation of DNA, thereby promoting colorectal cancer.
Studies suggest that folacin deficiency may be an important causeof a moderate hyperhomocysteinemia, which is an independent risk factor for cardiovascular disease (Boushey et al., 1995; Selhub et al.,1996). All but a few of over 75 studies, including a total of more than
15,000 investigated patients and controls, support this issue(Brattström, 1996). Basal hyperhomocysteinemia unmasked by a methionine load test is a marker for increased cardiovascular risk (Ueland
et al., 1992).
Some studies suggest that folacin deficiency may be responsible forup to 40% of the heart attacks and strokes suffered by American men
every year (Raeburn, 1995). Studies have consistently found that men
with elevated homocysteine levels in the blood are three times more
likely to have a heart attack over the next 5 years than men with normal
homocysteine levels. Elevated homocysteine levels can be caused by genetic defects in homocysteine metabolism, or by lack of vitamin B6 or folacin, and vitamin B12. Supplementation with folacin has been shown tomarkedly reduce elevated plasma homocysteine concentrations and reduce normal homocysteine concentrations (Brattström, 1996). Folacindoses of less than 1 mg/day may be effective. Supplementation with a
combination of folacin and vitamin B12 will secure the full homocysteine-lowering effect and prevent occurrence of a vitamin B12 deficiency
during the course of therapy.
Folacin deficiency is often associated with chronic alcoholism. Between 40 and 87% of alcoholics admitted to municipal hospitals in the
United States have low serum folacin, and between 40 and 61% havemegaloblastic anemia (Halsted and Tamura, 1979). Folacin deficiency
appears to be a result of a complex interaction between nutritional dep509

C H A P T E R T W E LV E

rivation and chronic alcohol ingestion. Alcohol specifically inhibits
hematopoiesis, and Halsted et al. (1973) reported that the jejunal uptakeof folacin was significantly greater in subjects who remained sober.
Hidiroglou et al. (1994) suggested that alcohol exerts its effect through(1) inhibition of B12-dependent methyl transfer from methyltetrahydrofolate to homocysteine; (2) diversion of formylated tetrahydrofolates toward serine synthesis; and (3) interaction of acetaldehyde with tetrahydrofolates, which thereby interferes with folacin coenzyme metabolism.
Both poor folacin status and moderate to excessive alcohol consumption have been associated with increased risk of colorectal cancer,
although the mechanisms through which these effects occur have notbeen established. Reports suggest that diminished folacin status and excessive alcohol intake—which may decrease S-adenosylmethionine levels, thereby depleting methionine—may induce hypomethylation ofDNA, thus promoting colorectal cancer (Giovannucci et al., 1993).
Folacin has been shown to have a role in neuropsychiatry. In many
countries there is a high incidence of folacin deficiency in psychiatric andpsychogeriatric patients, especially inpatients (Bottiglieri et al., 1995).
The deficiency can be associated with any diagnostic category but ismore common in patients with depression and dementia. There is someevidence that even a borderline deficiency can be harmful to the mental
state (Godfrey et al., 1990). There is growing evidence that whether the
deficiency is primary or secondary to the psychiatric disorder, folacin replacement may improve mental function when added to standard psychotropic medication.

Assessment of Status
Assessment of nutritional status of folacin can involve dietary evaluation, clinical signs, response to supplementation, and laboratory
analysis. Regarding dietary history, humans or animals (particularly
poultry) that have not received green leafy plant sources or organ meats
would suggest reduced folacin intake. Because liver contains a high percentage of stored folacin, concentration in this organ would serve as afolacin status indicator. On low-folacin diets, liver concentrations are
depleted in a few months. Clinical signs of folacin deficiency are extremely variable and are less precise than laboratory analysis to confirm
deficiency. Using a protocol of folacin depletion-repletion of rats, followed by assessment of growth and liver, serum, and erythrocyte folacin
concentrations, has been successful in evaluating bioavailability of food
folacin sources (Clifford et al., 1990, 1991).
In humans, a positive diagnosis of folacin deficiency is usually made510

FOLACIN

by the finding of subnormal serum and erythrocyte levels. Cutoff valuesof less than 0.3 ng/ml for serum folacin and less than 140 ng/ml for erythrocyte folacin are the basis for estimation of the prevalence of low values of the vitamin (Senti and Pilch, 1985). Patients with vitamin B12 deficiency exhibit all the clinical and hematological features and many ofthe biochemical features of folacin deficiency. Thus, erythrocyte folacin
assay is better used in conjunction with the values for serum vitamin B12.
Measurement of tissue content of several forms of folacin and urinary excretion of FIGLU, a histidine catabolite that cannot be normallymetabolized in folacin deficiency, is utilized to assess folacin nutrition. It
has been shown that the amount of FIGLU excreted in urine roughly
parallels the erythrocyte folate and hepatic folacin levels in both anemic
and nonanemic folacin-deficient patients and thus, like the erythrocyte
folate and hepatic folacin levels, appears to be a satisfactory index of tissue folacin stores (Herbert, 1967). A 24-hour urinary excretion level after a single oral dose of isotopically labeled folacin is a functional indicator of folacin status (Gregory et al., 1998).
Deficiencies of iron and vitamin C can be related to folacin status,
but it has not yet been proven that folacin requirement is increased in either iron deficiency anemia or scurvy. The frequent occurrence of combined iron and folacin deficiencies has led to the suggestion that iron deficiency may be responsible for the development of secondary folacin
deficiency. Iron deficiency has been reported to influence folacin metabolism in pregnant women. The stress of lactation in rats superimposedon iron deficiency was found to alter milk folacin concentration, resulting in folacin depletion in rat pups (Kochanowski et al., 1983). Bothiron and folacin are required for normal hematopoiesis, with anemia resulting from lack of either nutrient. Iron deficiency may mask thechanges in the developing erythroblasts but does not affect the white cellabnormalities. Anemia in humans is often assumed to be the result ofiron deficiency; however, Bailey et al. (1980) reported that folacin deficiency was much more prevalent than iron deficiency in a lower-income
pregnant Florida population. Whether the association of scurvy andmegaloblastic anemia is due to increased folacin requirements secondary
to lack of vitamin C or is merely the result of a double nutritional deficiency is unresolved. However, it is interesting to speculate that manycases of vitamin C deficiency could likewise be folacin deficiencies, sinceboth vitamins are rich in green plants and citrus fruits and lacking inmost other food sources.
The measurement of serum homocysteine was shown to be a good
indicator of folacin and vitamn B12 nutritional status (Krumdieck, 1990;
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C H A P T E R T W E LV E

Jacob, 1998). The folate-vitamin B12-requiring remethylation of homocysteine to methionine normally converts approximately 50% of available homocysteine back to methionine.
In most human patients with folacin deficiency, a combination offactors leads to negative folacin balance. Poor diet is usually the major
cause, since few diseases cause malabsorption or poor folacin utilization
to become so severe that good intake of the vitamin cannot overcome
losses. It is likely that severe folacin deficiency may occur in tropicalsprue and congenital specific malabsorption of folacin, despite a normal
dietary folacin content (Hoffbrand, 1978).

SUPPLEMENTATION
Folacin needs for livestock are often met by good practical diets,and for most species, substantial quantities of folacin are provided
through microbial synthesis. Nevertheless, field observations have been
made on folacin-insufficient diets. Green forage is an excellent source of
folacin. Supplementation of folacin would be most needed when animals
are in confinement without access to green grazing or preserved green
forages. The successful treatment of field cases of folacin deficiency with
supplemental folacin has demonstrated that commercial feeds do not always supply adequate quantities of the vitamin to poultry (Pesti et al.,1991).
Of farm livestock, poultry would most likely need supplemental folacin and then only under certain conditions. Newly weaned pigs may
also need additional folacin in their diets for optimum growth and feed
efficiency. The supplementation of gestating swine diets with folacin to
maximize sow productivity is indicated by research (Matte et al., 1984b;
Tremblay et al., 1989; Lindemann, 1993). There was consistent increase
in total and live pigs born when reproducing females received supplemental folacin during gestation. The increase has occurred at dietary
supplementation levels from 0.2 mg/kg (Easter et al., 1983) to 15 mg/kg(Matte et al., 1990) as well as with folacin injections throughout the first
12 weeks of gestation (Matte et al., 1984b, 1992). Matte et al. (1992)
observed that dietary supplementation of 15 mg/kg was necessary to
avoid the drop in concentrations of serum folates of sows in gestationand seemed to influence some traits of the reproductive performance. It
seems apparent and logical that supplementation must occur in earlygestation. The response appears to be greater in conditions of increased
ovulation (e.g., sows versus gilts, flushed versus nonflushed), which sug-

512

FOLACIN

gests that there may be breed differences in response to folacin supplementation (Lindemann, 1988).
Only young ruminants that do not have a fully developed rumen
would be expected to require a dietary source. Fluctuations in the concentration of serum folacin observed during the first months of life ofyoung ruminants may be an indication that synthesis of folacin by ruminal microflora is not sufficient to meet requirements during weaning(Girard et al., 1989a). The age effect on development of the rumen wasobserved, in which parenterally administered folacin markedly increased
serum folacin in 2-week-old heifers, while in 4-month-old heifers, the increase was less marked. For adult dairy cows, Girard et al. (1989b) reported that serum folacin can be increased by intramuscular injection of
folacin, but in cows during early lactation, this does not markedly affectserum and milk folacin. Girard et al. (1995) injected folacin (160 mg)
weekly into dairy cows from mating to 6 weeks postparturition; the supplementation achieved maximal production of milk and protein in multiparous cows.
Folacin may be of little benefit when poultry and swine receive only
low levels of sulfa drugs and consume grains relatively free of toxin-producing molds. However, since a large percentage of the U.S. corn crop
has some mold contamination, folacin supplementation should have apositive effect in many commercial poultry and hog operations as wellas in other livestock enterprises (Purser, 1981). This would likely be aneven more important consideration in developing tropical countries,
where conditions favoring mold growth are optimized. Individual responses to folacin supplementation to counteract mold effect will obviously vary with the class of livestock being fed, species of mold present,
and the levels of toxin encountered (Bhavanishankar et al., 1986).
Purser (1981) fed supplemental folacin to pigs weaned at 3, 4, or 5
weeks of age. At the end of the 4-week feeding period, the younger pigs
showed the greatest response to diets providing the supplemental vitamin.
Gadient (1986) found folacin to be very sensitive to heat and light,
slightly sensitive to moisture, and insensitive to oxygen. Frye (1978)
found that folacin can be lost during storage of premixes, particularly atelevated temperatures. After 3 months of room-temperature storage,
43% of the original folacin activity was lost. Verbeeck (1975) found folacin to be stable in premixes without minerals, but there may be as
much as 50% loss in a premix with minerals kept at room temperaturefor 3 months. Adams (1982) reported only 38% retention of folacin ac-

513

C H A P T E R T W E LV E

tivity in a premix without minerals after 3 weeks at 45oC. However, hereported 57% retention of activity after 3 months at room temperature.
One suggestion would be to almost double the amount of folacin in a
premix at the time of manufacture to ensure that poultry or swine receive the desired amount from that premix, since a 3- to 4-month period
is not an unreasonable amount of time from premix manufacture to customer purchase to (ultimately) diet mixing and feeding.
Slinger et al. (1979) reported processing and storage losses of folacin
in fish feeds of 5 to 10% for steam-pelleted crumbles and 3 to 7% for
extruded crumbles, depending on dietary level. Scott (1966) indicatedthat an adjustment of 10 to 20% in the folacin level in poultry feed may
be necessary because of pelleting losses. Coelho (1994) reported a 21%
loss of folacin in feeds as a result of pelleting and 2 weeks of storage.
It is important that women begin supplementation of folacin before
they plan to become pregnant, or at least to consume adequate folacin
if there is a chance of pregnancy occurring. They should not wait until
pregnancy is confirmed, because supplementation at that time is too late
to provide protection from birth defects. Neural tube defects develop
very early in pregnancy (18 to 26 days); therefore supplemental folacinmust be provided in the preconception period before a woman knows
she is pregnant.
Humans obtain folacin primarily from fruits and vegetables, and an
average western diet supplies 0.15 to 0.2 mg/day (Metz, 1995). Womenof childbearing age would need to increase their folacin intake threefold
to reach the level of 0.4 mg/day, the amount of folacin known to be effective against neural tube defects and which does not mask the diagnosis of pernicious anemia and/or vitamin B12 deficiency.
Supplementation with large doses of folacin will cure the macrocytic
anemia in human patients with the vitamin B12 deficiency, pernicious
anemia. However, folacin will not prevent the often irreversible neurological lesions of vitamin B12 deficiency or pernicious anemia. Therefore,
large quantities (more than 1 mg/day) of supplemental folacin should
not be taken indiscriminately as they may obscure a diagnosis of vitamin
B12 deficiency. The U.S. Public Health Service cautioned against recommending daily intakes above 1 mg/day, except under physician supervision (Anonymous, 1996).
To reduce the risk of neural tube defects, the U.S. Food and Drug
Administration (FDA) has required (since January 1, 1998) that folacin
be added to most enriched flour, breads, corn meals, rice, noodles, macaroni, and other grain products (Anonymous, 1996). These foods were514

FOLACIN

chosen for folacin fortification because they are staple products for mostof the U.S. population, and because they have a long history of being
successful vehicles for improving nutrition by reducing the risk of classic nutrient deficiency diseases. The FDA is requiring that the abovenoted food products be fortified with folacin at a level of 140 mg/100 g.
This supplementation level will likely provide at least 0.4 mg folacin
daily. The purpose of mandatory folacin supplementation in the UnitedStates is to reduce neural tube defects; however, benefits will surely bederived in providing protection against megaloblastic anemia, cardiovascular disease, and various forms of cancer (e.g., colorectal), and inpreventing other conditions that result from folacin deficiency. As an example, assuming that correction of the homocysteinemia diminishes therisk of vascular disease and given the apparent high prevalence of thetrait, a vast opportunity for prevention of coronary disease by giving
these patients supplemental folacin may exist. Likewise, folacin deficiency enhances tumor development in rats, which is preventable with
folacin supplementation (Henning et al., 1997).
Crystalline folacin, produced by chemical synthesis, is available forfeeds, foods, and pharmaceuticals. Although folacin is only sparingly
soluble in water, the sodium salt is quite soluble and is used in injections
as well as feed supplements (McGinnis, 1986; Tremblay et al., 1986).
For humans, folacin supplements (monoglutamate form) ranging from100 to 1,000 μg/day have been recommended by different investigators.
Oral supplementation appears to be desirable to maintain maternal
stores and to keep pace with the increased folacin turnover that is seen
in rapidly growing tissue. Synthetic folacin supplements are convenient
to take and highly available; natural forms are approximately 50% lessavailable than synthetic forms.

TOXICITY
Folacin generally has been regarded as a nontoxic vitamin (NRC,1987). Acute intravenous toxicity is very low, with the LD50 as follows(in milligrams per kilogram of body weight): mice, 600; rats, 500; rabbits, 410; and guinea pigs, 120. In rats, most of the deaths occurred
within 30 minutes of injection (Anonymous, 1961). Rabbits given 50
mg/kg/day intraperitoneally for 10 weeks were possibly retarded in
growth and did not differ in blood picture, number of deaths, or generalappearance, but did show signs of renal injury at autopsy. Folacin has a
low acute and chronic toxicity for humans. In adults, no adverse effects

515

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were noted after 400 mg/day for 5 months or after 10 mg/day for 5 years(Brody, 1991). The danger of excess folacin is indirect in that the folacin
supplements can obscure the diagnosis of vitamin B12 deficiency; they
can prevent anemia while permitting neurological damage.

■ REFERENCES
Abad, A.R., and Gregory, J.F. (1987). J. Nutr. 117, 866.
Achón, M., Reyes, L., Alonso-Aperte, E., Ubeda, N., and Varela-Moreiras, G.
(1999). J. Nutr. 129, 1204.
Adams, C.R. (1982). In Vitamins—The Life Essentials. Nutrition Institute, National Feed Ingredients Association, NI-82, 1, Des Moines, Iowa.
Anonymous (1961). Vitamin Manual, p. 53. Upjohn, Kalamazoo, Michigan.
Anonymous (1996). Nutr. Rev. 54(3), 94.
Arai, S., Nose, T., and Hashimoto, Y. (1972). Bull Freshwater Res. Lab. Tokyo22, 69.
Bailey, L.B. (1995). In Folate in Health and Disease (L.B. Bailey, ed.), p. 123.
Marcel Dekker, New York.
Bailey, L.B., Manan, C.S., and Dimpeno, D. (1980). Am. J. Clin. Nutr. 33, 1997.
Baker, H., Jaslow, S.P., and Frank, O. (1978). J. Am. Geriatr. Soc. 26, 218.
Baker, S.J., and DeMaeyer, E.M. (1979). Am. J. Clin. Nutr. 32, 368.
Balaghi, M., and Wagner, C. (1992). J. Nutr. 122, 1391.
Balaghi, M., Horne, D.W., Woodward, S.C., and Wagner, C. (1993). Am. J. Clin.
Nutr. 58, 198.
Baugh, C.M., and Krumdieck, C.L. (1971). Ann. N. Y. Acad. Sci. 186, 7.
Bechtel, H.E. (1964). Feedstuffs 36(45), 18.
Bhandari, S.D., and Gregory, J.F. (1992). J. Nutr. 122, 1847.
Bhavanishanker, T.N., Shantha, T., and Ramesh, H.P. (1986). Nutr. Rep. Int. 33,
603.
Blakley, R.L., and Benkovic, S.J. (1984). Folates and Pterins. Wiley, New York.
Bottiglieri, T., Crellin, R.F., and Reynolds, E.H. (1995). In Folate in Health andDisease (L.B. Bailey, ed.), p. 435. Marcel Dekker, New York.
Boushey, C.J., Beresford, S.A.A., Omenn, G.S., and Motulsky, A.G. (1995).
JAMA 274, 1049.
Bower, C. (1995). Nutr. Rev. 53, S33.
Brattström, L. (1996). J. Nutr. 126, 1276S.
Brody, T. (1991) In Handbook of Vitamins (L.J. Machlin, ed.), 2nd Ed., p. 453.
Marcel Dekker, New York.
Brown, G.M. (1962). J. Biol. Chem. 237, 536.
Butterworth, C.E. (1993). Pennington Cent. Nutr. Ser. 3, 196.
Butterworth, C.E. Jr., Hatch, K.D., Macaluso, M., Cole, P., Sauberlich, H.E.,
Soong, S.J., Borst, M., and Baker, V.V. (1992). JAMA 267, 528.
Caprelli, J.L., Bohlay, P., and Barre, D. (1994). Pratique-Medicale-et-Chirurgicale-de-l’Animal-de-Compagnie 29, 395.
Carl, G.F., and Smith, M.L. (1995). J. Nutr. 125, 1245.
Carl, G.F., Hudson, F.Z., and McGuire, B.S. (1997). J. Nutr. 127, 2231.
516

FOLACIN

Carroll, F.D., Gross, H., and Howell, C.E. (1949). J. Anim. Sci. 8, 290.
Cartwright, G.E., and Wintrobe, M.M. (1949). Proc. Soc. Exp. Biol. Med. 71,
54.
Cartwright, G.E., Tatting, B., Kurth, D., and Wintrobe, M.M. (1952). Blood 7,992.
Carvalho da Silva, A., deAngelis, R.C., Pontes, M.A., and Mansurguerios, M.F.
(1955). J. Nutr. 56, 199.
Centers for Disease Control (1992). Morb. Mortal. Wkly. Rep. 41, RR-14 (September 11, 1992).
Cerecedo, L.R., and Mirone, L. (1947). Arch. Biochem. 12, 154.
Chandler, C.J., Wang, T.T.Y., and Halsted, C.H. (1986). J. Biol. Chem. 261,
928.
Clifford, A.J., Jones, A.D., and Bills, N.D. (1990). J. Nutr. 120, 1640.
Clifford, A.J., Heid, M.K., Peerson, J.M., and Bills, N.D. (1991). J. Nutr. 121,
445.
Coelho, M.B. (1994). In BASF Technical Symposium, p. 60. Clarion, Arkansas.
Cohen, N.L., Reyes, P.S., and Briggs, G.M. (1971). Lab. Anim. Sci. 21, 350.
Colman, N. (1982). Nutr. Rev. 49, 225.
Cowey, C.B., and Woodward, B. (1993). J. Nutr. 123, 1594.
Cravo, M.L., Mason, J.B., and Dayal, Y. (1992). Cancer Res. 52, 5002.
Creek, R.D., and Vasaitis, V. (1963). Poult. Sci. 42, 1136.
Cruces-Blanco, C., Segura-Carretero, A., Fernandez-Guitierrez, A., and RomanCeba, M. (1994). Anal. Lett. 27, 1339.
Cunha, T.J., Colby, R.W., Bustad, L.K., and Bone, J.F. (1948). J. Nutr. 36, 215.
Davenport, D.J., Ching, R.J.W., Hunt, J.H., Bruyette, D.S., and Gross, K.L.
(1994). J. Nutr. 124, 2559S.
Dhur, A., Galan, P., and Hereberg, S. (1991). Prog. Food Nutr. Sci. 15, 43.
Draper, H.H., and Johnson, B.C. (1952). J. Nutr. 46, 123.
Dumoulin, P.G., Girard, C.L., Matte, J.J., and St-Laurent, G.J. (1991). J. Anim.
Sci. 69, 1657.
Duncan, P.L., and Lovell, R.T. (1991). Twenty-second Ann. Confer. World
Aquacul. Soc. San Juan, Puerto Rico, June 16–20, 1991.
Duncan, P.L., Lovell, R.T., Butterworth, C.E., Jr., Freeberg, L.E., and Tamura, T.
(1993). J. Nutr. 123, 1888.
Easter, R.A., Anderson, P.A., Michel, E.J., and Corley, J.R. (1983). Nutr. Rep.
Int. 28, 945.
Ek, J. (1980). J. Pediatr. 97, 288.
Fleming, A.F., and Dawson, I. (1972). Br. Med. J. 4, 236.
Foo, S.K., and Shane, B. (1982). J. Biol. Chem. 257, 13587.
Frye, T.M. (1978). Proc. Roche Vitam. Nutr. Update Meet. Arkansas Nutr.
Conf. p. 54.
Gadient, M. (1986). Proc. Nutr. Conf. Feed Manuf., College Park, Maryland, p.
73.
Giovannucci, E., Stampfer, M.J., and Colditz, G.A. (1993). J. Natl. Cancer Inst.
85, 875.
Girard, C.L., and Matte, J.J. (1998). J. Dairy Sci. 81, 1412.
Girard, C.L., Matte, J.J., and Roy, G.L. (1989a). Br. J. Nutr. 61, 595.
Girard, C.L., Matte, J.J., and Tremblay, G.F. (1989b). J. Dairy Sci. 72, 3240.
517

C H A P T E R T W E LV E

Girard, C.L., Matte, J.J., and Lévesque, J. (1992). J. Anim. Sci. 70, 2847.
Girard, C.L., Chiquett, J., and Matte, J.J. (1994). J. Anim. Sci. 72, 1023.
Girard, C.L., Matte, J.J., and Tremblay, G.F. (1995). J. Dairy Sci. 78, 404.
Giugliani, E.R.J., Jorge, S.M., and Goncalves, A.L. (1985). Am. J. Clin. Nutr.
41, 330.
Glynn, S.A., and Albanes, D. (1994). Nutr. Cancer, 22, 101.
Godfrey, P.S.A., Toone, B.K., and Carney, M.W.P. (1990). Lancet 336, 392.
Green, R., and Jacobsen, D.W. (1995). In Folate in Health and Disease (L.B. Bailey, ed.), p. 75. Marcel Dekker, New York.
Gregory, J.F. (1989). Adv. Food Nutr. Res. 33, 1.
Gregory, J.F., Sartain, D.B., and Day, B.P.F. (1984). J. Nutr. 114, 341.
Gregory, J.F., Bhandari, S.D., Bailey, L.B., Toth, J.P., Baumgartner, T.G., andCerda, J.J. (1991). Am. J. Clin. Nutr. 53, 736.
Gregory, J.F., Williamson, J., Bailey, L.B., and Toth, J.P. (1998). J. Nutr. 128,
1907.
Halsted, C.H., and Tamura, R. (1979). Problems in Liver Disease (C.S. Davidson, ed), p. 91. Stratton, New York.
Halsted, C.H., Robles, E.A., and Mezey, E. (1973). Gasteroenterology 64, 526.
Harper, A.F., Lindemann, M.D., Chiba, L.I., Combs, G.E., Handlin, D.L., Kornegay, E.T., and Southern, L.L. (1994). J. Anim. Sci. 72, 2338.
Heald, F.P. (1975). Med. Clin. North Am. 59, 1329.
Heimburger, D.C., Alexander, C.B., Birch, R., Butterworth, C.E. Jr., Bailey,
W.C., and Krumdieck, C.L. (1988). JAMA 259, 1525.
Henning, S.M., Swendseid, M.E., and Coulson, W.F. (1997). J. Nutr. 127, 30.
Herbert, V. (1962). Trans. Assoc. Am. Physicians 75, 307.
Herbert, V. (1967). Am J. Clin. Nutr. 20, 562.
Herbert, V. (1981). Proc. Fla. Symp. Micronutr. Hum. Nutr. Univ. Fla.,
Gainesville, p. 121.
Herbert, V., and Zalusky, R. (1962). J. Clin. Invest. 41, 1263.
Hercberg, S., Bichon, L., Galan, P., Christides, J.P., Carroget, C., and Potier deCourcey, G. (1987). Nutr. Rep. Int. 35, 915.
Hidiroglou, N., Camilo, M.E., Beckenhauer, H.C., Tuma, D.L., Barak, A.J.,
Nixon, P.F., and Selhub, J. (1994). Biochem. Pharm. 47, 1561.
Hoffbrand, A.V. (1978). Handbook Series in Nutrition and Food, Section E: Nutritional Disorders (M. Rechcigl, ed.), Vol. 2, p. 55. CRC Press, Boca Raton, Florida.
Hosokawa, H. (1989). The vitamin requirements of fingerling yellowtail, Seriola quinqueradiata, Ph.D. dissertation. Kochi University, Japan.
Jacob, R.A. (1998). Nutr. Rev. 56, 212.
Jacob, R.A., Wu, M., Henning, S.M., and Swendseid, M.E. (1994). J. Nutr. 124,
1072.
Johnson, B.C., James, M.F., and Krider, J.L. (1948). J. Anim. Sci. 7, 486.
Kang, S.S., Wong, P.W.K., and Malinow, M.R. (1992). Ann. Rev. Nutr. 12, 279.
Keagy, P.M., and Oace, S.M. (1984). J. Nutr. 114, 1252.
Kim, Y., Miller, J.W., DaCosta, K.A., Nadeau, M., Smith, D., Selhub, J. Zeisel,
S.H., and Mason, J.B. (1994). J. Nutr. 124, 2197.
Kochanowski, B.A., Smith, A.M., Picciano, M.F., and Sherman, A.R. (1983). J.
Nutr. 113, 2471.
518

FOLACIN

Krumdieck, C.L. (1990). In Present Knowledge in Nutrition (M.L. Brown, ed.),6th Ed., p. 179. International Life Sciences Institute/Nutrition Foundation,
Washington, D.C.
Leichter, J., Switzer, V.P., and Landymore, A.F. (1978). Nutr. Rep. Int. 18, 475.
Lévesque, J., Girard, C.L., Matte, J.J., and Brisson, G.J. (1993). Livestock Prod.
Sci. 34, 71.
Lindemann, M.D. (1988). Feedstuffs 60(46), 15.
Lindemann, M.D. (1993). J. Anim. Sci. 71, 239.
Lindemann, M.D., and Kornegay, E.T. (1986). J. Anim. Sci. 63(Suppl. 1),35(Abstr).
Lindemann, M.D., and Kornegay, E.T. (1989). J. Anim. Sci. 67, 459.
Lindemann, M.D., Blodgett, D.J., Kornegay, E.T., and Schurig, G.G. (1993). J.
Anim. Sci. 71, 171.
Loosli, J.K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.), p. 25.
Academic Press, San Diego, California.
Lowik, M.R.H., Berg, H., Schrijver, J., Odink, J., Wedel, M., and Van Houten,
P. (1992). J. Am. Coll. Nutr. 11, 673.
Maiani, G., Polito, A., Ranaldi, L., Azzini, E., Raguzzini, A., Mobarhan, S., andFerro-Luzzi, A. (1992). Age Nutr. 3, 48.
Matoth, Y., Pinkas, A., and Sroka, C. (1965). Am. J. Clin. Nutr. 16, 356.
Matte, J.J., and Girard, C.L. (1999). J. Anim. Sci. 77, 159.
Matte, J.J., Girard, C.L., and Brisson, G.J. (1984a). J. Anim. Sci. 59, 158.
Matte, J.J., Girard, C.L., and Brisson, G.J. (1984b). J. Anim. Sci. 59, 1020.
Matte, J.J., Girard, C.L., and Brisson, G.J. (1990). J. Anim. Sci. 68(Suppl. 1),
370(Abstr.).
Matte, J.J., Girard, C.L., and Brisson, G.J. (1992). Livestock Prod. Sci. 32, 131.
Matte, J.J., Girard, C.L., and Tremblay, G.F. (1993). J. Anim. Sci. 71, 151.
McGinnis, C.H. (1986). Bioavailability of Nutrients in Feed Ingredients, p. 1.
National Feed Ingredient Association (NFIA), Des Moines, Iowa.
McNulty, H., McPartlin, J.M., and Weir, D.G. (1993). J. Nutr. 123, 1089.
McPartlin, J., Scott, J.M., Halligan, A., Darling, M., and Weir, D.G. (1993).
Lancet 341(8838), 148.
Metz, J. (1995). Med. J. Aust. 163, 231.
Muldoon, R.T., Ross, D.M., and McMartin, K.E. (1996). J. Nutr. 126, 242.
Murphy, M., Keating, M., Boyle, P., Weir, D.G., and Scott, J.M. (1976).
Biochem. Biophys. Res. Commun. 71, 1017.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
519

C H A P T E R T W E LV E

(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed. National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
O’Broin, S., and Kelleher, B. (1992). J. Clin. Pathol. 45, 344.
O’Dell, B.L., and Hogan, A.G. (1943). J. Biol. Chem. 149, 323.
Ortega, R.M., Redondo, R., Andres, P., and Equileor, I. (1993). Int. J. Vit. Nutr.
Res. 63, 17.
Pesti, G.M., Rowland, G.N., and Ryu, K.S. (1991). Poult. Sci. 70, 600.
Plumb, J.A., Liu, P.R., and Butterworth, C.E., Jr. (1991). J. Appl. Aquacult. 1,
33.
Pollard, W.O., and Creek, R.D. (1964). Poult. Sci. 43, 1415.
Pölönen, I.J., Vahteristo, L.T., and Tanhuanpää, E.J. (1997). J. Anim. Sci. 75,
1569.
Porras-Tejero, E., and Lluch-Fernandez, M.D. (1993). Anales Espanoles de Pediatria 38, 113.
Purser, K. (1981). Anim. Nutr. Health April, 38.
Raeburn, P. (1995). Vitamin Deficiency Tied to Heart Attacks, San Francisco
Examiner, July 25.
RDA. (1980). Recommended Dietary Allowances, 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
RDA. (1989). Recommended Dietary Allowances, 10th Ed. National Academyof Sciences-National Research Council, Washington, D.C.
Robel, E.J. (1993). Poult. Sci. 72, 546.
Roberts, M.C. (1983). Aust. Vet. J. 60, 106.
Rong, N., Selhub, J., Goldin, B.R., and Rosenberg, I.A. (1991). J. Nutr. 121,
1955.
Rose, J.A. (1971). Lancet 2, 453.
Rothman, D. (1970). Am. J. Obstet. Gynecol. 108, 149.
Rush, D. (1992). Nutr. Rev. 50(1), 25.
Sauberlich, H.E., Kretsch, M.J., Skala, J.H., Johnson, H.L., and Taylor, P.O.
(1987). Am. J. Clin. Nutr. 46, 1016.
Savage, D.G., and Lindenbaum, J. (1995). In Folate in Health and Disease (L.B.
Bailey, ed.), p. 237. Marcel Dekker, New York.
Schweigert, B.S., German, H.L., Pearson, P.B., and Sherwood, R.M. (1948). J.
Nutr. 35, 89.
Scott, J.M., Weir, D.G., and Kirtie, P.N. (1995). In Folate in Health and Disease(L.B. Bailey, ed.), p. 329. Marcel Dekker, New York.
Scott, M.L. (1966). Proc. Cornell Nutr. Conf., p. 35. Ithaca, New York.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Seckington, I.M., Huntsman, R.G.,and Jenkins, G.C. (1967). Vet. Rec. 81, 158.
Selhub, J. (1989). Anal. Biochem. 182, 84.
Selhub, J., Jacques, P.F., Bostom, A.G., D’Agostino, R.B., Wilson, P.W.F., Belanger, J., O’Leary, D.H., Wolf, P.A., Rush, D., Schaefer, E.J., and Rosen520

FOLACIN

berg, I.H. (1996). J. Nutr. 126, 1258S.
Semchuk, G.M., Allen, O.B., and O’Connor, D.L. (1994). J. Nutr. 124, 1118.
Senti, F.R., and Pilch, S.M. (1985). J. Nutr. 115, 1398.
Shaw, W., Schreiber, R.A., and Zemp, J.W. (1973). Nutr. Rep. Int. 8, 219.
Sheffy, B.E. (1964). In Cornell Nutr. Conf., p. 159. Cornell University, Ithaca,
New York.
Shoda, R., Mason, J.B., Selhub, J., and Rosenberg, I.H. (1990). J. Nutr.
Biochem. 1(5), 257.
Siddons, R.C. (1978). Handbook Series in Nutrition and Food: Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 2, p. 123. CRC Press, Boca
Raton, Florida.
Slinger, S.J., Razzaque, A., and Cho, C.Y. (1979). Proc. World Symp. Finfish
Nutr. Fishfeed Technol. 2, 425.
Stancic, B., Piuko, J., Grafenau, P., Sijacic, L., Laurincik, J., Oberfranc, M., andSahinovic, R. (1993). J. Farm Anim. Sci. 26, 13.
Tamura, T., Shane, B., Baer, M.T., King, J.C., Margen, S., and Stokstad, E.L.R.
(1978). Am. J. Clin. Nutr. 31, 1984.
Tani, M., and Iwai, K. (1984). J. Nutr. 114, 778.
Taylor, L.W. (1947). Poult. Sci. 26, 372.
Thenen, S.W., Hwang, S.M., Blocker, D.E., and Meadows, C.A. (1991). Int. J.
Vitam. Nutr. Res. 61, 310.
Tremblay, G.F., Matte, J.J., Lemieux, L., and Brisson, G.J. (1986). J. Anim. Sci.
63, 1173.
Tremblay, G.F., Matte, J.J., Dufour, J.J., and Brisson, G.J. (1989). J. Anim. Sci.
67, 724.
Trugo, N.M.F., Donangelo, C.M., Koury, J.C., Freitas, L.A., and Feldheim, W.
(1991). Ecol. Food Nutr. 25, 333.
Ueland, P.M., Refsum, H., and Brattström, L. (1992). In Cardiovascular Disease, Hemostasis, and Endothelial Function (R.B. Francis Jr., ed.), p. 183.
Marcel Dekker, New York.
Vallet, J.L., Christenson, R.K., and Klemcke, H.G. (1999). J. Anim. Sci. 77,
1236.
Verbeeck, J. (1975). Feedstuffs 47(36), 4, 45.
Wagner, C. (1984). Nutrition Reviews: Present Knowledge in Nutrition (R.E.
Olson, H.P. Broquist, C.O. Chichester, W.J. Darby, A.C. Kolbye, and R.M.
Stalvey, eds.), 5th Ed. Nutrition Foundation, Washington, D.C.
Wagner, C. (1995). In Folate in Health and Disease (L.B. Bailey, ed.), p. 23. Marcel Dekker, New York.
Wagner, C. (1996). J. Nutr. 126, 1228S.
Wagonfeld, J.B., Dudzinsky, D., and Rosenberg, I.H. (1975). Clin. Res. 23,259(Abstr.).
Welch, A.D., Heinle, R.W., Sharpe, G., George, W.L., and Epstein, M. (1947).
Proc. Soc. Exp. Biol. Med. 65, 364.
White, W.E., Yielding, K.L., and Krumdieck, C.L. (1976). Biochem. Biophys.
Acta. 429, 689.
Wiese, A.C., Johnson, B.C., Mitchell, H.H., and Nevens, W.B. (1947). J. Dairy
Sci. 30, 87.
Young, R.J., Norris, L.C., and Heuser, G.F. (1955). J. Nutr. 55, 353.
521

chapter thirteen

V I T A M I N B1 2

INTRODUCTION
Vitamin B12 was the last vitamin to be discovered (1948) and themost potent of the vitamins, with the lowest concentrations required tomeet daily requirements. Vitamin B12 is unique in that it is synthesizedin nature only by microorganisms; therefore, it is usually not found inplant feedstuffs. Consequently, humans and other monogastric species
who subsist entirely on plant foods would be susceptible to vitamin B12
deficiency. It is also unique in that the trace element cobalt is an integral
part of the molecule.
The discovery of this vitamin was dramatic and made possible by
the combined efforts of microbiologists, biochemists, nutrition scientists, and physicians working in various laboratories. Three seemingly
unrelated conditions attributed to lack of the vitamin or its precursor
were identified: (1) a fatal anemia in humans, (2) a potent growth factor for monogastric species, and (3) a relationship to cobalt, the lack ofwhich resulted in wasting diseases in ruminants.

HISTORY
The history of vitamin B12 in human and animal nutrition is both
exciting and stimulating and has been reviewed (Sebrell and Harris,
1968; Folkers, 1982; Loosli, 1991). In 1824 Combe described a fatal
anemia, pernicious anemia, and suggested that it could be related to adisorder of the digestive tract. The existence of an unknown factor inliver, effective in treatment of pernicious anemia, was recognized in
1926 when Minot and Murphy showed that large amounts (120 to 240
g/day) of raw liver given by mouth daily would alleviate this previously
523

13

CHAPTER THIRTEEN

fatal disease. In 1920 Whipple provided liver in diets to dogs andshowed regenerated blood and a specific liver protein that was needed
for the formation of hemoglobin. Minot, Murphy, and Whipple received
the Nobel Prize in 1934 for liver therapy of pernicious anemia. Twoyears after the Nobel Prize was awarded it became apparent that the reason the dogs responded to liver was the iron content of the liver and not
the vitamin B12 content.
During the next 20 years following the Minot and Murphy discovery, research resulted in concentrating the activity of 400 g of liver to 1
mg of active substance. From 1929 onward, Castle postulated that pernicious anemia was due to the interaction of a dietary (extrinsic) factor
and an intrinsic factor produced by the stomach. Mixing beef muscle
and gastric juice prevented anemia; thus a factor in gastric juice was theintrinsic factor, while a different substance in beef muscle was the extrinsic factor. Castle took the extraordinary step of using his own stomach to process food for his anemic patients, then regurgitated his partially digested meals to supplement their diets.
For many years following the discovery that liver contained asubstance that could cause remission of pernicious anemia, scientists
tried unsuccessfully to isolate from liver the antipernicious anemia(APA) factor. Progress was slow because no experimental animal forlaboratory trials exhibited this condition, thus human patients withpernicious anemia were required for experimental studies. In 1947
Shorb of the University of Maryland reported that a factor (LLD factor) in liver extract required by the bacterium Lactobacillus lacatis
Dorner was in concentrations bearing an almost linear relationship to
the APA activity of the extract. Making use of this organism, Rickes
and coworkers in the United States in 1948, isolated, in crystalline
form, a factor from the liver that cured pernicious anemia. They
named this factor vitamin B12. Three weeks after the Rickes andcoworkers publication, a similar article from Smith of Great Britain
also indicated isolation of the APA factor. West (1948) confirmed theclinical activity of the vitamin, which prevented pernicious anemia
with a single dose of 3 to 6 μg. During these early years of isolating
the APA factor, the production of 15 mg of crystalline vitamin B12 required 1,000 kg of fresh liver.
Although isolated by the two laboratories in 1948, it was not until
1956 that the complicated vitamin B12 structure was ascertained. In
1961 Lenhert and Hodgkin reported the structure of the enzyme formof vitamin B12. In 1964 another Nobel Prize was awarded to Hodgkin524

V I T A M I N B1 2

for her part in the elucidation of the chemical structure of vitamin B12by x-ray crystallography.
Attempts to raise pigs and chickens by feeding all vegetable diets resulted in poor performance. In 1926 it was recognized that liver extracts
and other concentrates of animal origin stimulated growth of rats,
chicks, and pigs. Because the true nature of the active principle of suchanimal products was unknown, it was called animal protein factor(APF). It was also referred to as the chick growth factor, and the same
factor was found essential for hatchability. Manure from cattle (cowmanure factor) was also found to contain the factor. The APF found inanimal excrement was particularly potent in the summer, when feces
were rich in bacteria. This led to the isolation of a microorganism fromchicken feces and its cultivation in a simple fermentation medium. The
discovery of microbial production of vitamin B12 opened the way to economical industrial production of vitamin B12, which is based entirely on
bacterial fermentation.
Vitamin B12 became available for animal experiments in 1948 as the
crystalline vitamin B12 was isolated from liver and crude fermentation
products, which contained considerable concentrations of the vitamin.
To the surprise of researchers, the growth effect of pure vitamin B12 inanimals was not quite as good as that observed from APF supplements.
Eventually, it was found that crude vitamin B12 concentrates contained
more than one active principle, namely, vitamin B12, some essential
amino acids, and small concentrations of compounds with antibiotic activity. The growth-promoting effects of antibiotics were discovered practically as a by-product of research on vitamin B12 in animal nutrition.
These discoveries were important considerations to providing complete
confinement for certain classes of livestock.
The significance of vitamin B12 for ruminants was discovered to be
the requirement of cobalt by rumen microorganisms in order to synthesize the vitamin (McDowell, 1985, 1992). Cobalt, the central ion in vitamin B12, was shown to be a dietary essential for sheep in 1935 by Underwood and others in Australia (Underwood, 1977). The Australians
showed that the deficiency caused debilitating diseases of sheep known
as “coast disease” and “wasting disease.” In Florida in 1937, cobalt deficiency was reported by Becker and coworkers (1965) to be responsible
in part for “salt sick” cattle, describing a severe wasting disease. In
1951, Smith and coworkers at Cornell discovered that injections of vitamin B12 prevented all signs of cobalt deficiency.

525

CHAPTER THIRTEENCHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Vitamin B12 is now considered by nutritionists as the generic namefor a group of compounds having B12 activity. These compounds havevery complex structures. The empirical formula of B12 isC63H88O14N14PCo, and among its unusual features is the content of
4.5% cobalt. The structure of one B12 compound, cyanocobalamin, isshown in Fig. 13.1. Vitamin B12 resembles a porphyrin structure consisting of four pyrrole nuclei coupled directly to each other, with the inner nitrogen atom of each pyrrole coordinated with a single atom ofcobalt. The basic tetrapyrrole structure is the corrin nucleus, which positionally is a planar structure coupled below to the nucleotide 5,6-dimethylbenzimidazole and above to cyanide or some other derivative.
The large ring formed by the four reduced rings is called “corrin” because it is the core of the vitamin. The vitamin belongs to the corrinoidgroup of compounds that have a corrin nucleus; however, numerous
other corrinoids do not possess vitamin B12 activity. The name “cobalamin” is used for compounds in which the cobalt atom is in the centerof the corrin nucleus.
In vitamin B12 the base is coupled directly to the cobalt atom, and an
ester linkage from the phosphate group of the nucleotide to the propionicacid group of the D ring of the corrin nucleus adds further stability to themolecule. Cyanide, which lies above the planar ring, is attached to thecobalt atom—thus the name cyanocobalamin. The cyanide can be replaced by other groups, including OH (hydroxycobalamin), H2O (aquacobalamin), NO2 (nitrocobalamin), and CH3 (methylcobalamin). All thesecompounds are referred to as cobalamins and have activity. In addition,
several other compounds, referred to as “pseudo” vitamin B12 complexes
or vitamin B12-like factors that have some activity, have been isolated orsynthesized. Their structure differs regarding the nucleotide moiety. These
pseudovitamins are probably intermediates of the biosynthesis of vitamin
B12 and are found in sewage, manure, rumen contents, and residues fromfermentation. Some of these pseudovitamin B12 compounds are analogs
without a nucleotide, while others contain a nucleotide other than 5,6-dimethylbenzimidazole. Some analogs have incorporated other metals (e.g.,
Cu, Ni, Mn, Fe) into them, but without corrinoids have no biological activity. Some of the corrinoid pseudovitamins that are growth factors for
microorganisms not only have no vitamin B12 activity, but may be antagonists or antimetabolites of the vitamin.
526

V I T A M I N B1 2

Fig. 13.1 Structure of vitamin B12 (cyanocobalamin).

The isolation of coenzyme forms of vitamin B12 led to the recognition that cyanocobalamin is not the naturally occurring form of the vitamin but is rather an artifact that arises from the original isolation procedure, which absorbed the vitamin by activated charcoal.
Adenosylcobalamin, hydroxocobalamin, methylcobalamin, cyanocobalamin, and sulfitocobalamin have been determined in feedstuffs, with thefirst three being the most predominant forms in animal tissue (Farquharson and Adams, 1976).
The two coenzyme forms of cobalamin found in animals are adenosylcobalamin and methylcobalamin. Cyanocobalamin, however, is themost widely used form of cobalamin in clinical practice because of its
relative availability and stability (particularly to light). Most metabolicstudies utilize cyanocobalamin. Hydroxocobalamin is also used in pharmaceutical preparations, and is better retained after parenteral administration than is cyanocobalamin.
527

CHAPTER THIRTEEN

Vitamin B12 is a dark red, crystalline, hygroscopic substance, freely
soluble in water and alcohol but insoluble in acetone, chloroform, or
ether. Cyanocobalamin has a molecular weight of 1354 and is the mostcomplex structure and heaviest compound of all the vitamins. Oxidizing
and reducing agents and exposure to sunlight tend to destroy its activity. Losses of vitamin B12 during cooking are usually not excessive.

ANALYTICAL PROCEDURES
Several techniques are employed for analyses of vitamin B12. Chemical assays, including spectrophotometric and colorimetric procedures,
have been developed for pharmaceutical preparations but are not sensitive enough for determination of the vitamin in natural materials (Scottet al., 1982). Colorimetric procedures rely on measurement of cyanide
released or a color complex with 5,6-dimethyl-benzimidazole.
Microbiological assays for vitamin B12 are sensitive and can be applied to crude materials. Microbiological methods using L. leichmannii
can determine quantities less than 0.01 μg of the vitamin per milliliter ofassay solution. The organism responds, however, to deoxyribonucleosides and to several B12 pseudovitamins. Treatment of the sample withalkali destroys vitamin B12, leaving the deoxyribonucleosides intact; thus
the vitamin B12 plus pseudo forms can be determined by difference.
Also, response to deoxyribonucleosides is less serious, since L. leichmannii requires about 1,000 times more deoxyribonucleosides than vitamin B12 for growth.
The protozoans Euglena gracilis and Ochromonas malhamensis arealso successfully used for vitamin B12 determination. Vitamin B12 assays
involving animals are somewhat more difficult and time-consuming than
microbiological assays. Large stores of vitamin B12 found in young,growing animals reared from normal mothers present the biggest problem (Ellenbogen and Cooper, 1991). A biological assay for vitamin B12
uses growth of chicks hatched from eggs of vitamin B12-deficient hens,
and assays have been conducted with young rats born from depleted
dams. Thyroid-stimulating material often is added to assay diets to increase the B12 requirement in the young animal.
The radioisotope dilution methods for the assay of cobalamins havebeen replacing the microbiological methods and are used more widely
than most other methods. These assays measure the extent to which
cobalamin, after first being liberated from bound materials, competes
with radioactively labeled cyanocobalamin for binding sites on a protein(Ellenbogen and Cooper, 1991).
528

V I T A M I N B1 2

Ramoz et al. (1994) reported that the radioassay method was as reliable as the microbiological method for measuring vitamin B12 in ovine
milk, except for very low levels. A radioimmunoassay was found not todetect vitamin B12 concentration in plasma samples containing elevatedconcentrations of methylmalonic acid (Kennedy et al., 1992b). Muhammad et al. (1993) compared a competitive binding assay with a microbiological assay and found a high correlation between methods for estimating B12 concentrations in foods.

METABOLISM
Digestion, Absorption, and Transport
Passage of vitamin B12 through the intestinal wall requires intervention of certain carrier compounds able to bind the vitamin molecule. Vitamin B12 in the diet is bound to food proteins. In the stomach, the combined effect of gastric acid and peptic digestion releases the vitamin,
which is then bound to a nonintrinsic factor–cobalamin complex(Toskes et al., 1973). The nonintrinsic protein that is secreted in thesaliva has been named cobalophilin, formerly known as R-proteins, because of their rapid electrophoretic mobility compared with other cobalamin-binding proteins.
Vitamin B12 is bound preferentially to cobalophilin in the acidmedium of the stomach rather than to intrinsic factor. The B12 remains
bound to cobalophilin in the slightly alkaline environment of intestine
until pancreatic proteases (e.g., trypsin) partially degrade thecobalophilin protein and thereby enable B12 to become bound exclusively to intrinsic factor. Therefore, patients with pancreatic insufficiency absorb B12 poorly (Jorgensen et al., 1991), and this malabsorption is completely corrected by administration of pancreatic enzymes orpurified trypsin.
A prerequisite for intestinal absorption of physiological amounts ofcobalamin is binding to intrinsic factor. Intrinsic factor is a glycoprotein(mucoprotein) synthesized and secreted by parietal cells of the gastricmucosa. Atrophy of the fundus, where intrinsic factor is produced, andlack of free HCl (achlorhydria) are usually associated with pernicious
anemia (Behrns et al., 1994). Gastric juice defects are responsible formost cases of food-vitamin B12 malabsorption (Carmel, 1994). The formation of this intrinsic factor complex protects the vitamin from bacterial utilization and/or degradation as it traverses the lumen of the small
intestine to the terminal ileum, where absorption occurs (Ellenbogen529

CHAPTER THIRTEEN

and Highley, 1970). The intrinsic factor–B12 complex is transiently attached to an ileal receptor. The proximal small intestine does not havethe ability to enhance absorption of the vitamin—only the ileum has thisproperty. In the ileum, the intrinsic factor moiety of the intrinsic factor–B12 complex binds to a specific receptor protein on the microvillus
membrane of brush borders of intestinal epithelial cells. Next there istransport of vitamin B12 from the receptor intrinsic factor–B12 complex
through the epithelial cell to portal blood.
The absorption of vitamin B12 is limited by the number of intrinsic
factor–vitamin B12 binding sites in the ileal mucosa, so that not morethan about 1 to 1.5 μg of a single oral dose of the vitamin in humans
can be absorbed (Bender, 1992). The absorption is also slow; peak bloodconcentrations of the vitamin are not achieved for some 6 to 8 hours after an oral dose.
When B12 enters the portal blood, it is no longer bound to intrinsic
factor but to specific transport proteins called transcobalamins. Threebinding proteins have been identified in normal human serum and aredesignated as transcobalamin I, II, and III. The transcobalamins are synthesized by several tissues, including intestinal mucosa and liver, andhave been shown to deliver B12 to various tissues, such as liver, kidney,spleen, heart, lung, and small intestine (Rothenberg and Cotter, 1978).
Transcobalamin II appears to be primarily concerned with transport ofvitamin B12, whereas transcobalamin I is involved in storage of the vitamin.
The function of transcobalamin III is to provide a mechanism for returning vitamin B12 from peripheral tissues to the liver, as well as forclearance of other corrinoids without vitamin activity (e.g., undesired
analogs of B12), which may arise either from foods or from the products
of intestinal bacterial action and be absorbed passively across the lowergut (Bender, 1992). These corrinoids are then secreted into the bile,
bound to cobalophilins. Like dietary vitamin B12 bound to salivary
cobalophilin, the biliary cobalophilins are hydrolyzed in the duodenum,and the released vitamin B12 binds to intrinsic factor, permitting reabsorption into the ileum.
To summarize, B12 absorption for most species studied requires thefollowing: (1) adequate quantities of dietary B12, (2) normal stomach forbreakdown of food proteins for release of B12, (3) normal production ofcobalophilin (nonintrinsic factor) secreted in saliva, (4) normal stomach
for production of intrinsic factor for absorption of B12 through theileum, (5) normal pancrease (trypsin) required for release of bound B12530

V I T A M I N B1 2prior to combining the vitamin with the intrinsic factor, and (6) normal
ileum with receptor and absorption sites. Additional factors that diminish vitamin B12 absorption include deficiencies of protein, iron, and vitamin B6; thyroid removal; and dietary tannic acid (Anonymous, 1984).
Intrinsic-factor concentrates prepared from one animal’s stomach
do not in all cases increase B12 absorption in other species or in humans.
There are structural differences in the B12 intrinsic factor among species.
Likewise, species differences exist for B12 transport proteins (Polak etal., 1979). Intrinsic factor has been demonstrated in the human, monkey, dog, pig, rat, cow, ferret, rabbit, hamster, fox, lion, tiger, and leopard. It has not been detected in the guinea pig, horse, sheep, chicken, ora number of other species. The dog stomach produces only small
amounts of intrinsic factor, with larger amounts produced by the pancreas (Simpson et al., 1989).
Absorption of vitamin B12 does not completely depend on active intervention of the intrinsic factor. Both active and passive mechanisms exist for absorption of B12 (Herbert, 1990). The passive mechanism, simple diffusion, has low efficiency (approximately 1%) and is operative
throughout the digestive tract; it becomes practically important only inthe presence of large quantities of the vitamin, in excess of those present
in most foods.
About 3% of ingested cobalt is converted to vitamin B12 in the rumen. Of the vitamin B12 produced, only 1 to 3% is absorbed. In the rumen, iron interacts with cobalt so that iron deficiency enhances cobalt
absorption (Keen and Graham, 1989). As in most species, the absorptivesite for ruminants is the lower portion of the small intestine. Substantial
amounts of B12 are secreted into the duodenum and then reabsorbed in
the ileum.

Tissue Distribution and Storage
In normal human subjects, vitamin B12 is found principally in theliver; the average amount is 1.5 mg. Kidneys, heart, spleen, and brain
each contain about 20 to 30 μg (Ellenbogen and Cooper, 1991). Vitamin
B12 is stored in the liver in the largest quantities for most animals thathave been studied, but it is stored in the kidney of the bat. Vitamin B12
storage in humans can exceed the daily requirement by about 1,000fold.
Henderickx et al. (1964) reported a total retention of 20 to 23% of
an oral dose of vitamin B12 in pigs, about two-thirds of which was present in the liver. Even though vitamin B12 is a water-soluble vitamin, there

531

CHAPTER THIRTEEN

is a considerable degree of tissue storage. The great storage and long biological half-life (350 to 400 days in humans) of the vitamin provides
substantial protection against periods of deprivation.
To become metabolically active, vitamin B12 must be converted into
one of its various coenzyme forms. This transformation takes placemainly in liver but also in kidneys. Most of the cobalamins in humans
occur as two coenzymatically active forms, adenosylcobalamin andmethylcobalamin. In humans, methylcobalamin constitutes 60 to 80%
of total plasma cobalamin, while adenosylcobalamin is the major cobalamin in all cellular tissues, constituting about 60 to 70% in the liver andabout 50% in other organs (Ellenbogen and Cooper, 1991). In subjects
with pernicious anemia, methylcobalamin is disproportionately reduced
in relation to the other major components. Cyanocobalamin is converted within cells to either methylcobalamin, a coenzyme for methyltransferase, or adenosylcobalamin, the coenzyme for mutase.

Excretion
The main excretion of absorbed vitamin B12 is via urinary, biliary,and fecal routes. Total body loss ranges from 2 to 5 μg daily in humans(Shinton, 1972). Urinary excretion of the intact vitamin B12 by kidney
glomerular filtration is minimal. Biliary excretion via feces is the major
excretory route. Approximately 0.5 to 5 μg of cobalamin is secreted into
the alimentary tract daily, mainly in bile (Ellenbogen and Cooper, 1991).
Most of the cobalamin excreted in bile is reabsorbed; at least 65 to 75%
is reabsorbed in the ileum by means of the intrinsic factor, active transport mechanism.

FUNCTIONS
Vitamin B12 is an essential part of several enzyme systems that carryout a number of very basic metabolic functions. Specific biochemical reactions in which cobalamin coenzymes participate are of two types: (1)those that contain 5´-deoxyadenosine linked covalently to the cobalt
atom (adenosylcobalamin) and (2) those that have a methyl group attached to the central cobalt atom (methylcobalamin). A number of vitamin B12-dependent metabolic reactions have been identified in microorganisms; however, only three vitamin B12-dependent enzymes have been
discovered in animals: methylmalonyl CoA mutase and leucine mutase,which each require adenosylcobalamin, and methionine synthetase,which requires methylcobalamin. Most reactions requiring adenosylcobalamin can be classified as rearrangement reactions of the carbon
532

V I T A M I N B1 2

skeleton of several metabolic intermediates; a hydrogen atom moves
from one carbon atom to an adjacent one in exchange for an alkyl, acyl,
or electronegative group, which migrates in the opposite direction (Ellenbogen and Cooper, 1991). In all these rearrangement reactions,
adenosylcobalamin is an intermediate hydrogen carrier. The reactions
requiring methylcobalamin involve transfer or synthesis of one-carbon
units, for example, methyl groups.
Vitamin B12 is metabolically related to other essential nutrients, suchas choline, methionine, and folacin (Savage and Lindenbaum, 1995). Interrelationships of these nutrients with vitamin B12 (and in particular totransmethylation and biosynthesis of labile methyl groups) are discussed
in Chapters 12 and 14. Though the most important tasks of vitamin B12
concern metabolism of nucleic acids and proteins, it also functions inmetabolism of fats and carbohydrates. A summary of B12 functions
would include (1) purine and pyrimidine synthesis, (2) transfer of methylgroups, (3) formation of proteins from amino acids, and (4) carbohydrate and fat metabolism. A general function of B12 is to promote red
blood cell synthesis and to maintain nervous system integrity, which arefunctions noticeably affected in the deficient state.
Vitamin B12 is necessary in reduction of one-carbon compounds offormate and formaldehyde, and in this way it participates with folacin
in biosynthesis of labile methyl groups. Formation of labile methylgroups is necessary for biosynthesis of purine and pyrimidine bases,
which represent essential constituents of nucleic acids. Disorders of nucleic acid synthesis in vitamin B12 deficiency are connected with this. The
purine bases (adenine and guanine) as well as thymine are constituentsof nucleic acids, and with folacin deficiency there is a reduction inbiosynthesis of nucleic acids essential for cell formation and function.
Hence, deficiency of either folacin or B12 leads to impaired cell division
and alterations of protein synthesis; these effects are most noticeable inrapidly growing tissues.
Deficiency of B12 will induce folacin deficiency by blocking utilization of folacin derivatives. A vitamin B12-containing enzyme removes the
methyl group from methylfolate, thereby regenerating tetrahydrofolate(THF), from which is made the 5,20-methylene-THF required forthymidylate synthesis. Because methylfolate returns to the body’s folacinpool only via the vitamin B12-dependent step, vitamin B12 deficiency results in folacin being “trapped” as methylfolate, and thus becoming
metabolically useless. The “folate trap” concept explains why hematological damage of vitamin B12 deficiency is indistinguishable from thatof folacin deficiency by alleging that in both instances the defective syn533

CHAPTER THIRTEEN

thesis of DNA results from the same final common pathway defect,
namely, an inadequate quantity of 5,10-methylene-THF to participate
adequately in DNA synthesis (Herbert and Zalusky, 1962).
Metabolism of labile methyl groups plays a significant part inbiosynthesis of methionine from homocysteine. A vitamin B12-requiring
enzyme, 5-methyl-tetrahydrofolate-homocysteine methyltransferase,
catalyzes reformation of methionine from homocysteine according to
the following reaction:
5-Methyltetrahydrofolate + homocysteine ♦ℜ methionine +tetrahydrofolate
The mechanism of converting homocysteine to methionine has utilized amethyl group from folacin, the mechanism for maintaining folacin in a
reduced form. Activity of this enzyme is depressed in liver of vitamin
B12-deficient sheep (MacPherson, 1982), which could lead to a deficiency of available methionine that may account for impairment of nitrogen metabolism in vitamin B12-deficient sheep.
Overall synthesis of protein is impaired in vitamin B12-deficient animals. Wagle et al. (1958) demonstrated that rats and baby pigs deprivedof vitamin B12 were less able to incorporate serine, methionine, phenylalanine, and glucose into liver proteins. There is good reason to believethat impairment of protein synthesis is the principal reason for thegrowth depression that is frequently observed in animals deficient in vitamin B12 (Friesecke, 1980).
In the metabolism of animals, propionate of dietary or metabolic
origin is converted into succinate, which then enters the tricarboxylic
acid (Krebs) cycle. Because propionate is a three-carbon and succinate afour-carbon compound, this process requires the introduction of a onecarbon unit. Methylmalonyl-CoA isomerase (mutase) is a vitamin B12requiring enzyme (5´-deoxyadenosylcobalamin) that catalyzes the conversion of methylmalonyl-CoA to succinyl-CoA. Flavin and Ochoa(1957) established that for succinate production, the following steps are
involved:
Propionate + ATP + CoA ♦ propionyl-CoA
Propionyl-CoA + CO2 + ATP ♦ methylmalonyl-CoA (a)Methylmalonyl-CoA (a) ♦ methylmalonyl-CoA (b)Methylmalonyl-CoA (b) ♦ succinyl-CoA
Methylmalonyl-CoA (a) is an inactive isomer. Its active form (b) is con534

V I T A M I N B1 2verted into succinyl-CoA by a methylmalonyl isomerase, or methylmalonyl mutase (fourth reaction).
Vitamin B12 is a metabolic essential for all animal species studied,and vitamin B12 deficiency can be induced with the addition of high dietary levels of propionic acid. However, metabolism of propionic acid isof special interest in ruminant nutrition because large quantities are produced during carbohydrate fermentation in the rumen. Propionate production proceeds normally, but in cobalt or vitamin B12 deficiency, its
rate of clearance from blood is depressed, and methylmalonyl-CoA accumulates. This results in increased urinary excretion of methylmalonicacid and in loss of appetite because impaired propionate metabolism
leads to higher blood propionate levels, which are inversely correlated tovoluntary feed intake (MacPherson, 1982). Injection of cobalt-deficientanimals with vitamin B12 produces overnight improvement in appetite,whereas oral dosing with cobalt takes 7 to 10 days to produce the same
effect.
A further important function of vitamin B12 in intermediary metabolism consists of maintaining glutathione and sulfydryl groups of enzymes in the reduced state (Marks, 1975). The reduced activity of glyceraldehyde-3-phosphate dehydrogenase, which needs glutathione as acoenzyme, is possibly responsible for carbohydrate metabolism being
impaired in vitamin B12 deficiency. Vitamin B12 also influences lipid metabolism via its effect on the thiols.

REQUIREMENTS
Vitamin B12 requirements are exceedingly small; an adequate allowance is only a few micrograms per kilogram of feed, making B12 themost potent of vitamins. Estimated requirements of vitamin B12 for various animals and humans are presented in Table 13.1.
The vitamin B12 requirements of various species depend on the levels of several other nutrients in the diet. Excess protein increases theneed for B12 as does performance level (NRC, 1994). The B12 requirement depends on levels of choline, methionine, and folacin in the dietand is interrelated with ascorbic acid metabolism (Scott et al., 1982).
The requirements for both vitamin B12 and folacin are reduced when thediet contains an abundance of compounds that can supply methylgroups. Sewell et al. (1952) showed that B12 has a sparing effect on themethionine needs of the pig. A reciprocal relationship occurs between
B12 and pantothenic acid in chick nutrition, with pantothenic acid sparing the B12 requirement. Dietary ingredients may also affect the require535

CHAPTER THIRTEEN■ Table 13.1
AnimalBeef cattle
Dairy cattle
ChickenJapanese quail
Turkey
Sheep
Swine
Horse
Goat
Dog
Cat
Mink
RabbitFish
Rat
Hamster
Human

Vitamin B12 Requirements for Various Animals and Humans
Purpose or Class

Requirementa

Reference

Adult
Calf (milk replacer)
Adult
Leghorn, 0–6 weeks
Leghorn, 6–18 weeks
Laying (100-g intake)
Broilers, 0–6 weeks
Broilers, 6–8 weeks
All classes
All classes
Adult
Growing-finishing
Breeding-lactating
Adult
Adult
Growing
Growing
All classes
All classes
Pacific salmon
Yellowtail
Growing
Growing
Infants
Children
AdultsPregnancy-lactation

Microbial synthesisb

NRC (1996)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1981)
NRC (1985a)
NRC (1986)
NRC (1982a)
NRC (1977)
NRC (1993)
NRC (1993)
NRC (1995)
NRC (1995)
RDA (1989)
RDA (1989)
RDA (1989)
RDA (1989)

0.07 ppm
Microbial synthesisb
9 μg/kg
3 μg/kg
4 μg/kg
10 μg/kg
7 μg/kg
3 μg/kg
3 μg/kg
Microbial synthesisb
5–20 μg/kg
15 μg/kg
Microbial synthesis
Microbial synthesisb
26 μg/kg
20 μg/kg
30 μg/kg
Microbial synthesis
15–20 μg/kg
53 μg/kg
50 μg/kg
10 μg/kg
0.3–0.5 μg/day
0.7–1.4 μg/day
2 μg/day
2.2–2.6 μg/day

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b). Human data are expressed as μg/day.
bOnly young ruminants have a dietary need for B
12 prior to ruminal development.
Practical vitamin B12 deficiency is a secondary result of cobalt deficiency. Suggested cobalt
requirements for ruminants range from 0.07 to 0.2 mg/kg of diet.

ment, as wheat bran has been known to reduce availability of vitamin
B12 in humans (Lewis et al., 1986).
Dietary need depends on intestinal synthesis and tissue reserves atbirth. Intestinal synthesis probably explains frequent failures to produce
B12 deficiency in pigs and rats on diets designed to be B12 free. The deficiency can be readily produced in rats, however, when coprophagy iscompletely prevented (Barnes and Fiala, 1958). Coprophagous animals
and poultry on deep litter receive excellent supplies of B12 from microbial fermentation. Litter would be a less valuable source of vitamin B12
under cold conditions, in which bacterial numbers are greatly reduced.
Poultry obtain some vitamin B12 by direct absorption of the vitamin produced by bacterial synthesis in the intestine (NRC, 1994), however, the536

V I T A M I N B1 2amount from this source is not reliable.
Intestinal microfloral synthesis of vitamin B12 has been demonstrated in common carp, channel catfish, Nile tilapia, rainbow trout,ayu, and goldfish (NRC, 1993). Sugita et al. (1991) found a close relationship between the amount of vitamin B12 and the viable counts of
Bacteroides type A in the intestinal contents of the various fish studied.
They found that this bacterium was present in the intestinal contents offish that do not require vitamin B12 and was absent in fish that do require the vitamin.
Requirement for vitamin B12 in ruminant diets is closely associatedwith the requirement for cobalt since this trace mineral is a componentof the B12 molecule. Ruminant animals have the ability to synthesize vitamin B12 provided they are supplied with an adequate dietary supply ofcobalt (0.07 to 0.2 ppm) and have a normally functioning rumen. Under
typical conditions a rumen would be functional for synthesis of all B vitamins at 6 to 8 weeks of age. Therefore, only young ruminants that do
not have a fully developed rumen would be expected to require a dietary
source of B12.
Cobalt content of the diet is the primary limiting factor for synthesis of vitamin B12 by ruminal microflora. However, studies indicate thatsynthesis of vitamin B12 could be restricted even when the diet is adequate in cobalt, as several factors can influence synthesis and perhaps
utilization. For animals on high-concentrate diets, there is a decrease invitamin B12 synthesis, and more analogs are produced than the vitaminitself (Sutton and Elliot, 1972). These natural analogs have little or novitamin B12 activity. Lopez-Guisa and Satter (1992) suggested thatcobalt and copper requirements higher than NRC recommendations
may aid in digestibility of low-quality forages (i.e., corn crop residues),
but cobalt was without effect for digestibility of high-quality forages(Hussein et al., 1994).
The established dietary cobalt requirements for ruminants are 0.1 to
0.2 mg/kg of diet (NRC, 1985b, 1989a, 1996). It has been suggested
that the requirement for vitamin B12 in dairy cattle is between 0.34 and0.68 μg/kg of live weight (NRC, 1989a). Precise estimates of minimum
cobalt requirements are difficult because of the influence of many variables, such as seasonal changes in herbage cobalt concentrations, selective grazing habits, and soil contamination. Under grazing conditions,lambs are the most sensitive to cobalt deficiency, followed by mature
sheep, calves, and mature cattle (Andrews, 1956).
Ruminants have higher vitamin B12 requirements than nonruminants, presumably because of its involvement in the metabolism of pro537

CHAPTER THIRTEEN

pionic acid. Vitamin B12 is essential as a cofactor for methyl-malonylCoA isomerase, an enzyme necessary for propionic acid utilization thatis produced in much greater quantities in ruminants. Experiments withsheep suggest an oral requirement for growing lambs of some 200
μg/day, about 10 times the reported oral requirement of other species
per unit of food intake (Marston, 1970). The surprisingly high requirement of ruminants for cobalt arises partly from the low efficiency of production of vitamin B12 from cobalt by the rumen microorganisms andpartly from the low efficiency of absorption of vitamin B12.
It has been reported that grass in healthy areas contains around 0.1
mg of cobalt per kg (dry basis) of cobalt or more, on the average, ascompared with 0.05 to 0.07 mg/kg for deficient areas. As little as 0.1
mg/kg has restored sick animals to health. Further evidence of 0.1 mg/kg
as the dietary requirement for sheep was provided by Mohammed(1983), who fed various levels of cobalt and found that vitamin B12 andpropionic acid in rumen concentrations were maximal at 0.1 mg ofcobalt per kg.
Vitamin B12 requirements for humans have been estimated fromthree different types of studies (Ellenbogen and Cooper, 1991): (1) determination of the amount necessary to treat megaloblastic anemia fromvitamin B12 deficiency, (2) comparison of blood and liver concentrationin normal and cobalamin-deficient subjects, and (3) body stores andturnover rates of the vitamin. Obviously, the requirement for B12 will be
substantially higher for humans lacking intrinsic factor or other conditions that affect absorption and metabolism of the vitamin (see Metabolism).
For adults, recommendations vary from 1 to 2 μg of vitamin B12
daily. Higher vitamin B12 requirements would be for pregnancy; during
the last half, the fetus removes 0.2 μg daily from maternal stores, andduring lactation, 0.3 μg is lost daily in breast milk (Herbert, 1990).

NATURAL SOURCES
The origin of vitamin B12 in nature appears to be microbial synthesis. It is synthesized by many bacteria but apparently not by yeasts or bymost fungi. There is little evidence that the vitamin is produced in tissuesof higher plants or animals. A few reports have suggested limited B12synthesis by a few plants, but in insignificant quantities in relation to animal requirements. Synthesis of this vitamin in the alimentary tract is of
considerable importance for animals; if sufficient cobalt is available, ruminants are independent of external sources of vitamin B12.
538

V I T A M I N B1 2
■ Table 13.2

Vitamin B12 Concentrations of Various
Foods and Feedstuffs (ppb, dry basis)

Blood meal
Corn, grain
Crab meal
Distiller’s solubles
Fish solubles
Fish meal, anchovy
Fish meal, herring
Fish meal, menhaden
Fish meal, tuna
Horse meat
Liver meal
Meat meal
Milk, skim, cow’s
Poultry by-product meal
Soybean meal
Spleen, cow
Wheat, grain
Whey, cow
Yeast

49
0
475
3
1007
233
467
133
324
142
542
72
54
322
0
247
1
20
1

Source: NRC (1982b).

Foods of animal origin are reasonably good sources, including meat,
liver, kidney, milk, eggs, and fish (Table 13.2). Milk and milk products
are good sources of vitamin B12, but the vitamin content is reduced considerably in whey (Sato et al., 1997). Kidney and liver are excellent
sources, and these organs are richer in vitamin B12 from ruminants thanfrom most nonruminants. In one study, goat milk was found to containsignificantly less vitamin B12 than cow milk (Dostalova, 1994). Vitamin
B12 presence in tissues of animals is due to the ingestion of vitamin B12
in animal foods or from intestinal or ruminal synthesis. Among the richest sources are fermentation residues, activated sewage sludge, and manure.
Plant products are practically devoid of vitamin B12. The vitamin
B12 reported in higher plants in small amounts may result from synthesis by soil microorganisms and excretion of the vitamin into soil, withsubsequent absorption by the plant. Root nodules of certain legumes
contain small quantities of B12. Certain species of seaweed (algae) havebeen reported to contain appreciable quantities of vitamin B12 (up to 1
μg/g of solids). Seaweed does not synthesize vitamin B12, but it is synthesized by the bacteria associated with seaweed and then concentrated
by the seaweed (Scott et al., 1982).
Little is known about the bioavailability of orally ingested B12 in
539

CHAPTER THIRTEEN

foods and feeds. Dagnelie et al. (1991) reported that vitamin B12 fromalgae is largely unavailable. Providing algae to B12-deficient children was
ineffective in elevating blood parameters compared to B12 from fish
sources. Vitamin B12 was found to be more available in pasteurized milk
(1.91 pmol/ml) than in raw milk (1.54 pmol/ml), sterilized milk (1.25
pmol/ml), and dried milk (1.27 pmol/ml), indicating that heat treatment
probably affects the bioavailability (Fie et al., 1994).
To supply the ruminant with vitamin B12, the important consideration is dietary sources of cobalt. Although most feeds are adequate incobalt, the element is deficient in forages for the grazing ruminant inmany parts of the world (McDowell, 1985, 1992). Concentration ofcobalt in crops and forages is dependent on soil factors, plant species,
stage of maturity, yield, pasture management, climate, and soil pH. Soilcontaining less than 2 mg/kg of cobalt is generally considered deficient
for ruminants (Corrêa, 1957). Raising the pH by liming reduces thecobalt uptake by the plant and may increase the severity of the deficiency. Plants grown on 15-ppm cobalt soil that is neutral or slightly
acid may contain more cobalt than those grown on 40-ppm cobalt alkaline soil (Latteur, 1962). High rainfall tends to leach cobalt from thetopsoil. This problem is often aggravated by rapid growth of forage during the rainy season, which dilutes the cobalt content. Plants have varying degrees of affinity for cobalt, some being able to concentrate the element much more than others. Legumes, for example, generally havegreater ability to concentrate cobalt than do grasses (Underwood,1977). Cobalt is needed by the N-fixing bacteria in the root nodules oflegumes.

DEFICIENCY
The result of vitamin B12 deficiency in humans is megaloblastic anemia (pernicious anemia) and neurological lesions. Vitamin B12 deficiency
in humans usually is conditioned by a deficiency of intrinsic factor necessary for its absorption or is found in humans consuming strict vegetarian diets. In animals, pernicious anemia—or in fact any anemia—is
not characteristic of vitamin B12 shortage. In rats, guinea pigs, swine,
and poultry, vitamin B12 functions as a growth factor, although mild
anemia does occur in a small percentage of deficient swine. In ruminants, vitamin B12 deficiency is closely associated with their requirement
for cobalt, since the trace mineral is a component of the B12 molecule.
In all species, as vitamin B12 deficiency progresses, depletion of vitamin
B12 (and cobalt) occurs concurrently in serum and tissue reserves.
540

V I T A M I N B1 2

Effects of Deficiency
Ruminants
Vitamin B12 deficiency can occur in young ruminants as long as the
microflora of the forestomachs is not yet far enough developed andhence unable to furnish sufficient amounts of the vitamin. Lassiter et al.
(1953) demonstrated vitamin B12 deficiency in calves less than 6 weeks
old that received no dietary animal protein. Clinical signs characterizing
the deficiency included poor appetite and growth, muscular weakness,demyelination of peripheral nerves, and poor general condition. Younglambs (up to 2 months of age), if weaned early, likewise had a need for
dietary vitamin B12 (NRC, 1985b). In vitamin B12-deficient lambs, there
was a sharp decrease of vitamin B12 concentrations in blood and liver
before signs like anorexia, loss of body weight, and a decrease in hemoglobin concentration were observed. Sheep were more sensitive to cobalt
deficiency than cattle, and Mburu et al. (1993) suggested that sheep are
likewise less resistant to low dietary cobalt than goats, based on blood
parameters.
As cobalt is required for biosynthesis of vitamin B12, lack of cobalt
may cause deficiency of the vitamin in adult ruminants. Cobalt-deficient soils occur in large areas of many countries, therefore grazing ruminants may be particularly affected by the deficiency. With the exception of phosphorus and copper, cobalt deficiency is the most extensivemineral limitation to grazing livestock in tropical countries (McDowell,1997).
Cobalt deficiency signs are not specific, and it is often difficult todistinguish between an animal with cobalt deficiency and malnutrition
due to low intake of energy and protein and an animal that is diseasedor parasitized. Acute clinical signs of cobalt deficiency include lack ofappetite, rough hair coat, thickening of the skin, anemia (normocytic
and normochromic), wasting away (Fig. 13.2), and eventually death, if
the animals are not moved to “healthy” pastures or if cobalt supplements are not made available.
Cobalt deficiency has been reported to reduce lamb survival and increase susceptibility to parasitic infection in cattle and sheep (Ferguson
et al., 1988; Suttle and Jones, 1989). Cobalt deficiency was associatedwith photosensitization of lambs, characterized by a swollen head (Hesselink and Vellema, 1990). The condition responded to two injections ofvitamin B12 3 weeks apart. Cobalt-deficient ewes produced fewer lambs
and had more stillbirths and neonatal mortality than cobalt-sufficient541

CHAPTER THIRTEEN

A

B
Fig. 13.2 (A) A cobalt-deficient heifer that had access to an iron-copper salt
supplement. Note severe emaciation, which resulted from failure to synthesize
B12. Her blood contained 6.6 g of hemoglobin per 100 ml on February 25,
1937. (B) The same heifer fully recovered with an iron-copper-cobalt salt supplement while on the same pasture. (Courtesy of R.B. Becker, University of
Florida.)controls (Fisher and MacPherson, 1991). Lambs from deficient ewes
were also slower to start suckling. Auricular myocardial necrosis was reported in cobalt-deficient sheep (Mohammed and Lamand, 1986).
At necropsy, the body of a severely affected cobalt-deficient animalpresents a picture of extreme emaciation, often with a total absence ofbody fat. The liver is fatty, the spleen hemosiderized, and in some animals there is hypoplasia of the erythrogenic tissue in the bone marrow(Filmer, 1933). The anemia in lambs is normocytic and normochromic,
but the mild anemia is not responsible for the main signs of cobalt deficiency. Inappetence and marasmus invariably precede any considerable
542

V I T A M I N B1 2

degree of anemia. Clinical signs of cobalt deficiency are identical tothose of simple starvation, and may indicate that the signs observed withadequate cobalt may be simply due to the inappetence caused by the lackof cobalt. The first discernible response to cobalt feeding or parenteralvitamin B12 is rapid improvement in appetite and body weight.
Two other conditions attributed to cobalt deficiency are ovine whiteliver disease and Phalaris staggers (Graham, 1991; McDowell, 1992;
Kennedy et al., 1994b). Ovine white liver disease is characterized by hepatic lipidosis and emaciation. At necropsy, affected lambs had pale,swollen, and friable fatty livers and showed accumulation of lipofuscin(Kennedy et al., 1994b). Alteration in choline synthesis presumably
leads to impaired lipid mobilization, but white liver disease may be complicated by other factors. Similarly, the persistent neural effects of
Phalaris spp., inducing Phalaris staggers, have been shown to be preventable with oral cobalt supplementation, but not with B12.
Prior to the recognition of cobalt deficiency in livestock in many
parts of the world, cattle could be maintained on deficient pastures only
if they periodically were relocated to so-called healthy ground. Cobalt
deficiency can be prevented by moving animals for a few months every
year to a healthy region, preferably during the rainy season. An example of the necessity of periodically moving animals was illustrated in adisease condition known as “togue” in Espirito Santo, Brazil (Tokarnia
et al., 1971). The disease was observed when animals stayed for longer
than 60 to 180 days on certain pastures. Sick animals isolated themselves from the rest of the herd; were apathetic; showed loss of appetite,rough hair coat, and dry feces; and lost body condition. If the animals
were not moved from the pasture, they died, but if they were taken to a
pasture where the disease did not occur, the animals recovered quickly.
Cobalt subclinical deficiencies or borderline states are extremelycommon and are characterized by low production rates unaccompanied by clinical manifestations or visible signs (McDowell, 1985). Subclinical deficiencies often go unnoticed, thereby resulting in great economic losses to the livestock industry. No estimate can be made of theeffect of cobalt subdeficiency on animal performance in general, but inmany areas of the world, it is one of the major causes of poor production.

Swine
The general signs of vitamin B12 deficiency in pigs are comparable
to those observed in other species, principally a loss of appetite, variable
feed intake, and a dramatic growth decline (Fig. 13.3). In addition,
543

CHAPTER THIRTEEN

A

B

Fig. 13.3 (A) Vitamin B12-deficient pig. Note rough hair coat and dermatitis.
(B) Control pig. (Courtesy of the late D.V. Catron and Iowa State University.)

sometimes there is rough skin and hair coat, vomiting and diarrhea,
voice failure, and slight anemia (Catron et al., 1952). Microcytic tonormocytic anemia is typical; however, observations on anemia are notunanimous and are sometimes contradictory. Nervous disorders occur inthe pig, including increased excitability, and unsteady gait (i.e., hind leg
incoordination). The thymus and spleen become atrophied, while liver
and tongue are frequently enlarged as a result of proliferation of granulomatous tissue.
In the reproducing animal, litter size and pig survival are reduced.
Abortions, small litters and birth weights, some deformities, and inability to rear young occur in breeding sows. Later estrus, fewer corpora544

V I T A M I N B1 2

lutea, and fewer embryos are produced in B12-deficient animals. During
reproduction and lactation, vitamin B12 supplementation was shown toincrease birth weight and survival of young pigs (Vestal et al., 1950).
Frederick and Brisson (1961) found that sows deficient in vitamin B12
had fewer pigs, and these had a lower viability than the pigs born fromsows that were supplemented with vitamin B12. Successive litters fromdeficient sows became progressively weaker. In a study of sows supplemented with 80 to 100 μg of vitamin B12 daily during pregnancy, there
were improved piglet and litter weights and a decreased percentage ofstillbirths (Reinisch and Gebhardt, 1987). Under some conditions, reproductive performance of sows were improved by inclusion of higher
than recommended levels of dietary vitamin B12 (Cunha, 1977). The response was evidenced by an increase in litter size and birth weight ofpigs.

Poultry
In growing chicks, turkey poults, and quail, vitamin B12 deficiency
reduces body weight gain, feed intake, and feed conversion. Vitamin B12
deficiency in growing chicks and turkeys may result in a nervous disorder and defective feathering. It has also been related to leg weakness andperosis; however, this appears to be a secondary effect. Perosis may occur in vitamin B12-deficient chicks or poults when the diet lacks choline,
methionine, or betaine as sources of methyl groups. Addition of B12 may
prevent perosis under these conditions because of its effect on synthesisof methyl groups. Additional clinical signs in B12 deficiency include anemia, gizzard erosion, and fattiness of heart, liver, and kidneys. Poor
feathering and mortality are the most obvious signs of vitamin B12 deficiency, and gizzard erosions may also appear (NRC, 1994).
In hens, body weight and egg production are maintained despite deficiency, but B12 has an important influence on egg size (Scott et al.,1982). However, Squires and Naber (1992) reported that both egg production and hen weight increased with vitamin B12 supplementation, as
well as increased hatchability and egg weight.
Hatchability of incubated eggs may be severely reduced if thebreeder diet contains inadequate vitamin B12. Changes that manifest
themselves in vitamin B12-deficient chick embryos (Olcese et al., 1950)
may be summarized as (1) general hemorrhagic condition; (2) fatty liverin varying degrees; (3) heart often enlarged and irregular in shape; (4)kidneys pale or yellow, sometimes hemorrhagic, incidence of perosis; (5)myoatrophy of the leg; (6) fewer myelinated fibers in the spinal cord;and (7) high incidence of embryonic malpositions. Hypertrophy of the545

CHAPTER THIRTEEN

thyroid gland has also been repeatedly observed (Ferguson and Couch,1954).
The most obvious change in B12-deficient embryos is myoatrophy ofthe leg, a condition characterized by atrophy of thigh muscles (Olcese etal., 1950). Two to 5 months may be needed to deplete hens of vitamin
B12 stores to such an extent that progeny will hatch with low vitamin B12
reserves. The rate of depletion is most rapid when hens are fed high-protein diets (Scott et al., 1982). Chicks that hatch without adequate carryover of vitamin B12 from the dam have a high rate of mortality. Vitamin
B12-deficient embryos die at about day 17.

HorsesVitamin B12 deficiency has not been described in the horse. It has
been concluded that for mature horses, supplemental vitamin B12 is notnecessary, as the vitamin can be synthesized in the large intestine, where
it is absorbed (NRC, 1989b). Davies (1971) subsequently demonstrated
increased vitamin B12 concentration in gut contents from stomach torectum. Values (ng/ml) were as follows: stomach, 4; small intestine, 5;cecum, 30; ventral colon, 84; dorsal colon, 197; and small colon, 301.
These increasing concentrations were presumably the consequence ofsynthesis by intestinal anaerobes. Of 97 gut microbe isolates, 47% were
found to produce vitamin B12 in vitro.
Even with low dietary cobalt intake, B12 deficiency would not be expected because dietary cobalt levels greatly exceed the theoretical needfor vitamin B12 synthesis. In addition, horses have remained in good
health while grazing pastures so low in cobalt that cattle and sheep confined to them have died (Filmer, 1933).

Other Animal Species
BATS
Neurological impairment associated with experimentally induced
vitamin B12 deficiency was produced in bats (Vieira-Makings et al.,1990). Methionine, valine, and isoleucine have been shown to delay onset of the deficiency.

DOGS AND CATS
Vitamin B12-deficient kittens exhibited poor growth, lethargy, emaciation and a high level of methylmalonic acid excretion (Keesling andMorris, 1975; Vaden et al., 1992). Morris (1977) reported that kittens
given a vitamin B12-deficient diet at first grew normally for 3 to 4546

V I T A M I N B1 2

months, after which growth ceased. Subsequently, body weight was lostat an accelerating rate until supplementation was initiated with parenteral vitamin B12, which restored weight gain.
Vitamin B12 deficiency has been described in the dog (NRC, 1985a).
Earlier reports noted reduced growth (Arnrich et al., 1952) and impaired reproduction (Campbell and Phillipps, 1952) with B12 deficiency.
An inherited intestinal vitamin B12 malabsorption disorder was reported
in dogs as a result of a defective brush-border expression of intrinsic factor–vitamin B12 receptor (Fyfe et al., 1989, 1991). Vitamin B12-deficientdogs developed chronic inappetence, lethargy, chronic nonregenerative
anemia, and failure to thrive at 12 weeks of age. Vitamin B12 deficiency
has been identified in 50% of German shepherds with degenerative
myelopathy; however, the clinical signs were not responsive to vitamin
B12 treatment (Toenniessen and Morin, 1995).

FISH
Salmon and trout fed low dietary B12 showed high variability innumbers of fragmented erythrocytes and in hemoglobin values, with apropensity for microcytic, hypochromic anemia (NRC, 1993). Channel
catfish and grass shrimp fed a vitamin B12-deficient diet exhibited reduced growth rates (NRC, 1993; Shiau and Lung, 1993a). John and
Mahajan (1979) observed reduced growth and lower hematocrit in rohu
fed a vitamin B12-deficient diet. Japanese eel were found to require vitamin B12 for normal appetite and growth (Arai et al., 1972). Intestinal
microflora synthesis of vitamin B12 was shown to be sufficient to meetthe requirements for carp, tilapia, rainbow trout, ayu, and goldfish(NRC, 1993; Shiau and Lung, 1993b). Of 746 bacteria strains tested incarp, 8% of the bacteria consumed vitamin B12 and the remaining 92%
produced it (Sugita et al., 1994). More than 50% of the obligate anaerobes, especially the genus Clostridium, produced large amounts of vitamin B12.

LABORATORY ANIMALSIn practically all laboratory animals, growth is retarded in vitamin
B12 deficiency (NRC, 1995), and there are changes in relative weights ofcertain organs. In B12-deficient rats, there are kidney lesions and mucoid
structures in the urinary bladders, with reduced muscle mass and fibrotic degeneration of heart. In rats, deficiency in the diet of the mothercan result in hydrocephalus, eye defects, and bone defects in the newborn, and mothers may eat the offspring. In mice deficient in B12, there
is death of young, retarded growth, and renal atrophy. Hamsters receiv547

CHAPTER THIRTEEN

ing low dietary vitamin B12 exhibited reduced feed efficiency, but there
was no apparent effect on body weight or blood parameters (Scheid etal., 1950). Under practical conditions, vitamin B12 deficiency would not
be expected in most laboratory animals because of intestinal synthesis
and the significant amounts of the vitamin obtained by coprophagy.

MINK
Mink kits deficient in vitamin B12 exhibit anorexia, loss of bodyweight, and severe fatty degeneration of liver (NRC, 1982a). Such deficiency is unlikely in these animals, which are commonly fed diets rich inanimal protein. Oral vitamin B12 supplementation, but not injections,
had some effect in preventing iron-deficiency anemia, indicating an influence on intestinal iron absorption (Pezacka et al., 1992).

NONHUMAN PRIMATES
Megaloblastic anemia reminiscent of pernicious anemia in humans
has not been reported in vitamin B12-deficient monkeys. However, abnormalities of the central and peripheral nervous systems in several
species of monkeys have been attributed to B12 deficiency as a result ofvegetarian diets (NRC, 1978). Agamanolis et al. (1976) described demyelination and axon loss in the papillomacular bundle and optic nerve,
degeneration within the optic chiasm, and loss of nuclei in all six layersof the lateral geniculate body of rhesus monkeys fed a B12-deficient dietfor 4 years.

RABBITSHigh urinary and fecal excretion rates of B12 have been reported in
rabbits receiving diets practically devoid of the vitamin (NRC, 1977). As
a result of coprophagy, rabbits should not be deficient in vitamin B12, assuming adequate cobalt is available.

HumansIn humans, pernicious anemia, a fatal megaloblastic anemia withneurological involvement, is the result of vitamin B12 deficiency. This
megaloblastic anemia is not found in animals. This finding by itself does
not elucidate whether the condition is due to deficiency of B12 or folacin.
Either vitamin B12 or folacin supplementation will cure the megaloblastic anemia; however, folacin is ineffective in preventing degenerativechanges in the nervous system. In macrocytic anemia, erythrocytes arelarger than normal and show great variation in size and normal hemoglobin saturation. The bone marrow shows a megaloblastic pattern of548

V I T A M I N B1 2

red cell maturation as opposed to the usual normoblast pattern.
In addition to megaloblastic anemia, the most prominent signs andsymptoms of vitamin B12 deficiency are weakness, tiredness, lightheadedness, pale and smooth tongue with inflammation (Fig. 13.4), dyspnea,
splenomegaly, leukopenia, thrombocytopenia, achlorhydria, paresthesia, neurological changes, loss of appetite, loss of weight, and low serum
cobalamin levels. The condition results in stiffness of limbs, progressive
paralysis, mental disorders, diarrhea, and finally death.
The basic pathological lesion is demyelination of nervous system tissue, seen mainly in the posterolateral columns of the spinal cord but occurring also as foci of demyelination in the cerebral white matter (Metz,
1993). Neurological B12 deficiency results in axon degeneration of
nerves in the spinal cord. The deficiency produces patchy, diffuse, andprogressive demyelination. The clinical picture of the diffuse, uneven demyelination is one of an insidiously progressive neuropathy, often beginning in the peripheral nerves and progressing centrally to involve theposterior and lateral columns of the spinal cord (Herbert, 1990). In addition to subacute combined degeneration of the cord, hypovitaminosis
B12 may give rise to a severe psychosis with extensive mental deterioration.
The function of B12 and folacin in DNA synthesis accounts for someof the pathological findings when deficiency of either of these vitamins
occurs. The most rapidly proliferating tissues of the body, such as bonemarrow for blood cell production, have the greatest requirement for
DNA synthesis and thus are principally affected by megaloblastic anemia when severe deficiency of either B12 or folacin occurs. Disturbed division and nuclear maturation of proliferating epithelial cells can be observed in buccal mucosal scrapings and intestinal biopsies ofB12-deficient patients. The proliferating epithelial cells, as is true forbone marrow, have a high B12 and folacin requirement for DNA synthesis.
At the biochemical level the lesion responsible for neuropathy wasoften attributed to defective functioning of the adenosylcobalamin-dependent folacin-independent methylmalonyl CoA mutase reaction,rather than of the cobalamin-dependent folacin-dependent methionine
synthetase reaction (Metz, 1992). However, it is postulated that in vitamin B12 deficiency, impaired methionine synthetase activity leads in turnto deficiencies of methionine and S-adenosylmethionine (SAM). Low
levels of cerebrospinal SAM are related to abnormal myelination in inherited disorders that affect vitamin B12 and folacin metabolism (Metz,
1993). Because SAM is a key intermediary in methylation reactions, de549

CHAPTER THIRTEEN

A

Fig. 13.4 Pernicious
anemia. In addition tomegaloblastic anemia,
an additional sign withvitamin B12 deficiency isa pale, smooth tongue
with inflammation (A),which is found in onethird to one-half of pernicious anemia patients.
The normal tongue (B)has papilla. (Courtesyof R.R. Streiff, Veterans
Administration, University of Florida.)

Bficiency of SAM could be expected to impair methylation reactions inmyelin; thus, methyl group deficiency could result in demyelination andclinical neuropathy (Metz, 1993). A report of studies on infants andchildren with inborn errors of the methyl-transfer pathway provided
further evidence of an association between reduced cerebrospinal fluid
SAM levels and abnormal myelination in the nervous system (Surtees etal., 1991).
Vitamin B12 deficiency in humans is influenced by one or more of
the following considerations:
1. Dietary intake—Inadequate vitamin B12 intake is occasionally seen in
geriatric patients and in vegetarians. A completely vegetarian diet (one devoid of meat, eggs, and dairy products) can produce B12 deficiency if consumed for several years. In some areas of India, religion and poverty resultin a basic diet of polished rice, cereals, some vegetables, and fruit, with aninadequate intake of milk and eggs (Rothenberg and Cotter, 1978). Inci550

V I T A M I N B1 2

dence of serum vitamin B12 deficiencies were great in Thailand in womenpracticing vegetarianism for 6 to 10 years (Tungtrongchitr et al., 1993). In
Mexico, 62% of breast milk samples from women consuming high-vegetable diets were deficient in vitamin B12 (Black et al., 1994). Vitamin B12
deficiency was very prevalent in lactating Guatemalan women and their infants at 3 months postpartum (Casterline et al., 1997). There have also beenreports of cobalamin deficiency in infants breast-fed by strictly vegetarian
mothers (Higginbottom et al., 1978). Breast-fed babies of mothers who arevitamin B12 deficient may develop megaloblastic anemia and neurologic
symptoms in the first few months of life. Graham et al. (1992) described vitamin B12 deficiency in six infants 8 to 15 months old. Consistent clinicalsigns were irritability, anorexia, and failure to thrive, associated withmarked developmental regression and poor brain growth. After follow-up
for several years, two of four children tested had mild to borderline intellectual retardation.
2. Failure of absorption or transport—Pernicious anemia is most commonly acquired because of a failure to secrete intrinsic factor. This may be
due to deranged stomach activity or may follow total gastrectomy. Total
gastrectomy in humans always produces cobalamin deficiency, since it completely removes the site and source of intrinsic factor secretion.
Patients with certain small intestine defects have failure of B12 absorption. Impaired absorption of cobalamin is a regular manifestation of tropical sprue. Patients with lesions such as blind loops or small bowel diverticula have demonstrated that an inappropriate bacterial overgrowth instagnant areas may introduce into the intestinal stream sufficient organisms
to absorb all or adsorb much of the dietary cobalamin. Fish tapeworm (Diphyllobothrium latum) infestation is a well-recognized cause of impaired B12
absorption, as the worm sequesters the vitamin as it progresses through thesmall intestine. Other conditions that result in reduced B12 absorption ortransport include excessive intake of alcohol or certain drugs, chronic pancreatitis, diseased B12 ileal receptors, and abnormalities related to B12 transport proteins.
3. Storage—Vitamin B12 stores in the human body exceed the daily requirement by about 1,000-fold (Ellenbogen and Cooper, 1991), which helps
explain why clinical B12 deficiency due to dietary insufficiency is not morecommon. It may take as long as 5 to 7 years after cessation of intrinsic-factor secretion before any outward signs of pernicious anemia are evident.
Moreover, the small intestine contains microflora that can synthesize significant amounts of cobalamin.
4. Heredity—Pernicious anemia, inherited as an autosomal dominant
trait, chiefly affects persons past middle age and results in lack of intrinsic551

CHAPTER THIRTEEN

factor production. The incidence of pernicious anemia in the general population is one to two per 1,000 and about 25 per 1,000 among relatives of
pernicious anemia patients (Ellenbogen and Cooper, 1991).

Neural tube defects (NTDs) are associated with inadequate dietaryfolacin (see Chapter 12, Deficiency, Humans). Evidence indicates that vitamin B12 is also an independent risk factor for NTDs, and the enzyme
methionine synthase is involved either directly or indirectly, as this is the
only enzyme in metabolism that requires both folate and vitamin B12 tofunction. Further evidence that methionine synthase is defective inmothers who have NTD-affected pregnancies has been reported (Scott etal., 1995). Methionine synthase is required to demethylate 5-CH3THF
with the methyl group used to remethylate homocysteine to produce methionine and SAM, the latter being required for methylation reactions.
Low levels of the active form of folacin could impair synthesis of DNA
and SAM. Alternatively, impaired activity of methionine synthase might
in itself decrease DNA or SAM biosynthesis. Any of the above events
could lead to an NTD (Scott et al., 1995).
Carriers of the defective enzyme methionine synthase may also be atrisk of other diseases in later life. Homocysteine, the substrate of the enzyme may be inadequately metabolized, leading to hyperhomocysteinemia, a condition that is known to predispose to cardiovascular and cerebrovascular diseases (Clark et al., 1991; Herzlich et al., 1996). Such a
defect would be exacerbated by low plasma folate and/or plasma vitamin B12 levels (see Chapter 12, Deficiency, Humans).

Assessment of Status
A positive diagnosis of vitamin B12 deficiency is usually made byfinding subnormal serum and tissue B12 concentrations. Low serum levels are associated with low body content of the vitamin. Microbiological assay methods or radioisotopic dilution techniques may be used forthis purpose. The normal range for vitamin B12 in human serum (orplasma) is 200 to 900 pg/ml. Values between 150 and 200 pg/ml
strongly suggest B12 deficiency (Rothenberg and Cotter, 1978). A serum
concentration of vitamin B12 below 110 pmol/L is associated with megaloblastic bone marrow, incipient anemia, and myelin damage, and below
150 pmol/L, there are early bone marrow changes (Bender, 1992).
Analysis of serum levels of both B12 and folate can sometimes differentiate which nutrient is deficient for a given megaloblastic anemia(see Chapter 12). However, serum vitamin B12 concentration has beenshown to have limitations in specificity and sensitivity in diagnosing vi-

552

V I T A M I N B1 2

tamin B12 deficiency and predicting response to therapy in subjects withclinical deficiency syndromes. Frequently, both serum folacin and vitamin B12 concentrations are found to be low or low normal in a patientwith megaloblastic anemia, making distinction between the two syndromes difficult (Lindenbaum et al., 1994; Savage et al., 1994). In addition to problems with diagnosis, the serum B12 concentration cannot be
used to monitor response to therapy because it increases in all subjects
who receive parenteral B12 regardless of whether they were deficient ornot.
Contrary to serum vitamin B12, elevations of methylmalonic acidand total homocysteine are very sensitive and specific in diagnosing vitamin B12 deficiency and can be used to help differentiate vitamin B12
deficiency from folacin deficiency (Stabler et al., 1996). Elevated total
homocysteine concentrations that may have been attributed to folacin
deficiency in elderly subjects may in many instances be the result of vitamin B12 deficiency even though serum vitamin B12 concentrations arewithin normal limits. Vitamin B12 deficiency is confirmed by determining serum levels of homocysteine and methylmalonic acid. Urinary
methylmalonic acid is a sensitive, convenient, and noninvasive indicator for vitamin B12 versus folacin deficiency and has been used as ascreening procedure in elderly populations (Norman and Morrison,
1993).
Different methods involving various radioactive isotopes of cobalt
are available for determining the absorption of an oral dose of radioactive B12 in humans. The Schilling urinary excretion test is one of the simplest and most generally used. The basis of this test is giving large dosesof unlabeled B12 to saturate blood binding protein so any absorbed radioactive B12 will be excreted by the kidney. Other procedures include
direct determination of fecal radioactivity after a test dose of B12-labeledcobalt (i.e., 58Co or 56Co) or measurement of hepatic uptake of B12.
Biochemical detection of B12 deficiency in both humans and animals
includes increased urinary excretion of formiminoglutamic acid (FIGLU)
following loading with histidine and a greater percentage of serum folate as 5-methyl-THF rather than THF. Since vitamin B12 participates indemethylation of methylfolate, patients with pernicious anemia show an
accumulation of the methylfolate and an increased excretion of FIGLU.
For deamination of histidine, THF is required rather than methyl-THF.
In laying poultry, vitamin B12 concentrations in egg yolk can be used
as a status indicator. Squires and Naber (1992) reported that egg yolk
vitamin B12 concentrations respond rapidly to dietary changes in the553

CHAPTER THIRTEEN

level of this vitamin and are indicative of the vitamin B12 status of thehen. At four times the dietary requirement, efficiency of transfer to eggs
remained nearly constant for vitamin B12 (Naber and Squires, 1993).
Hence, vitamin B12 fortification of eggs is easily accomplished.
In ruminants, the best indicators of cobalt deficiency are low levelsof cobalt and B12 in tissues, loss of appetite, elevated blood pyruvate,
and elevated urinary methylmalonic acid. A vitamin B12 enzyme(methylmalonyl-CoA mutase) is required for conversion of propionate
to succinate and, with the vitamin deficiency, methylmalonic acid is excreted. Methylmalonic acid concentrations would be higher in all species
with B12 deficiency but particularly high in ruminants because of thelarge quantities of propionic acid metabolized. Other indicators of low
vitamin B12 status have been increased plasma homocysteine (Kennedy
et al., 1994b) and low liver methionine synthase activity in sheep(Kennedy et al., 1992a).
In sheep consuming low dietary vitamin B12, plasma vitamin B12
concentration decreased below the lower limit of normal after 6 weeks,
and plasma methylmalonic acid concentration increased above the upper limit after 10 weeks (Kennedy et al., 1994a). However, Fisher and
MacPherson (1990) suggested that serum methylmalonic acid concentrations are less variable and provide a more accurate diagnosis of cobalt
deficiency than serum vitamin B12. Normal concentrations of serum
methylmalonic acid were tentatively suggested as being less than 2
μmol/L; subclinically cobalt deficient, 2 to 4 μmol/L; and cobalt deficient greater than 4 μmol/L (Paterson and MacPherson, 1990). Graham(1991) suggested that in ruminants, plasma B12 is marginally deficient at
380 to 760 pmol/L and deficient when under 380 pmol/L. In goats,
serum vitamin B12 concentrations below 200 pg/ml were indicative of
cobalt deficiency (Mburu et al., 1994).
The levels of cobalt in the livers of sheep and cattle are sufficiently
responsive to changes in cobalt intake to have value in the detection ofcobalt deficiency, with liver vitamin B12 an even more reliable criterion.
Values of 0.10 μg or less of vitamin B12 per gram wet weight are “clearly
diagnostic of cobalt deficiency disease” (Underwood, 1979). Livercobalt concentrations in the range of 0.05 to 0.07 ppm (dry basis) or below are critical levels indicating deficiency (McDowell, 1985). While
herbage and tissue analyses are helpful in diagnosing the deficiency, thedefinite proof is the prompt improvement in feed intake following injection of vitamin B12.

554

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SUPPLEMENTATION
Vitamin B12 is produced by fermentation and is available commercially as cyanocobalamin for addition to feed. Vitamin B12 is only
slightly sensitive to heat, oxygen, moisture, and light (Gadient, 1986;
Coelho, 1994). Verbeeck (1975) reported vitamin B12 to have good stability in premixes with or without minerals regardless of mineral source.
Scott (1966) indicated that there is apparently little effect of pelleting on
vitamin B12 content of feed. Vitamin leaching is a problem when feed isadministered in water for fish. Pannevis and Earle (1994) reported a
90% loss of vitamin B12 after only 30 seconds through water leaching of
the feed.
Results of a large number of animal experiments are about equallydivided between those reporting a positive response to dietarycyanocobalamin and those reporting little or no response. Variable responses may be due to several factors: initial body stores, environmental sources of the vitamin (such as molds, soil, and animal excreta), microbial synthesis in the intestinal tract, and adequacy or deficiency ofother nutrients that influence B12 requirements.
Vitamin B12 is normally added to diets of all classes of swine andpoultry. Swine and poultry raised in confinement, in management systems where there is less access to feces for coprophagy, should have agreater dietary requirement for the vitamin. Although dietary supplements would be recommended, injections of vitamin B12 are often given
to animals with a poor health appearance. Animals coming into a feedlot are sometimes given B12 injections, along with other vitamins, as insurance against not quickly adapting to new feeding regimes. This use of
B12 may be warranted under certain conditions in which stress, disease,or parasites lower feed intake, impair ruminal function, and/or reduce
intestinal absorption.
Intramuscular administration of vitamin B12 at the rate of 100 μg
each week or of 150 μg every second week were shown to produce a
rapid remission of all signs of deficiency in lambs and are just as effective as cobalt administered orally at the rate of 7 mg per week (Andrews
and Anderson, 1954). For rapid correction of cobalt deficiency in cattle,
intramuscular administration of vitamin B12 at 500 to 3,000 μg per headis recommended, which may be repeated weekly (Graham, 1991). Intramuscular administration of vitamin B12 to cobalt-deficient animals produced overnight improvement in appetite, whereas oral dosing with555

CHAPTER THIRTEEN

cobalt required 7 to 10 days to produce the same effect (MacPherson,1982). Although parenteral vitamin B12 injections prevent cobalt deficiency in ruminants, it is more convenient and economical to supplement the diet with cobalt, allowing the microorganisms to synthesize thevitamin for subsequent absorption by the host.
Since many forages and some concentrate feeds do not supply 0.10
ppm cobalt, an adequate dietary allowance, cobalt supplementation isneeded. Ruminants that consume concentrates are more likely to receive
adequate dietary cobalt than grazing livestock; however, growth responses to supplemental cobalt were demonstrated in steers fed finishing diets based on barley grain (Raun et al., 1968), sorghum grain, andsilage (Morris and Gartner, 1967).
Available cobalt must be present in the animal diet for ruminal andintestinal synthesis to occur. Cobalt deficiency in ruminants can be cured
or prevented through treatment of soils or pastures with cobalt-containing fertilizers or by direct oral administration of cobalt to the animals
through free-choice mineral supplements (McDowell, 1985, 1992). In
deficient areas, where the pastures require regular fertilizer applications,adequate cobalt intakes can usually be ensured by including cobalt salts
or oxide ores. Often, use of fertilizer cobalt is impractical and uneconomical for extensive range conditions in developing countries (McDowell et al., 1984).
Cobalt deficiency in grazing ruminants can best be prevented by direct oral administration of cobalt through free-choice mineral supplements. During critical periods cobalt supplementation must be continuous. Cobalt supplementation in either the first or second half ofpregnancy only did not fully alleviate adverse effects on ewe reproductive performance and lamb viability (Fisher and MacPherson, 1991).
Cobalt-deficient livestock respond quickly to cobalt treatment and recover appetite, vigor, and weight; this serves as an easy practical test to
determine whether cobalt deficiency exists. Large, frequent injections ofvitamin B12 can effectively prevent or cure cobalt deficiency but aremuch more expensive. Oral dosing or drenching with dilute cobalt solutions are satisfactory if the doses are regular and frequent. Dosing sheep
twice weekly with 2 mg of cobalt or once weekly with 7 mg of cobalt,
or dosing cattle with 5 to 10 times those amounts, depending on theirsize and age, is fully adequate for severely deficient regions (Underwood,1981).
An additional method of providing cobalt, developed in Australia,
is the use of an orally administered, heavy pellet (bullet), made of cobalt
oxide plus finely divided iron, that remains in the reticulorumen for an556

V I T A M I N B1 2

extended period. Pellets may be lost through regurgitation or become ineffective because of formation of a surface coating of calcium phosphate. The addition of a steel grinder, which provides an abrasive action,
reduces the surface coating and extends the usefulness of the pellet. A
multiple slow-release trace element (including cobalt) rumen bolus isavailable (Judson et al., 1988; Ritchie et al., 1991).
The need for continual cobalt supplementation is difficult to assess,as the incidence of cobalt deficiency can vary greatly from year to year,
from undetectable mild deficiency to an acute stage. Lee (1963) illustrated this variation in a 14-year experiment with sheep in southern Australia. Half the ewes, replacements, and progeny were dosed with cobalt
and remained healthy. The undosed half had the following performancefor the 14 years: in 2 years, lambs were unthrifty, but there were no
deaths; in 3 years, growth rate of the lambs was slightly retarded; in 4
years, 30 to 100% of the lamb crop was lost; in 5 years, the performance of the remaining stock was as good as that of dosed animals.
The most important cobalt supplements produced for feed use arecobalt carbonate, a fine powder containing over 46% cobalt; cobalt sulfate crystal, a fine granular crystal containing 21% cobalt; and cobalt
sulfate monohydrate, a powder containing 33% cobalt. Carbonate,chloride, sulfate, and oxide forms of cobalt were proposed as satisfactory dietary sources of the mineral (Ammerman and Miller, 1972).
However, observations by Ammerman et al. (1982) suggested that theoxide form may be considerably less available than either the carbonate
or sulfate form.
For human therapy, during the stage of relapse of pernicious anemia, intramuscular doses of vitamin B12 at the rate of 15 to 30 μg/day
should be given (Marks, 1975). A single injection of 100 μg or more willproduce complete remission in any patient whose vitamin B12 deficiency
is not complicated by unrelated systemic disease or other factors. Whenpernicious anemia is due to inadequate absorption, 1 μg of the vitaminby injection daily is adequate therapy. Remission is sustained for life bymonthly injections of 100 μg of vitamin B12 (Herbert, 1990). In patientstreated successfully with intramuscular vitamin B12, the first response
occurs within about 2 days of the start of treatment and consists of an
intense feeling of well-being and an increase in appetite.
Single, large oral doses (1,000 μg) of vitamin B12 without intrinsic
factor have proven effective in treatment of pernicious anemia. Absorption of a small amount of B12 from massive doses is independent of theaction of intrinsic factor and is believed to occur by a “mass-action” effect, resulting in diffusion of some of the vitamin. An oral dose of at557

CHAPTER THIRTEEN

least 150 μg/day is deemed necessary to maintain the pernicious anemia
patient. Single weekly oral doses of 1,000 μg satisfactorily maintainsome pernicious anemia patients (Ellenbogen and Cooper, 1991). For
vegetarian patients who have a normal secretion of intrinsic factor, 1.0
to 1.5 μg/day of vitamin B12 orally is sufficient to prevent pernicious
anemia.
Vitamin B12 supplementation is particularly important for individuals consuming all-vegetable diets. Supplementation of B12 should beconsidered by vegetarians, particularly for lactating mothers to provide
vitamin B12 to their offspring. If malabsorption and chronic diarrhea are
combined with low dietary intake of vitamin B12, as is the case for many
children in the Third World, depletion of vitamin B12 stores may result(Paerregaard et al., 1990). Premature infants require additional folacin
and vitamin B12 to reduce the severity of the anemia of prematurity(Worthington-White et al., 1994). When high amounts of supplemental
folacin are administered (e.g., to prevent NTDs), the risk of undiagnosed
vitamin B12 deficiency could be reduced by simultaneous supplementation with generous doses (1 mg/day) of vitamin B12. A 20% prevalenceof abnormal vitamin B12 metabolism was found in 64 patients with human immunodeficiency virus (HIV) who were referred for neurologicalevaluation (Kieburtz et al., 1991). Vitamin B12 deficiency may be a frequent and treatable cause of neurological dysfunction in patients with
HIV infection.
Supplementation with large doses of vitamin B12 has falsely been advocated for various disorders in humans. The red color of the vitamin,along with its almost total lack of known toxicity, makes it an almost
ideal placebo. It is, unfortunately, used in large quantities to defraud unsuspecting buyers.

TOXICITY
Addition of vitamin B12 to food in amounts far in excess of need orabsorbability appears to be without hazard. Dietary levels of at least several thousand times the requirement are safe (NRC, 1987; Ellenbogen
and Cooper, 1991). However, the maximum tolerable amount of dietarycobalt for ruminants is estimated at 5 ppm. Cobalt toxicosis in cattle ischaracterized by mild polycythemia; excessive urination, defecation, andsalivation; shortness of breath; and increased hemoglobin, red cellcount, and packed cell volume. Although most researchers consider themargin between safe and toxic doses wide enough to make toxicity under natural conditions unlikely, several reports have appeared in the lit558

V I T A M I N B1 2

erature concerning cobalt toxicity. These reports almost invariably have
involved management mistakes in formulating mineral mixtures.

■ REFERENCES
Agamanolis, D.P., Chester, E.M., Victor, M., Kark, J.A., Hines, J.S., and Harris,
J.W. (1976). Neurology 26, 905.
Ammerman, C.B., and Miller, S.M. (1972). J. Anim. Sci. 35, 681.
Ammerman, C.B., Henry, P.R., and Loggins, P.R. (1982). J. Anim. Sci. 55(Suppl.
1), 403.
Andrews, E.D. (1956). N. Z. J. Agric. 92, 239.
Andrews, E.D., and Anderson, J.P. (1954). N. J. Z. Sci. Tech. A35, 483.
Anonymous (1984). Vitamin B12. Roche Tech. Bull. Hoffmann-LaRoche, Nutley, New Jersey.
Arai, S., Nose, T., and Hashimoto, Y. (1972). Bull. Freshwater Res. Lab. Tokyo22, 69.
Arnrich, L., Lewis, E.M., and Morgan, A.F. (1952). Proc. Soc. Exp. Biol. Med.
80, 401.
Barnes, R.H., and Fiala, G. (1958). J. Nutr. 65, 103.
Becker, R.B., Henderson, J.R., and Leighty, R.B. (1965). Fla. Univ. Agric. Exp.
Stn. Tech. Bull., 699.
Behrns, K.E., Smith, C.O., and Sarr, M.G. (1994). Dig. Dis. Sci. 39(2), 315.
Bender, D.A. (1992). Nutritional Biochemistry of the Vitamins, p. 294. Cambridge University Press, Cambridge, England.
Black, A.K., Allen, L.H., Pelto, G.H., DeMata, M.P., and Chávez, A. (1994). J.
Nutr. 124, 1179.
Campbell, J.E., and Phillips, P.H. (1952). J. Nutr. 47, 621.
Carmel, R. (1994). Dig. Dis. Sci. 12, 2516.
Casterline, J.E., Allen, L.H., and Ruel, M.T. (1997). J. Nutr. 127, 1966.
Catron, D.V., Richardson, D., Underkofler, L.A., Maddock, H.M., and Friedland, W.C. (1952). J. Nutr. 47, 461.
Clark, R., Daly, L., Robinson, K.H., Naughtey, E., Cahalane, S., Fowler, B., andGraham, I. (1991). N. Engl. J. Med. 324, 1149.
Coelho, M.B. (1994). In BASF Technical Symposium, p. 60. Clarion, Arkansas.
Corrêa, R. (1957). Arquivos do Instituto Biologicos 24, 199.
Cunha, R.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Dagnelie, P.C., VanStaveren, W.A., and VandenBerg, H. (1991). Am. J. Clin.
Nutr. 53, 695.
Davies, M.E. (1971). Br. Vet. J. 127, 34.
Dostalova, J. (1994). Vyzivay 49, 43.
Ellenbogen, L., and Cooper, B.A. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 491. Marcel Dekker, New York.
Ellenbogen, L., and Highley, D.R. (1970). Fed. Proc. Fed. Am. Soc. Exp. Biol.
29, 633(Abstr.).
Farquharson, J., and Adams, J.F. (1976). Br. J. Nutr. 36, 127.
Ferguson, E.G.W., Mitchell, G.B., and MacPherson, A. (1988). Vet. Rec. 124,
20.
559

CHAPTER THIRTEEN

Ferguson, R.M., and Couch, J.R. (1954). J. Nutr. 54, 361.
Fie, M., Zee, J.A., and Amiot, J. (1994). Lait 74, 461.
Filmer, J.F. (1933). Aust. Vet. J. 9, 163.
Fisher, G.E.J., and MacPherson, A. (1990). Br. Vet. J. 146, 120.
Fisher, G.E.J., and MacPherson, A. (1991). Res. Vet. Sci. 50, 319.
Flavin, M., and Ochoa, S. (1957). J. Biol. Chem. 229, 965.
Folkers, K. (1982) In B12 (D. Dolphin, ed.), p. 1. Wiley, New York.
Frederick, G.L., and Brisson, G.J. (1961). Can. J. Anim. Sci. 41, 212.
Friesecke, H. (1980). Vitamin B12. Hoffmann-La Roche, Basel, Switzerland.
Fyfe, J.C., Jezyk, P.F, Giger, U., and Patterson, D.F. (1989). J. Am. Anim. Hosp.
Assoc. 25, 533.
Fyfe, J.C., Ramanujam, K.S., Ramaswamy, K., Patterson, D.F., and Seetharm, B.
(1991). J. Biol. Chem. 266, 4489.
Gadient, M. (1986). Proc. Nutr. Conf. Feed Mfr., p. 73. University of Maryland,College Park.
Graham, S.M., Arvela, O.M., and Wise, G.A. (1992). J. Pediatr. 121, 710.
Graham, T.W. (1991). Vet. Clin. North Am. Food Anim. Proc. 7, 153.
Henderickx, H.K., Teague, H.S., Redman, D.R., and Grifo, A.P. (1964). J. Anim.
Sci. 23, 1036.
Herbert, V. (1990). Nutrition Reviews: Present Knowledge in Nutrition (R.E.
Olson, H.P. Broquist, C.O. Chichester, W.J. Darby, A.C. Kolbye, and R.M.
Stalvey, eds.), 5th Ed., p. 170. Nutrition Foundation, Washington D.C.
Herbert, V., and Zalusky, R. (1962). J. Clin. Invest. 41, 1263.
Herzlich, B.C., Lichstein, E., Schulhoff, N., Weinstock, M. Pagala, M., Ravindran, K., Namba, T., Nieto, F.J., Stabler, S.P., Allen, R.H., and Malinow,
M.R. (1996). J. Nutr. 126, 1249S.
Hesselink, J.W., and Vellema, P. (1990). Tijdschrift-voor-Diergeeneskunde 115,
789.
Higginbottom, M., Sweetman, L., and Nyhan, W. (1978). N. Engl. J. Med. 299,
317.
Hussein, H.S., Fahey, G.C., Wolf, B.W., and Berger, L.L. (1994). J. Dairy Sci. 77,
3432.
John, M.J., and Mahajan, C.L. (1979). J. Fish Biol. 14, 127.
Jorgensen, B.B., Pedersen, N.T., and Worning, H. (1991). Aliment. Pharmacol.
Ther. 5, 207.
Judson, G.J., Brown, T.H., Kempe, B.R., and Turnbull, R.K. (1988). Aust. J.
Exp. Agr. 28. 299.
Keen, C.L., and Graham, T.W. (1989). In Clinical Biochemistry of Domestic Animals (J.J. Kaneko, ed.), 4th Ed., p. 753. Academic Press, San Diego California.
Keesling, P.T., and Morris, J.G. (1975). J. Anim. Sci. 41, 317(Abstr.).
Kennedy, D.G., Blanchflower, W.J., Scott, J.M., Weir, D.G., Molloy, A.M.,
Kennedy, S., and Young, P.G. (1992a). J. Nutr. 122, 1384.
Kennedy, D.G., Blanchflower, W.J., Young, P.G., and Davidson, W.B. (1992b).
Biol. Trace Elem. Res. 35, 153.
Kennedy, D.G., Kennedy, S., Blanchflower, W.J., Scott, J.M., Weir, D.G., Molloy,
A.M., and Young, P.B. (1994a). Brit. J. Nutr. 71, 67.
Kennedy, D.G., Young, P.B., Blanchflower, W.J., Scott, J.M., Weir, D.G., Molloy,
A.M., and Kennedy, S. (1994b). Int. J. Vit. Nutr. Res. 64, 270.
560

V I T A M I N B1 2

Kieburtz, K.D., Giang, D.W., Schiffer, R.B., and Vakil, N. (1991). Arch. Neurol.
48, 312.
Lachance, A.P. (1998). Nutr. Rev. 56, 34S.
Lassiter, C.A., Ward, G.M., Huffman, C.F., Duncan, C.W., and Webster, H.D.
(1953). J. Dairy Sci. 36, 997.
Latteur, J.P. (1962). Cobalt Deficiencies and Subdeficiencies in Ruminants. Centre d’Information du Cobalt, Brussels, Belgium.
Lee, H.J. (1963). In Animal Health, Production and Pasture (A.H. Worden,
K.O. Sellers, and D.E. Tribe, eds.), p. 662. Longmans, New York.
Lewis, N.M., Kies, C., and Fox, H.M. (1986). Nutr. Rep. Int. 34, 495.
Lindenbaum, J., Rosenberg, I., Wilson, P., Stabler, S.P., and Allen, R.H. (1994).
Am. J. Clin. Nutr. 60, 2.
Loosli, J.K. (1991). In Handbook of Animal Science (P.A. Putnam, ed.), p. 25.
Academic Press, San Diego, California.
Lopez-Guisa, J.M., and Satter, L.D. (1992). J. Dairy Sci., 75, 247.
MacPherson, A. (1982). Roche Vitamin Symposium: Recent Research on the Vitamin Requirements of Ruminants, p. 1. Hoffmann-La Roche, Basel,
Switzerland.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
Marston, J.R. (1970). Br. J. Nutr. 24, 615.
Mburu, J.N., Kamau, J.M.Z., and Badamana, M.S. (1993). Int. J. Vit. Nutr. Res.
63, 135.
Mburu, J.N., Kamau, J.M.Z., Badamana, M.S., and Mbugua, P.N. (1994). Bull.
Anim. Health Prod. Africa 42, 141.
McDowell, L.R. (1985). In Nutrition of Grazing Ruminants in Warm Climates(L.R. McDowell, ed.), p. 329. Academic Press, Orlando, Florida.
McDowell, L.R. (1992). Minerals in Animal and Human Nutrition Academic
Press, San Diego, California.
McDowell, L.R., Conrad, J.H., and Ellis, G.L. (1984). In Symposium on Herbivore Nutrition in Subtropics and Tropics-Problems and Prospects (F.M.C.
Gilchrist and R.I. Mackie, eds.) p. 67. The Science Press, Pretoria, South
Africa.
McDowell, L.R. (1997). Minerals for Grazing Ruminants in Tropical Regions,
3rd Ed. University of Florida Press, Gainesville, FL.
Metz, J. (1992). Annu. Rev. Nutr. 12, 59.
Metz, J. (1993). Nutr. Rev. 51, 12.
Mohammed, R. (1983). Sub-clinical Cobalt Deficiency in Sheep. Ph.D. dissertation, Univ. of Clermont, Clermont, France.
Mohammed, R., and Lamand, M. (1986). Ann. Res. Vet. 17, 447.
Morris, J.G. (1977). In Kal Kan Symposium for Treatment of Dog and Cat Diseases, p. 15. Ohio State University, Wooster.
Morris, J.G., and Gartner, R.J.W. (1967). Agric. Sci. 68, 1.
Muhammad, K., Briggs, D., and Jones, G. (1993). Food Chem. 48, 431.
Naber, E.C., and Squires, M.W. (1993). Poult. Sci. 72, 1046.
Norman, E.J., and Morrison, J.A. (1993). Am J. Med. 94, 589.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
561

CHAPTER THIRTEEN(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed. National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Olcese, O., Couch, J.R., Quisenberry, J.H., and Pearson, P.B. (1950). J. Nutr. 41,
423.
Paerregaard, A., Hjelt, K., and Krasilnikoff, P.A. (1990). J. Pediatr. Gastroenterol. Nutr. 11, 351.
Pannevis, M.C., and Earle, K.E. (1994). J. Nutr. 124(Suppl.), 2633S.
Paterson, J.E., and MacPherson, A. (1990). Vet. Rec. 126, 329.
Pezacka, E.H., Jacobsen, D.W., Luce, K., and Green, R. (1992). Biochem. Biophys. Res. Comm. 184, 832.
Polak, D.M., Elliot, J.M., and Haluska, M. (1979). J. Dairy Sci. 62, 697.
Ramoz, J.J., Saez, T., Bueso, J.P., Sanz, M.C., and Fernandez, A. (1994). Vet.
Res. 25, 405.
Raun, N.S., Stables, G.L., Pope, L.S., Harper, O.F., Waller, G.R., Renbarger, R.,
and Tillman, A.D. (1968). J. Anim. Sci. 27, 1695.
RDA. (1989). Recommended Dietary Allowances, 10th Ed. National Academyof Sciences-National Research Council, Washington, D.C.
Reinisch, F., and Gebhardt, G. (1987). In Symposium Vitamin and Ergotropika
und Podiumsdiskussion zur Verzehrsregulation Reinhardsbrunn, E. Germany.
Ritchie, N.S., Lawson, D.C., and Parkins, J.J. (1991). In Tracae Elem. Man
Anim. Proc. Int. Symp. (TEMA-7), p. 15. Dubrovnik, Yugoslavia.
Rothenberg, S.P., and Cotter, R. (1978). In Handbook Series in Nutrition andFood, Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 3, p. 69.
CRC Press, Boca Raton, Florida.
Sato, K., Wang, X., and Mizoguchi, K. (1997). J. Dairy Sci. 80, 2701.
Savage, D.G., and Lindenbaum, J. (1995). In Folate in Health and Disease (L.B.
Bailey, ed.), p. 237. Marcel Dekker, New York.
Savage, D.G., Lindenbaum, J., Stabler, S.P., and Allen, R.H. (1994). Am. J. Med.
96, 239.
Scheid, H.E., McBride, B.H., and Schweigert, B.S. (1950). Proc. Soc. Exp. Biol.
Med. 75, 236.
562

V I T A M I N B1 2

Scott, J.M., Weir, D.G., and Kirke, P.N. (1995). In Folate in Health and Disease(L.B. Baily, ed.), p. 329. Marcel Dekker, New York.
Scott, M.L. (1966). Proc. Cornell Nutr. Conf., p. 35. Cornell University, Ithaca,
New York.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Sebrell, W.H. Jr., and Harris, R.S. (1968). The Vitamins, p. 119. Academic Press,
New York.
Sewell, R.F., Cunha, T.J., Shawver, C.B., Ney, W.A., and Wallace, H.D. (1952).
Am. J. Vet. Res. 13, 186.
Shiau, S.Y., and Lung, C.Q. (1993a). Comp. Biochem. Phys. A. 105, 147.
Shiau, S.Y., and Lung, C.Q. (1993b). Aquaculture 117, 157.
Shinton, N.K. (1972). Br. Med. J. 1, 556.
Simpson, K.W., Morton, D.B., and Batt, R.M. (1989). Am. J. Vet. Res. 50, 1233.
Squires, M.W., and Naber, E.C. (1992). Poult. Sci. 71, 2075.
Stabler, S.P., Lindenbaum, J., and Allen, R.H. (1996). J. Nutr. 126, 1266s.
Sugita, H., Kuruma, A., Hirato, C., Ohkoshi, T., Okada, R., and Deguchi, Y.
(1994). Aquaculture 119, 425.
Sugita, H., Miyajima, C., and Deguchi, Y. (1991). Aquaculture 92, 267.
Surtees, R., Leonard, J., and Austin, S. (1991). Lancet 338, 1550.
Suttle, N.F., and Jones, D.G. (1989). J. Nutr. 119, 1055.
Sutton, A.L., and Elliot, J.M. (1972). J. Nutr. 102, 1341.
Toenniessen, J.G., and Morin, D.E. (1995). Compendium Cont. Ed. Pract. Vet.
17, 217.
Tokarnia, C.H., Guimaraes, J.A., Canella, C.F.C., and Döbereiner, J. (1971).
Pesqui. Agropecu. Bras. 6, 61.
Toskes, P.P., Deren, J.J., Fruiterman, J., and Conrad, M.E. (1973). Gastroenterology 65, 199.
Tungtrongchitr, R., Pongpaew, P., Prayurahong, B., Changbumrung, S., Vudhivai, N., Migasena, P., and Schelp, F.P. (1993). Int. J. Vit. Nutr. Res. 63,
201.
Underwood, E.J. (1977). Trace Elements in Human and Animal Nutrition, Academic Press, New York.
Underwood, E.J. (1979). In Proc. Nutr. Conf., p. 203. University of Florida,
Gainesville, Florida.
Underwood, E.J. (1981). The Mineral Nutrition of Livestock, Commonwealth
Agricultural Bureaux, London, England.
Vaden, S.L., Wood, P.A., Ledley, F.D., Cornwell, P.E., Miller, R.T., and Page, R.
(1992). J. Am. Vet. Med. Assoc. 200, 1101.
Verbeeck, J. (1975). Feedstuffs 47(36), 4, 45.
Vestal, C.M., Beeson, W.M., Andrews, P.N., Hutchings, L.M., and Doyle, L.P.
(1950). Purdue Agric. Exp. Stn. Mimeo No. 50. Purdue University,
Lafayette, Indiana.
Vieira-Makings, E., Van-der Westhuyzen, J., and Metz, J. (1990). Int. J. Vitam.
Nutr. Res. 60, 41.
Wagle, S.R., Mehta, R., and Johnson, B.C. (1958). J. Biol. Chem. 230, 137.
West, R. (1948). Science 107, 398.
Worthington-White, D.A., Behnke, M., and Gross, S. (1994). Am J. Clin. Nutr.
563

chapter fourteen

CHOLINEINTRODUCTION
Choline is considered essential to the animal organism and is utilized both as a building unit and as an essential component in regulationof certain metabolic processes. Choline is tentatively classified as one of
the B-complex vitamins even though it does not entirely satisfy the strict
definition of a vitamin. Choline, unlike B vitamins, can be synthesized in
the liver, is required in the body in greater amounts, and apparently
functions as a structural constituent rather than as a coenzyme. Also, existence of choline in essential body constituents was recognized long before the first vitamin was discovered. Regardless of classification,choline is an essential nutrient for all animals and a required dietary
supplement for some species (e.g., poultry, swine, and fish). Choline is
likewise an essential nutrient for humans, and studies have investigatedthe benefits of choline treatment for certain diseases, such as cancer andAlzheimer’s disease.

HISTORY
Choline was isolated by Streker from the bile of pigs in 1849 and by
Von Balb and Hirschbrunn from an alkaloid of white mustard seed(Sinapis alba) in 1852 (Griffith and Nyc, 1971). Streker isolated thecompound from lecithin, to which he gave the name “choline.” Thechemical structure of choline was established in 1867 by Bayer (Scott etal., 1982). Choline’s acceptance as a biologically essential compound resulted from studies in 1929 in which acetylcholine was isolated from thespleen of a horse (Dale and Dudley, 1929).
Choline’s role in nutrition was not known until the 1930s, following
565

14

CHAPTER FOURTEEN

the discovery of insulin in 1922 by Banting and Best. It was observed thatfatty degeneration of the liver associated with insulin deprivation in dogs
could be corrected by feeding either raw pancreas or lecithin. In 1932
choline was discovered to be the active component of pure lecithin previously shown to prevent fatty livers (“lipotropic effect”) in rats (Best etal., 1934). Betaine, considered to be only a methyl donor, was found tohave a similar lipotropic effect in rats and dogs. Various studies in the late
1930s and early 1940s determined that the lipotropic activity of choline
related to transfer of methyl groups and interrelationships and sparing effects of choline on dietary methionine, folacin, and vitamin B12.
In 1940, Jukes showed that choline is required for normal growth
and the prevention of perosis, a leg disorder in turkeys; he found thatthe amount required to prevent perosis was greater than that required
for normal growth. Choline was found to prevent a “spraddled-leg”
condition in swine in the 1950s and 1960s. More recently, choline has
been found to be effective for various human conditions involving mobilization of liver lipids, and studies have investigated the role of choline(and other methyl donors) on carcinogenesis and the neurotransmitterrelated syndromes (e.g., senile dementia and Alzheimer’s disease).

CHEMICAL STRUCTURE AND PROPERTIES
Choline is a β-hydroxyethyltrimethylammonium hydroxide and is
depicted in Fig. 14.1. The prominent feature of choline’s chemical structure is its triplet of methyl groups, which enables it to serve as a methyl
donor. Pure choline is a colorless, viscid, strongly alkaline liquid that isnotably hygroscopic. Choline is soluble in water, formaldehyde, and alcohol and has no definite melting or boiling point. The chloride salt of
this compound, choline chloride, is produced by chemical synthesis foruse in the feed industry, although other forms exist. Choline chloride exists as deliquescent white crystals that are very soluble in water and alcohols. Aqueous solutions are almost pH neutral.
Choline is ubiquitously distributed in all plant and animal cells,
mostly in the form of the phospholipids phosphatidylcholine (lecithin,see Fig. 14.1), lysophosphatidylcholine, choline plasmalogens, andsphingomyelin-essential components of all membranes (Zeisel, 1990).
Lecithin is the predominant phospholipid (> 50%) in most mammalian
membranes. In the lung, disaturated lecithin is the major active component of surfactant (Brown 1964), lack of which results in a respiratory
distress syndrome in premature infants. Choline is a precursor for thebiosynthesis of the neurotransmitter acetylcholine (Fig. 14.1). Glyc566

CHOLINE

Fig. 14.1 Structural formulas for freecholine, acetylcholine, and lecithin. R′
refers to any fatty acid.

erophosphocholine and phosphocholine are storage forms for choline
within the cytosol and principal forms found in milk (Rohlfs et al.,1993).

ANALYTICAL PROCEDURES
Determination of choline is complicated by its various forms in biological materials. Free choline can be extracted with water or alcohol,
but more precise procedures must be used for extraction of total choline.
Precautions have to be taken for the preparation of biological tissues,
which contain enzymes that rapidly hydrolyze phosphate esters ofcholine in nervous tissues when animals die (Chan, 1991). Much of theevidence for presence of free choline in biological materials is unreliableowing to delay in the preparation of extracts, with resulting release ofcholine by autolysis (Griffith and Nyc, 1971). For example, dog liver567

CHAPTER FOURTEEN

contained 0 to 43 mg choline per kilogram if extracted immediately after death of the animal and 136 to 164 mg choline per kilogram if extracts were made 5 hours postmortem.
The classic and most widely used procedure for quantitative determination of choline is the colorimetric reineckate method (AOAC,
1984). This procedure involves the precipitation of choline as a reinecke
salt for a colorimetric reaction. Other methods of analysis include enzymatic, fluorometric, gas chromatographic, photometric, and polarographic techniques. Analyses by high-performance liquid chromatograhy and gas chromatography-mass spectrometry (Pomfret et al.,
1989; Zeisel and da Costa, 1990) have permitted the measurement ofcholine-containing compounds in tissues. Enzymatic radioisotopic assay
and gas chromatography provide high sensitivity and specificity andhave been widely used in recent years (Chan, 1991). Microbiological assay for choline generally employs Neurospora crassa, while biologicalassay methods make use of conversion of choline to acetylcholine in isolated tissues, as well as growth response of chicks. Growth response may
be an unreliable measure of choline concentration as it is influenced byother constituents (e.g., methionine) in choline-limiting diets (Pesti et al.,1981).

METABOLISM
Choline is present in the diet mainly in the form of lecithin, with lessthan 10% present as either the free base or sphingomyelin. Choline is released from lecithin and sphingomyelin by digestive enzymes of the gastrointestinal tract, although 50% of ingested lecithin enters the thoracic
duct intact (Chan, 1991).
Choline is released from lecithin by hydrolysis in the intestinal lumen. Both pancreatic secretions and intestinal mucosal cells contain enzymes capable of hydrolyzing lecithin in the diet. Phospholipase A2(which cleaves the β-fatty acid moiety) is found in pancreatic juice andin the intestinal brush border. Within the gut mucosal cell, phospholipase A1 cleaves the α-fatty acid, and phospholipase B cleaves both fatty
acids. Quantitatively, digestion by pancreatic lipase is the most important process (Zeisel, 1990). The net result is that most ingested lecithin
is absorbed as lysophosphatidylcholine (deacylated in the β position).
Within the cells of the gut wall, lysophosphatidylcholine can be deacylated to form glycerophosphocholine, or it can be acylated to reconstitute lecithin.
Choline is absorbed from the jejunum and ileum mainly by an en568

CHOLINE

ergy- and sodium-dependent carrier mechanism. In the guinea pig,choline was taken up by ileal cells about three times faster than by jejunal cells (Hegazy and Schwenk, 1984). Preferential location of the transport system in ileal cells supports earlier findings in the rat and hamster(Sanford and Smyth, 1971). These results are expected since choline isreleased from lecithin in the proximal and middle small intestine.
Only one-third of ingested choline appears to be absorbed intact.
The remaining two-thirds is metabolized by intestinal microorganisms
to trimethylamine, which is excreted in the urine between 6 and 12
hours after consumption (De La Huerga and Popper, 1952). In contrast,
when an equivalent amount of choline is consumed as lecithin, less urinary trimethylamine is excreted, with most of the metabolite appearing
in urine 12 to 24 hours after consumption. Dietary choline is the principal factor governing excretion, with presence or absence of othersources of protein and of fat having relatively little effect. Absorbed
choline is transported into the lymphatic circulation primarily in theform of lecithin bound to chylomicra, it is transported to the tissues predominantly as phospholipids associated with the plasma lipoproteins.
Choline is present in all tissues as an essential component of phospholipids in membranes of all types. All tissues accumulate choline, but
uptake by liver, kidney, mammary gland, placenta, and brain are of especial importance. Uptake of choline by mammary cells enables this tissue to concentrate choline almost 70-fold versus maternal blood (Chaoet al., 1988). Most species can synthesize choline, as lecithin, by the sequential methylation of phosphatidylethanolamine (Fig. 14.2) by phosphatidylethanolamine-N-methyltransferase. This activity is actually due
to enzymes that use S-adenosylmethionine (SAM) as the methyl donor.
Choline synthesis activity is low in male rats and absent from chicks until about the thirteenth week of age. The activity is greatest in liver, but
is also found in many other tissues.
Choline is released in free form in the tissues by the actions of phospholipase C, which cleaves the circulating form (lecithin) to yield adiglyceride and phosphorylcholine. Free choline can be oxidized by the
mitochondrial enzyme choline dehydrogenase to yield betaine aldehyde,
which is then converted by the cytosolic enzyme betaine aldehyde dehydrogenase to betaine. Betaine is the actual source of methyl groups. Onlya small fraction of choline is acetylated, but that amount provides theimportant neurotransmitter acetylcholine. This step involves the reaction of choline with acetyl CoA and is catalyzed by choline acetyltransferase localized in cholinergic nerve terminals, as well as in certain othernonnervous tissues (e.g., placenta).
569

CHAPTER FOURTEEN

Fig. 14.2 Metabolic pathway for the synthesis of choline and related compounds. Folacin and vitamin B12 are required for synthesis of methyl groups
and metabolism of the one-carbon unit. (Adapted from Scott et al., 1982.)

In ruminants, dietary choline is rapidly and extensively degraded in
the rumen from studies with both sheep (Neill et al., 1979) and cattle(Atkins et al., 1988; Sharma and Erdman, 1988). In sheep fed forage diets, choline is rapidly degraded to trimethylamine with only 10 to 15%of the dietary choline escaping rumen degradation; however, the degreeof degradation in different diets is unknown (Neill et al., 1979). Estimates of rumen degradation have ranged from 85 to 99%. In vivo studies with dairy cows, in which choline intake was increased up to 303
g/day over controls, there was an increase of only 1.3 g/day in choline
flow to the duodenum (Sharma and Erdman, 1988).
Work with sheep (Neill et al., 1979) and goats (Emmanual and Kennelly, 1984) suggests that ruminants must metabolize and utilize choline
in a different manner than monogastric animals. Choline absorption
must be very limited in all ruminants because of (1) almost complete
degradation of dietary choline in the rumen, (2) only limited supplies
from any rumen protozoa that might escape rumen degradation, and (3)570CHOLINE

rumen bacteria, which are devoid of choline.

FUNCTIONS
Choline functions in four broad categories in the animal body:
1. Choline is a metabolic essential for building and maintaining cellstructure. As a phospholipid it is a structural part of lecithin (phosphatidylcholine), certain plasmalogens, and the sphingomyelins. Choline is incorporated into phospholipid by being converted to phosphoryl choline, then tocytidine diphosphate choline, and finally reacting with phosphatidic acid tolecithin. The phospholipids and total fatty acids present are affected by nutritional state and the type of fatty acids present in the diet. Lecithin is a partof animal cell membranes and lipid transport moieties in cell plasma membranes. Phospholipids exist in the cell membrane bilayers, and it is thoughtthat one of the primary roles of phospholipids is to regulate cell membrane
porosity by changing the ionic characteristics of the membrane. Lecithin isalso an essential component of very-low-density lipoprotein, the bloodtransport molecule for hepatic triacylglycerol (Lombardi et al., 1966). In theprevention of perosis, choline is required as a constituent of the phospholipids needed for normal maturation of the cartilage matrix of the bone. Various metabolic functions and synthesis of choline are depicted in Fig. 14.2.
2. Choline plays an essential role in fat metabolism in the liver. The first
discovered function of dietary choline dealt with prevention of fatty liver indepancreatized dogs, and later in rats, chicks, and other species. Owing to
the basic function of choline in membrane structure, the lack of choline ismanifested in a variety of phospholipid-related functions, such as fatty liverand lesions of the kidney and impairment of lipoprotein metabolism.
Choline prevents abnormal accumulation of fat (fatty livers) by promoting
its transport as lecithin or by increasing the utilization of fatty acids in theliver itself. Choline is thus referred to as a lipotropic factor because of its
function of acting on fat metabolism by hastening removal or decreasing
deposition of fat in liver.
3. Choline is essential for the formation of acetylcholine, the agent released at the termination of the parasympathetic nerves (Wauben and Wainwright, 1999). It makes possible the transmission of nerve impulses frompresynaptic to postsynaptic fibers of the sympathetic and parasympathetic
nervous systems. For example, acetylcholine released by the stimulated vagus nerve causes a slowing of heartbeat, and oviduct contraction resultsfrom the action of acetylcholine. Acetylcholine is the most common neuro571

CHAPTER FOURTEEN

transmitter in the nervous system. Apparently, brain tissue lacks the ability
to synthesize sufficient choline (Ansell and Spanner, 1971) for neural function. However, apparently circulating choline is the major source of choline
for acetylcholine synthesis.
4. A fourth function of choline is as a source of labile methyl groups forformation of methionine from homocystine and of creatine from
guanidoacetic acid. However, practical significance of the choline-homocystine interrelationship is of no real importance to feeding animals since natural proteins contain very little of the metabolic intermediate homocystine(Ruiz et al., 1983). Methyl groups also function in the synthesis of purine
and pyrimidine, which are used in the production of DNA. Methionine isconverted to S-adenosylmethionine in a reaction catalyzed by methionine
adenosyl transferase. S-adenosylmethionine is the active methylating agent
for many enzymatic methylations. A disturbance in folacin or methionine
metabolism results in changes in choline metabolism and visa versa (Zeisel,1990). The involvement of folacin, vitamin B12, and methionine in methylgroup metabolism, and of methionine in de novo choline synthesis, may allow these substances to substitute in part for choline. Severe folacin deficiency has been shown to cause secondary liver choline deficiency in rats
(Kim et al., 1994).

The demand for choline as a methyl donor is probably the major
factor that determines how rapidly a diet deficient in choline will induce
pathology. The pathways of choline and 1-carbon metabolism intersect
at the formation of methionine from homocysteine. Methionine is regenerated from homocysteine in a reaction catalyzed by betaine-homocysteine methyltransferase, in which betaine, a metabolite of choline,
serves as the methyl donor (Finkelstein et al., 1982). Large increases inchick hepatic betaine-homocysteine methyltransferase can be produced
under methionine-deficient conditions, especially in the presence of excess choline or betaine (Emmert et al., 1996). To be a source of methylgroups, choline must be converted to betaine, which has been shown toperform methylation functions as well as choline in some cases. However, betaine fails to prevent fatty livers and hemorrhagic kidneys. A diagram of the choline cycle, including related compounds and donations
to the body’s methyl pool, is shown in Fig. 14.3.
Since choline contains biologically active methyl groups, methionine
can partly be spared by choline and homocysteine. Research with lactating dairy cattle suggests that a high proportion of dietary methionine
is used for choline synthesis (Erdman and Sharma, 1991). Discoveries insignal transduction are beginning to reveal other important roles for572

CHOLINE

Fig. 14.3 Choline cycle showing metabolic generation and use of active methylgroups. (Adapted from Umbrett, 1960.)

choline phospholipids. Signal transduction is a new area of cell biology
research and is the process by which hormones and other substancestransmit messages from the cell’s surface to its interior, and by which a
broad spectrum of cell activities—such as growth, gene expression, ion
transport, and energy utilization—are regulated. Evidence shows thatcholine phospholipids play a central role in this process. Disruptions in
phospholipid metabolism can interfere with this process and may underlie certain disease states, such as cancer and Alzheimer’s disease(Canty and Zeisel, 1994).
The choline phospholipid metabolite plasmalogen has been studied
in relation to myocardial infarction. High levels of plasmalogen arefound in the sarcolemma, the cell membrane of heart muscle. An adverse
573

CHAPTER FOURTEEN

effect of acute myocardial ischemia may be the result of plasmalogen
breakdown during an ischemic attack (Canty and Zeisel, 1994).

REQUIREMENTS
Choline, unlike most vitamins, can be synthesized by most species,
although in many cases not in sufficient amounts or rapidly enough tosatisfy all the animal’s needs. For example, when intake of precursors oraccessory factors—such as methionine, vitamin B12, or folacin—is insufficient, poultry, dogs, cats, monkeys, pigs, rabbits, and certain fish (e.g.,bass, catfish, carp and trout) have been shown to require exogenous
choline. Also, young animals show a higher need for choline than adults.
Dietary factors such as methionine, betaine, myo-inositol, folacin, andvitamin B12, or the combination of different levels and composition offat, carbohydrate, and protein in the diet, as well as the age, sex, caloric
intake, and growth rate of animals, all have influence on the lipotropic
action of choline and thereby on the requirement of this nutrient (Mookerjea, 1971). Dietary betaine can spare choline, since choline functions
as a methyl donor by forming betaine. In relation to protein level, alarger choline effect on litter size and piglet and litter weight was observed for gilts fed a 12% protein diet than for those fed a 16% protein
diet (Maxwell et al., 1987).
Studies have shown that vitamin B12 and folacin reduce requirement
for choline in chicks and rats (Welch and Couch, 1955). Folacin and vitamin B12 are required for the synthesis of methyl groups and metabolism of the one-carbon unit. Biosynthesis of labile methyl from a formate
carbon requires folacin, while B12 plays a role in regulated transfer of themethyl group from tetrahydrofolic acid. Therefore, marked increases incholine requirement have been observed under conditions of folacin
and/or vitamin B12 deficiency.
The two principal methyl donors functioning in animal metabolism
are choline and methionine, which contain biologically labile methylgroups that can be transferred within the body. This phenomenon iscalled transmethylation. Methionine furnishes a methyl group that can
combine with ethanolamine to form choline and, in reverse, methylgroups from choline (du Vigneaud et al., 1939). Therefore, dietary adequacy for both methionine and choline directly affect requirements ofeach other. Other than exogenous sources of methyl groups from choline
and methionine, methyl group formation from de novo synthesis of formate carbons is reduced with folate and/or vitamin B12 deficiencies.
574

CHOLINE

Most animals can synthesize sufficient choline for their needs provided enough methyl groups are supplied. As an example, methionine in
the pig can completely replace that portion of the choline needed fortransmethylation. Thus, at methionine levels in excess of the physiological requirement, 4.3 mg of methionine—or 3.69 mg with hydroxyl(OH) group considered in molecular weight of choline—provides thesame methylating capacity as 1 mg of choline (NRC, 1998). Kroening
and Pond (1967) fed 5-kg pigs a low-protein diet (12%) supplemented
with three levels of dl-methionine—0, 0.11, and 0.22%. The addition of
1,646 mg of choline per kilogram of diet improved the weight gain andfeed conversion of pigs fed the control and the 0.11% but not the 0.22%
methionine-supplemented diet. Young poultry, on the contrary, are unable to benefit from methionine or betaine as a dietary replacement forcholine unless methylaminoethanol or dimethylaminoethanol is in thediet, as they appear unable to methylate aminoethanol when fed a purified diet (Jukes, 1947). Later studies showed that the chick can synthesize microsomal methylaminoethanol and choline from S-andenosylmethionine, but, unlike the pig, at an insufficient rate to cover its needs(Norvell and Nesheim, 1969).
Calculations from lactating dairy goats on rates of methyl grouptransfer revealed that only 6% of methionine methyl groups were derived from choline, while 28% of choline methyl groups were derived
from methionine (Emmanual and Kennelly, 1984). This suggests thatconsiderable dietary methionine is used for choline synthesis.
The metabolic needs for choline can be supplied in two ways: by dietary choline or by choline synthesis in the body, which makes use of labile methyl groups. For selected species, body synthesis sometimes cannot take place fast enough to meet choline needs for rapid growth, andthus clinical signs of deficiency result. Since choline functions in the prevention of fatty livers, hemorrhagic kidneys, and perosis, it does not act
as a true vitamin since it is incorporated into phospholipids (via cytidine
diphosphocholine). Therefore, unlike a typical B vitamin, the choline
molecule becomes an integral part of the structural component of liver,kidney, or cartilage cells (Scott et al., 1982).
In general, males are more sensitive to choline deficiency than females (Wilson, 1978). Growth hormone seemed to increase the choline
requirement in rats independent of its ability to promote growth and increase food intake (Hall and Bieri, 1953). Cortisone and hydrocortisone
have been reported to decrease severity of renal necrosis, and hydrocortisone reduced the amount of hepatic lipid in choline-deficient rats (Olson, 1959).
575

CHAPTER FOURTEEN

Excess dietary protein increases the young chick’s choline requirement. Ketola and Nesheim (1974) observed that over three times asmuch choline was needed for maximum growth of chicks fed a diet containing 64% protein versus 13%. Diets high in fat aggravate choline deficiency and thus increase requirement. Fatty liver is generally enhanced
by fats containing a high proportion of long-chain saturated fatty acids(Hartroft, 1955). Choline deficiency develops to a greater degree in rapidly growing animals with deficiency lesions more severe in these animals.
Use of antibiotics or prevention of coprophagy in rats decreases dietary choline requirement (Barnes and Kwong, 1967). Likewise, dietaryrequirement of choline is reduced in germ-free rats. Intestinal flora convert choline to trimethylamine, a lipotropically inactive compound thatis excreted in urine. Suppression of intestinal flora by antibiotics would
result in choline being more available than as inactive trimethylamine.
Recommended dietary allowances for choline or amounts typicallyfed for a number of animal species are listed in Table 14.1. Requirements for choline have generally been determined through the use of purified diets, and recommendations often do not take into accountbioavailability from feedstuffs, individual animal variation, or effects ofother dietary factors. Typical choline requirements for monogastric animals range from 1,000 to 2,000 mg/kg of feed. In contrast to monogastric animals, no requirements for choline have been established for ruminant animals except for milk-fed calves, in which 260 mg of choline
per liter of synthetic milk prevented choline deficiency signs. The NRC(1989a) suggests that milk replacers for calves should contain 0.26%
choline.
Choline is an essential nutrient for humans on the basis of deficiency
signs for those receiving experimental diets low in choline (Zeisel et al.,1991) and for individuals receiving total parenteral nutrition devoid ofcholine (Buchman et al., 1992). Choline is likely not essential when diets
contain excess methionine and folacin. Human choline requirements
have not been established (RDA, 1989). Based on a caloric intake of
2,000 kcal/day, one might expect the typical human diet to range incholine content from 410 to 1375 mg/kg dry matter. When compared torequirements for other animal species and, in particular, mature animals,
it seems unlikely that choline deficiency would occur (DuCoa L.P., 1994).

NATURAL SOURCES
All naturally occurring fats contain some choline, and thus it is sup576

CHOLINE■ Table 14.1

Choline Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Beef cattle
Dairy cattle

Adult
Calf milk replacer
Adult
Leghorn, 0–6 weeks
Leghorn, 6–12 weeks
Broilers, 0–3 weeks
Broilers, 6–8 weeks
Starting and growing
Breeding
Growing, 0–4 weeks
Growing, 8–16 weeks
Adult
Growing, 3–5 kg
Growing, 5–120 kg
Adult
Adult
Adult
Adult
Growing
All classes
All classes
CatfishLake trout
Common carp
All classes

Tissue synthesis
2,600 mg/kg liter
Tissue synthesis
1,300 mg/kg
900 mg/kg
1,300 mg/kg
750 mg/kg
2,000 mg/kg
1,500 mg/kg
1,600 mg/kg
950 mg/kg
Tissue synthesis
0.6 g/kg
0.3–0.5 g/kg
1.00–1.25 g/kg
Tissue synthesis
Tissue synthesis
2,400 mg/kg
1,250 mg/kg
0.12% choline chloride
1,000 mg/kg
400 mg/kg
1,000 mg/kg
1,500 mg/kg
Unknown

NRC (1984a)
NRC (1989a)
NRC (1989a)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
NRC (1985b)
NRC (1998)
NRC (1998)
NRC (1998)
NRC (1989b)
NRC (1981b)
NRC (1986)
NRC (1985a)
NRC (1977)
NRC (1978a)
NRC (1993)
NRC (1993)
NRC (1993)
RDA (1989)

ChickenJapanese quail
Turkey
Sheep
Swine
Horse
Goat
Cat
Dog
Rabbit
Rat
Fish
Human

aExpressed as per unit of animal feed on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a,b) (except for dairy calf).

plied by all feeds that contain fat. Choline is almost always found in theform of lecithin or sphingomyelin in most foods. Egg yolk (1.7%), glandular meats (0.6%), and brain and fish (0.2%) are the richest animal
sources, and germ of cereals (0.1%), legumes (0.2 to 0.35%), andoilseed meals are the best plant sources (DuCoa L.P., 1994). Most oil
seeds, such as peanuts, cottonseed, and soybeans, are good sources ofcholine because of their relatively high phospholipid content. Choline
content in foods is extremely variable, and much of this variation can be
attributed to biological variation. However, a large portion of the reported variability may be due to differences in procedures for choline
analysis.
Corn is low in choline, with wheat, barley, and oats containing approximately twice as much choline as corn. Since betaine can spare therequirements of choline, it would be useful to know the concentrationsof betaine in feeds. Unfortunately, most feedstuffs contain only small
amounts of betaine. However, wheat and wheat by-products apparently
577

CHAPTER FOURTEEN

contain over twice as much betaine as choline. Thus, the choline needs ofswine or poultry fed wheat-based diets would be much lower than those
fed diets based on other grains. Sugar beets are also high in betaine.
Choline is largely absent in most fruits and vegetables. However,other foods consumed by animals and humans can be good sources. Infant formulas based on soy protein have much less choline than humanor cow’s milk (Zeisel et al., 1986). Choline content of typical foods andfeedstuffs is shown in Table 14.2.
Little is known of the biological availability of choline in natural
feedstuffs. Using a chick assay method, soybean, canola, and peanut
meals were found to contain a substantial proportion of unavailable
choline (Emmert and Baker, 1997). Dehulled regular soybean meal andwhole soybeans were tested and appeared to range in availability from60 to 75% (Molitoris and Baker, 1976). However, soybean lecithin
products were found to be equivalent to choline chloride in bioavailability (Emmert et al., 1997). March and MacMillan (1980) presented
evidence that choline bioavailability in rapeseed meal is lower than insoybean meal. Canola meal, although three times as rich in total choline
as soybean meal, was found to have less bioavailable choline (Emmert
and Baker, 1997). In chicks, production of trimethylamine (resulting
from bacterial degradation of choline) in the intestine was greater inchicks fed rapeseed meal than in those fed soybean meal.

DEFICIENCY■ Table 14.2

Typical Choline Concentrations in Foods and Feedstuffs (ppm, dry basis)

Alfalfa meal, dehydrated
Bakery waste, dehydrated
BarleyBlood meal, dehydrated
Brewer's grains, dehydrated
Buttermilk (cattle), dehydrated
ChickenCitrus pulp, dehydrated
Copra (coconut) meal
Corn, yellow
Corn gluten feed
Cottonseed meal
Crab meal
Fish meal, anchovy

1,370
1,005
1,177
848
1,757
1,891
6,288
867
1,036
620
1,518
2,965
2,179
4,036

Meat and bone meal
Liver meal
Milk, skim, dehydrated
Molasses, sugarcane
Oats
Peanut meal
Potato, dehydrated
Rice
Rice bran
SesameSorghum
Soybean meal
Wheat
Wheat bran

1,996
12,281
1,480
1,012
1,116
2,120
2,879
1,076
1,357
1,655
737
2,916
1,053
1,797

Sources: Selected from United States-Canadian Tables of Feed Composition (NRC,1982) and DuCoa L.P. (1994).

578

CHOLINE

The most common signs of choline deficiency include poor growth,
fatty liver, perosis, hemorrhagic tissue (particularly in the kidney andcertain joints), and hypertension. In general, severity of clinical signs inanimal species is influenced by other dietary factors, including methionine, vitamin B12, folacin, and dietary fat (see Requirements). When feedintake—and consequently growth—are depressed by choline deficiency,
severity of choline deficiency is then reduced. Clinical signs and lesions
in choline-deficient animals are summarized in Table 14.3.

Effects of Deficiency
Ruminants
The ability of ruminants to synthesize the B vitamins including
choline is well known. In cattle and sheep, orally ingested choline is rapidly degraded in the rumen, but rumen microbes synthesize choline. It
has been found that ruminal synthesis and/or the unsupplemented diet
do not always supply enough choline to meet feedlot cattle demands.
Improved performance of feedlot cattle has been related to use of dietarycholine, although not consistently in all experiments. Reports from
Washington (Swingle and Dyer, 1970) and Maryland (Rumsey, 1975)
showed increased gains by as much as 6 to 7% and improved feed efficiency by 2.5 to 8% for finishing cattle when supplemented with 500 to
750 ppm dietary choline. In other experiments with growing cattle, noresponse occurred after addition of dietary choline (Wise et al., 1964;
Harris et al., 1966). Thus, choline, under certain conditions of high-concentrate feeding, may be limiting in the diet.
Researchers have shown interest in choline in view of its possible
lipotropic effect in high-producing dairy cows. Atkins et al. (1983) andErdman et al. (1984) studied the effects of supplementing choline on
roughage adequate diets for dairy cattle and concluded that added
choline had little effect on milk production, although slight increases
were seen in feed intake and milk fat percentage. Limited research indicates that supplemental choline may be needed to obtain optimum
milk fat, possibly through its effect on hepatic lipid metabolism. Lactating cows fed supplemental choline showed an increase in milk fat
percentage and fat-corrected milk (Erdman et al., 1984). In a later experiment, Erdman and Sharma (1991) were unable to show a positiveeffect on milk fat percentage from choline by postruminal infusion.
However, these experiments confirm earlier postruminal infusion studies in which increasing postruminal choline increased milk yield up to
3.1 kg/day.
579

■ Table 14.3

Clinical Signs and Lesions in Choline-Deficient Animals

Signs and Lesions

Rat

Fatty Liver
Cirrhosis
Hemorrhagic kidney
Growth failure
Hypolipemia

+
+
+
+
+

PerosisImpaired
reproduction
Impaired lactation
Reduced egg
production
Muscle defects
Spraddled hind legs++

Mouse Hamster
+
+a
+b
+

+

Fish

RabbitMonkey

Dog

Cat

Calf

Pig

Chicken Duck Turkey

+

+

+
+

+
+

+

+

+

+

+

+

+

+

+
+a
+b
+

+

+
+

+

+b
+

+b
+

+

+

+

+

+

+

+

+
+

+
+

+

+

+

+
+

Hemorrhagic lesions
Arterial sclerosis
Myocarditis andnecrosis
Bradycardia
Hypertension

+
+

+
+

+
+
+

+

AnemiaEdema, ascites

+
+

+

+

+

Sources: Modified from Wilson (1978). Information also from NRC (1988, 1993).
Note: In many cases, lesions have not been studied or reported in various species.
aProduced with difficulty; see text.
bRenal degeneration only.

+

+

+

CHOLINE

An apparent choline deficiency syndrome was produced with a synthetic milk diet containing 15% casein (Johnson et al., 1951). Within 6
to 8 days, calves developed extreme weakness with labored breathing
and were unable to stand. Supplementation with 260 mg of choline perliter of milk replacer prevented these deficiency signs.

Swine
Choline deficiency in the young pig results in unthriftiness, poor
conformation (short-legged and pot-bellied), lack of coordination in
movements, characteristic lack of proper rigidity in joints (particularly
the shoulders), fatty infiltration of liver, characteristic renal glomerular
occlusion, and some tubular epithelial necrosis (Cunha, 1977). These
clinical signs resulted from low-methionine diets (0.8%) and were prevented with 1.6% dietary methionine.
Spraddled-leg condition is a problem occasionally seen in newborn
pigs, and some evidence suggests that incidence has a strong genetic
component. However, the condition is often attributed to choline deficiency and is prevented by supplementation of the vitamin. Whether folacin and B12 are involved in the condition is unknown, but under conditions of deficiencies of these vitamins, choline requirements are
increased. Spraddled legs can be described as a congenital disorder inwhich the newborn pig cannot stand or walk because of the leg condition (Fig. 14.4). Nursing is also hindered, thereby affecting weaning
weights.
Spraddled-leg condition started to appear when swine producers began to decrease feed allowances given sows during gestation from 2.7 to
3.2 kg to 1.4 to 2.0 kg daily (Cunha, 1977), which resulted in reduced
intakes of both choline and methionine. Studies from Colombia, SouthAmerica (J.H. Maner, personal communication to Cunha, 1977), revealed death losses due to spraddled-leg condition. Some of these pigs
recuperated by the tenth day after birth, which could indicate that thecondition can be corrected through the sow’s milk. Other reports haveindicated that a high proportion of baby pigs affected by spraddled legs
were able to recover after a few days, especially if the hind legs werebound temporarily to allow them to move and suckle.
Protein, methionine, and sulfate concentrations of the diet influencecholine requirements. A study of the effects of protein level and supplemental choline on the reproductive performance of gilts indicated thatcholine increases pig weight and litter weight (Maxwell et al., 1987).
When dietary methionine is high, often there is no demonstrated response to supplementation of choline. This may be expected due to their581

CHAPTER FOURTEEN

Fig. 14.4 Choline deficiency. Spraddled hind legs—the condition shown
here—has been produced with a purified ration and is prevented by choline
supplementation. Other factors may be involved. (Courtesy of T.J. Cunha andWashington State University.)mutual sparing effect. Kroening and Pond (1967) showed increased gain
for 3-week-old pigs fed synthetic diets containing 1,640 ppm choline
and less than 0.31% methionine; although choline tended to improvefeed per gain in all diets containing up to 0.42% methionine. An investigation of the relationship of methionine, choline and sulfate in the dietof weanling swine indicated that choline and sulfate or methionine andsulfate improved performance (Lovett et al., 1985).
Research reports have shown that sows without choline have a significantly lower conception rate and farrowing rate and farrow significantly fewer total pigs and fewer live pigs per litter. No difference wasfound in the average birth weight, but sows with choline supplementation weaned significantly more pigs per litter, and sows without choline
farrowed a slightly higher percentage of pigs with spraddled legs. Pigs
from choline-deficient sows were unthrifty in appearance and became
increasingly so with age (Ensminger et al., 1947).
The NRC-42 committee on swine nutrition in 1976 evaluated the582

CHOLINE

effects of supplemental choline (770 mg/kg of diet) during gestation andlactation on litter size at birth and at weaning. Nine stations participated, with 22 trials and 551 sows. The diet was a 15.0% protein cornsoybean meal type during gestation, and 7.5% beet pulp was substituted
for an equal amount of corn during the lactation period. Results indicated that sows fed supplemental choline farrowed more pigs per litter(10.54 versus 9.89) and more live pigs per litter (9.33 versus 8.64), andweaned more pigs per litter (7.72 versus 7.29).
Betaine, the methyl donor derived from choline, has been shown tobe beneficial to swine (Odle, 1996). Shurson et al. (1994) concluded that
inclusion of betaine for more than 30 days prior to slaughter could, inmost cases, produce positive carcass effects, specifically reduced back fat
and increased yield. On the contrary, however, Øverland et al. (1999)
fed swine high fat diets supplemented with betaine but failed to improve
growth performance, carcass quality, or nutrient digestibility.
Haydon et al. (1995) reported positive betaine effects only with low
lysine:calorie feeds. Campbell et al. (1995) reported that when a methionine-deficient basal diet was supplemented with 0.125% betaine,
feed efficiency was improved to the same degree as when methionine
was supplemented at 0.23%.

Poultry
Growth retardation and perosis result from choline deficiency inyoung poultry. Perosis is the primary clinical sign of choline deficiency
in chicks and turkey poults, whereas bobwhite quail develop enlarged
hocks and bowed legs (NRC, 1994). Perosis is characterized first by pinpoint hemorrhages about the hock joint, followed by an apparent flattening of the tibiometatarsal joint (Scott et al., 1982). Progressively, theAchilles tendon slips from its condyles, thus rendering the bird relatively
immobile. Some studies indicated that in prevention of perosis, choline
is required for the phospholipids needed for normal maturation of the
cartilage matrix of bone.
Adult chickens probably synthesize sufficient choline to meet requirements for egg production. Minimal dietary choline does not affecthatchability with either chickens or turkeys, but Japanese quail and theirdeveloping embryos readily express general signs of deficiency (Latshaw
and Jensen, 1972; NRC, 1994). Supplementary choline may be necessary for maintenance of egg size in quail (NRC, 1994). Contrary to some
reports, 500 ppm supplemental choline to Leghorn hens increased egg
weight while reducing specific gravity (Tapia Romero et al., 1985). Also,
the choline growth requirement for quail is apparently higher than that583

CHAPTER FOURTEEN

for chicks or poults.
Choline requirement of growing chicks decreases with age, and it isgenerally not possible to produce a deficiency at an age over 8 weeks. It
was observed that methylation of aminoethanol to methylaminoethanol
seems to be the rate-limiting step in choline biosynthesis for young birds.
High levels of dietary methionine or other methyl donors, therefore,cannot completely spare the chick’s requirement for dietary choline,
which is in contrast to the situation with growing mammals such as thepig or the rat.
Apparently, choline requirement of laying hens can be influenced bycholine level in the diet of the growing pullet (Scott et al., 1982). Hens
that received choline-free diets after 8 weeks of age were able to synthesize all the choline required for good egg production. Those that received choline supplements in the growing diet required supplementalcholine in the laying diet for maximum egg production. The deficiency
signs noted in these hens were a reduction in egg production and an increase in fat content of liver. Even with choline deficiency, however,choline content of the egg was not affected by low dietary choline.
Despite lack of evidence that laying chickens require a dietary
source of choline for maximum egg production, addition of choline topractical diets markedly reduces the amount of fat in the liver (NRC,
1994). However, a number of reports with chicks and turkey poults did
not find fatty livers in chicks deficient in choline (Ruiz et al., 1983). A
choline response in laying chickens is likely to occur only if inadequate
daily sulfur amino acid is provided.
Addition of 0.1% of supplemental methionine resulted in no response in laying hens to supplemental choline (Crawford et al., 1969).
It appears that benefits from supplemental choline in layer diets occur
mainly when supplemental methionine is just adequate to meet methionine requirements. Miles and Harms (1983) demonstrated that the addition of 0.11% choline plus 0.1% sulfate could essentially spare all
supplemental methionine in broiler diets. However, in turkey poult diets(Harms and Miles, 1984), responses to sulfate and choline addition were
not equivalent to the addition of supplemental methionine. Pesti et al.
(1980), using young chicks, found that supplementation with methyl
donors from either 0.23% choline or 0.23% betaine was equivalent to
supplementation with 0.23% methionine in 21-day chick experiments,using basal diets containing 0.31% methionine and 0.43% cystine.
Spires et al. (1982) found that supplemental choline could replace up to
two-thirds of the supplemental methionine required in broiler diets from0 to 47 days in diets containing 0.30% methionine and 0.43% cystine584

CHOLINE

in the starter phase, and 0.25 and 0.42% methionine and cystine, respectively, in the finisher phase.

HorsesNo studies of choline requirements or deficiencies have been undertaken in the horse.

Other Animal Species
DOGS AND CATS
In growing cats, choline deficiency results in perilobular infiltrationof the liver, hypoalbuminemia, and decreased growth (NRC, 1986). In
young puppies, choline deficiency results in reduced appetite and growthrate, fatty metamorphosis of the liver, and atrophic changes of the thymus. Choline-deficient dogs with fatty livers show an increased rate ofhepatic phospholipid synthesis following choline supplementation(NRC, 1985a).

FISH
Dietary choline is required for optimum growth; feed utilization;and prevention of fatty livers, anemia, hemorrhages (in liver, kidney, andintestine), and other clinical signs in striped bass, salmon, trout, carp,
catfish, and bream (NRC, 1993; Griffin et al., 1994). Severity and nature of choline deficiency signs, as well as the time required for them toappear, seem to be highly variable and may depend on species or diet or
both (Ketola, 1976). Channel catfish fed casein-gelatin diets containing
excess methionine did not develop signs of choline deficiency; however,catfish fed diets adequate but not excessive in methionine did develop
deficiency signs (Wilson and Poe, 1988).
Rainbow trout fed a choline-deficient diet developed light yellowcolored livers, protruded eyes, anemia, and extended abdomens (Kitamura et al., 1967), while lake trout fed a choline-deficient diet for 12
weeks had depressed growth rate and increased liver fat content (Ketola,1976). Common carp were found to grow normally on a choline-deficient diet but had 10% higher liver lipid content (Ogino et al., 1970).
Contrary to other fish and different species, the red drum had reduced
liver lipid rather than high accumulation with choline deficiency (Craig
and Gatlin, 1996). Choline-deficient Japanese eels became anorexic, hadpoor growth, and exhibited a white-gray intestine. After 4 weeks,
shrimp fed a choline-deficient diet were significantly smaller than controls (NRC, 1993).
585

CHAPTER FOURTEENLABORATORY ANIMALS
Guinea pigs, like young poultry, easily develop choline deficiency
even with adequate dietary methionine. Clinical signs of choline deficiency in young guinea pigs include poor growth, anemia, muscular
weakness, and some adrenal subcutaneous hemorrhages (NRC, 1995).
In guinea pigs, choline supplementation altered carnitine homeostasis bydecreasing urinary excretion of carnitine (Daily and Sachan, 1995).
Hamsters showed poor appetite, reduced growth, and fatty livers on apeanut meal diet deficient in choline (Handler and Bernheim, 1949).
Both rats and mice exhibit fatty livers, but rats—unlike mice—also
develop cirrhosis or fibrosis of the liver. In young rats the kidney becomes hemorrhagic, owing presumably to a deficiency of choline for thephospholipid required to build cell structure at this critical growth period. Mice, on the other hand, develop hemorrhagic kidneys only withmore difficulty and diet manipulation. Administration of lecithin increased brain acetylcholine concentration and improved memory in micewith dementia (Chung et al., 1995). Severe folate deficiency caused secondary depletion of choline and phosphocholine in rat liver (Kim et al.,1994).

MONKEYS
Cebus monkeys, rhesus monkeys, and baboons fed choline-deficient
diets for 1 to 2 years developed fatty changes in the liver and varying degrees of hepatic fibrosis (NRC, 1978).

RABBITS
The signs of choline deficiency in rabbits have been described as retarded growth, fatty and cirrhotic liver, and necrosis of the kidney
tubules. Progressive muscular dystrophy was reported in rabbits fed a
low-choline diet for more than 70 days (NRC, 1977).

HumansIn humans, as in experimental animals, dietary choline can arrest
cirrhosis of the liver and reverse the fatty infiltration. However, these
conclusions are not definite, and some reports show no benefit ofcholine in the treatment of cirrhosis (Kuksis and Mookerjea, 1984). It
remains uncertain whether alcoholic cirrhosis in humans and animals is
caused by one or more nutritional deficiencies, but accepted dietary regimens for treatment involve a high-calorie, high-protein, low-fat dietsupplemented with choline and perhaps other lipotropic factors, such as
methionine and inositol.
586

CHOLINE

Choline is an essential nutrient for humans when excess methionineand folacin are not available in the diet. Healthy male subjects fed acholine-deficient diet for 3 weeks had depleted stores of choline in tissues and developed signs of incipient liver dysfunction (Zeisel et al.,1991). People receiving total parenteral nutrition have developed fatty
infiltration of the liver and hepatocellular damage. One study in humans
fed parenterally showed that supplemental choline (in the form oflecithin) could reverse fatty livers (Buchman et al., 1992).
The relationship of choline to atherosclerosis has been under study
for a number of years. Choline-deficient diets can result in arterial damage, with a high percentage of young rats maintained on low-choline diets developing pathological changes in the major arterial trunks and in
the coronary arteries. These changes are similar to those observed inearly stages of atherosclerosis of the aorta and large arteries in humans.
In a 3-year study of coronary thrombosis patients, there was a significant reduction in death rates and lowered blood cholesterol in atheromatous patients administered choline (DuCoa L.P., 1994). During acute
myocardial ischemia, plasmalogen (a choline phospholipid found in thecell membranes of heart muscle) was broken down (Canty and Zeisel,
1994). Further observations have included increases in high-density
lipoprotein and decreases in low-density lipoprotein following lecithintreatment in hypercholesterolemic humans (Childs et al., 1981).
Deficiency of choline enhances the initiating potency of several carcinogens (Mikol et al., 1983) and exerts a strong promoting effect withliver carcinogens. Choline deficiency alone can also lead directly to liver
tumor formation (Mikol et al., 1983). A number of possible mechanisms
have been proposed to account for the development of cancer as a resultof choline deficiency, including increased cell death followed by increased proliferation and regeneration, deceased DNA methylation andrepair, increased lipid peroxidation and free radical damage, and decreased methylation and detoxification of carcinogens (Canty and
Zeisel, 1994).
Reports indicate that lecithin and/or choline supplements may be
useful in preventing age-related memory deficits and certain neurological diseases (Kuksis and Mookerjea, 1984). Evidence suggests that dietary choline and lecithin produce clinical improvement by supplying
precursors for formation of the neurotransmitter acetylcholine. Notable
but sometimes limited success has been achieved with choline or lecithin
therapy for tardive dyskinesia syndrome (jerky involuntary movements
of face and extremities), Huntington’s disease (rapid involuntary andpurposeless movements), and Friedreich’s ataxia disease (loss of coordi587

CHAPTER FOURTEEN

nation in muscles of the extremities). Studies of both Alzheimer’s disease(deficiency of hippocampal cholinergic neurons) and senile dementia(forgetfulness) patients given choline or lecithin have shown mixed results (Wood and Allison, 1982). Smith et al. (1978), in a controlled experiment, noted some improvement in about 40% of the cases with advanced Alzheimer’s syndrome. In general, studies have found thatpatients with less severe symptoms show greater response to choline
supplementation. Thus, the use of choline in severe cases of Alzheimer’s
disease does not appear to be very helpful, but may be of value in earlystages of memory-loss disorders (DuCoa L.P., 1994).

Assessment of Status
Research information is very limited on detection methods to determine choline status of animals. Often the best indicator of status or needfor choline is observation of clinical signs attributable to choline deficiency (e.g., fatty livers or perosis) in particular species as well as beneficial performance responses when diets are supplemented with the vitamin.
Tissue levels of choline or its functional metabolites can be determined to evaluate choline status. There is evidence of a reduction ofacetylcholine in brains, kidneys, and intestines of rats deprived ofcholine 6 days after weaning. Choline administered to rats either by injection or by diet caused a dose-related increase in brain acetylcholine(Kuksis and Mookerjea, 1984). Studies on mechanism of liver fat accumulation have suggested that this is related to a lack of phosphatidylcholine synthesis. With choline deficiency, the hepatic phosphatidylcholine:phosphatidylethanolamine ratio is reduced, and is thus a meansof evaluating choline status.
Plasma choline levels in humans reflect dietary consumption (Hirsch
et al., 1978). There is the suggestion that fall of free plasma choline below the normal range may be related to a choline-deficiency state. Afterconsumption of 3 g of choline chloride in a meal, serum choline rose by
86% over the mean fasting level of 11.7 nmol/ml.

SUPPLEMENTATION
Choline is made synthetically, and, in the majority of cases, diet supplementation is from synthesized choline salts. Choline is synthesizedfrom natural gas via methanol and ammonia, which are reacted to produce trimethylamine (Griffith and Nyc, 1971). This trimethylamine is588

CHOLINE

subsequently reacted with ethylene oxide to produce choline. For feed
supplementation purposes, a chloride salt is produced by reacting the alkaline base with hydrochloric acid. Choline is available for feed use as
the 70 or 75% liquid or 25 to 60% dry powder. Choline chloride can beavailable on a cereal carrier; the product is obtained by spraying andthoroughly mixing aqueous choline chloride on a suitable cereal carrier
and then drying to a low moisture content. The 70 to 75% liquid is very
corrosive and requires special storage and handling equipment. It is notsuitable for inclusion in concentrated vitamin premixes, but rather ismost generally added singly to concentrate mixtures.
A variety of lecithin products that are derived from soybeans areavailable for use in feeds. The products range from crude fluid lecithin
to 50% lecithin codried with corn syrup solids to a dry deoiled soy
lecithin (Meyers, 1990). The advantage of a deoiled soybean lecithin isthat it can be handled as a dry feed ingredient. The dry deoiled lecithin
product also possesses those advantageous properties of soybean lecithinin general that impart important dietary formulation features, including
vitamin stabilization (thus protecting vitamins from oxidation); improvement of fat and vitamin utilization; and a source of choline, inositol, and growth-stimulating compounds.
Some research has shown that supplemental betaine, a product ofcholine oxidation, is beneficial to swine and poultry production (Lowry
et al., 1987; Odle, 1996). Supplemental betaine is available as betainehydrochloride (98.0% betaine on an anhydrous basis).
Response to dietary supplementation of choline will be most dependent on species, age of animals, protein and sulfur amino acid intake,dietary choline, and other choline-sparing nutrients. Unlike most vitamins, choline can be synthesized by various animals, although often in
insufficient amounts. It appears that choline deficiency in the young ofsome species (e.g., chick) may not be due to lack of ability to synthesize
choline but more likely to lack of ability to synthesize it at a rate sufficient for animal needs. Age is an important consideration; for example,
it is difficult to produce choline deficiency in growing chicks more than
8 weeks of age.
The young of many species (e.g., pig and rat) do not require supplementary choline if dietary methionine level is sufficiently high. On asynthetic milk diet containing 1.6% methionine, young pigs were found
not to require supplemental choline (Firth et al., 1953). Neumann et al.
(1949) reported that the young pig requires 0.1% dietary choline whenmethionine is present at 0.8 to 1.0% of the diet.

589

CHAPTER FOURTEEN

Methionine can furnish methyl groups for choline synthesis for most
species. Choline, however, is effective only in sparing methionine that
otherwise would be used to make up for a choline shortage. Methionine
is not used for choline synthesis if there is an adequate level of dietarycholine. In formulating typical poultry and swine diets, methionine isfrequently one of the most limiting amino acids. Therefore, it would be
impractical for marginal quantities of methionine to be wasted for synthesis of the vitamin when supplemental choline can be provided more
economically.
Interrelationships of choline and methionine are discussed in the
Functions and Requirements sections. In providing supplements of methionine and/or choline (or betaine), an additional nutrient, sulfur, must
be considered. Significance of a four-way interrelationship among methionine, choline, betaine, and sulfate has been reviewed (Ruiz et al.,
1983; Miles et al., 1986; Miles and Butcher, 1997). Sulfur is present ina number of body metabolites (e.g., mucopolysaccharides) and, if notadequately supplied in the diet, sulfur amino acids would likely be degraded. In feeding broilers, supplemental sulfate accompanied bycholine or methionine achieved a greater growth response than when either was fed alone (Miles et al., 1983). Data suggest that sulfate must bepresent for choline to spare a maximum amount of methionine. The
practical implication is that sulfate and choline need to be adequately
provided in diets so that the more expensive and often marginally deficient nutrient methionine is not used to provide either of these nutrients.
Most choline supplementation studies emphasize production benefits from providing the vitamin to young animals. However, researchand observations with adult swine have demonstrated improved litter
size at weaning, and supplementation may keep sows in the producing
herd longer (Cunha, 1977). The exact level of choline needed for sow rations is unknown. Until more research data are available, Cunha (1972)
suggested the following levels in situations where spraddled hind legs arelikely to occur: (1) during the first part of gestation, a daily level of
3,000 mg per sow and (2) the last month of gestation, when dailycholine should be increased to 4,200 to 4,500 mg.
Supplemental choline has been shown to be beneficial to young ruminants, such as young calves prior to ruminal development. However,
it is apparent that traditional supplements of choline in the form ofcholine chloride are extensively degraded in the rumen. For lactating
dairy cows or other ruminants that may have need for supplementalcholine, it should be protected from rumen degradation and available

590

CHOLINE

for absorption in the small intestine. A rumen-protected choline productthat could potentially be used as supplemental choline for ruminants isavailable. In vitro studies showed that at least 87% of the choline wasprotected from rumen degradation (Erdman and Sharma, 1991). Studies
of postruminal choline infusion and studies with rumen-protected formsof choline at 31 to 45 g/day supported a 1.9-kg increase in milk production (Erdman and Sharma, 1991; Erdman, 1992).
It is likely that a large portion of methionine is used as a methyl
donor for choline synthesis in ruminants. This suggests that choline
would be a more likely candidate for rumen-protected supplementation
than methionine (Erdman and Sharma, 1991). Diets in which methionine or, indirectly, protein was most limiting, would most likely benefit
from rumen-protected choline addition.
For humans, in view of the widespread occurrence of choline andmethionine in plant and animal foodstuffs, choline supplementation ismost warranted for individuals on special dietary regimens, including infants and young children fed diets deficient in protein and high in highly
refined products and patients on total parenteral nutrition. Also, supplemental choline has been used with some success to diminish shortterm memory loss associated with Alzheimer’s disease and certain otherneurological diseases. Supplemental choline has a positive benefit in reducing cancer and atherosclerosis (see Deficiency, Effects of Deficiency,
Humans). Cancer patients receiving methotrexate treatment should consider receiving choline supplementation, as some data suggest that fatty
infiltration of the liver associated with methotrexate treatment occurs
because of a disturbance in choline metabolism (Pomfret et al., 1990).
Because 5 to 14 mg of total choline per deciliter is present in humanmilk and is tolerated with no adverse reactions, and because deficiency
in young animals produces serious effects, it is recommended that atleast 7 mg of choline per 100 kcal be included in infant formulas notbased on milk (Kuksis and Mookerjea, 1984). Commercially prepared
infant formulas and milk products have been supplemented with choline
at these levels to ensure the presence of choline in an amount approximating that naturally occurring in milk.
The most widely used supplemental form of choline is choline chloride, which is stable in multivitamin premixes but is highly destructive
to various other vitamins in the premix (Frye, 1978; Coelho, 1991).
Choline is stable during processing and storage in pressure-pelleted andextruded feeds. Since the material is hygroscopic, containers of choline
should be kept closed when not in use.

591

CHAPTER FOURTEENTOXICITY
Experimental animal toxicity data on clinical signs of choline overdosage include salivation, trembling, jerking, cyanosis, convulsions, andrespiratory paralysis. Estimates of the oral LD50 of choline chloride inrats varied from 3.4 to 6.7 g/kg (Chan 1991). Bell (1985) reported thatwhen rats were exposed perinatally to 22 mg of soy lecithin preparation
daily, sensorimotor development and brain cell maturation were altered.
Choline levels somewhat above the requirement (868 to 2,000 ppm)
were shown to reduce rate and efficiency of gain in swine (Neumann etal., 1949; Southern et al., 1986). Derilo and Balnave (1980) reported reduced gain and efficiency in young broiler chicks fed a level of choline
only slightly in excess of the requirement. Studies with chickens suggestthat dietary choline double the requirement is safe, with swine having ahigher tolerance for choline (NRC, 1987). Work by Southern et al.
(1986) showed that excess choline, up to 2,000 mg/kg above the recommended level, had no adverse effect on swine performance.
In humans, high intakes of lecithin or choline produced acute gastrointestinal distress, sweating, salivation, and anorexia (Wood and Allison, 1982). However, therapeutic doses of choline chloride and ofcholine dihydrogen citrate administered in amounts ranging from 3 to
12 g/day have been used in the treatment of alcoholic cirrhosis for up to
4 months with no toxic effects reported. In treatment of tardive dyskinesia (a movement disorder generally associated with the intake of antipsychotic medication), up to 16 g/day of choline has been used; inother cases, lecithin at doses greater than 100 g/day for more than 4months has been used with no evidence of ill effects (Kuksis and Mookerjea, 1984).

■ REFERENCES
Ansell, G.B., and Spanner, S. (1971). Biochem J. 122, 741.
AOAC (Association of Official Agricultural Chemists), (1984). Official Methodof Analysis. AOAC, Washington, D.C.
Atkins, K.B., Erdman, R.A., and Vandersall, J.H. (1983). J. Dairy Sci. 66(Suppl.
1), 175.
Atkins, K.B., Erdman, R.A., and Vandersall, J.H. (1988). J. Dairy Sci. 71, 109.
Barnes, R.H., and Kwong, E. (1967). J. Nutr. 92, 224.
Bell, J.M. (1985). Devel. Psychobiol. 18, 383.
Best, C.H., Channon, H.J., and Ridout, J.H. (1934). J. Physiol. (Lond.) 78, 409.
592

CHOLINE

Brown, E.S. (1964). Am J. Physiol. 207, 402.
Buchman, A.L., Dubin, M., Jenden, D., and Moukarzel, A. (1992). Gastroenterology 102, 1363.
Campbell, R.G., Cadogan, D.J., Morley, W.C., Uusitalo, R., and Virtanen, E.
(1995). J. Anim. Sci. 73(Suppl. 1), 82(Abstr.).
Canty, D.J., and Zeisel, S.H. (1994). Nutr. Rev. 52, 327.
Chan, M.M. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), 2nd Ed., p.
537. Marcel Dekker, New York.
Chao, C.K., Pomfret, E.A., and Zeisel, S.H. (1988). Biochem. J. 254, 33.
Childs, M.T., Bowlin, J.A., Ogilvie, J.T., Hazzard, W.R., and Albers, J.J. (1981).
Atherosclerosis 38, 217.
Chung, S., Moriyama, T., Uezu, E., Uezu, K., Hirata, R., Yohena, N., Masuda,Y., Kokubu, T., and Yamamoto, S. (1995) J. Nutr. 125, 1484.
Coelho, M.B. (1991). Vitamin Stability in Premixes and Feeds: A Practical Approach, p. 56. BASF Technical Symposium, Bloomington, Minnesota.
Craig, S.R., and Gatlin, D.M. III (1996). J. Nutr. 126, 1696.
Crawford, J.S., Griffith, M., Teekell, R.A., and Watts, A.B. (1969). Poult. Sci.
38, 620.
Cunha, T.J. (1972). Feedstuffs 44, 27.
Cunha, T.J. (1977). Swine Feeding and Nutrition, Academic Press, New York.
Daily, J.W. III, and Sachan, D.S. (1995). J. Nutr. 125, 1938.
Dale, H.H., and Dudley, H.W. (1929). J. Physiol. (Lond.) 68, 97.
De La Huerga, J., and Popper, H. (1952). J. Clin. Invest. 31, 598.
Derilo, Y.L., and Balnave, D. (1980). Poult. Sci. 21, 479.
DuCoa L.P. (1994). In Choline Functions and Requirements, p. 91.
DuPont/ConAgra Co., Higland, Illinois.
Du Vigneaud, V., Chandler, J.P., Moyer, A.W., and Keppel, D.M. (1939). J. Biol.
Chem. 131, 57.
Emmanual, B., and Kennelly, J.J. (1984). J. Dairy Sci. 67, 1912.
Emmert, J.L., and Baker, D.H. (1997). J. Nutr. 127, 745.
Emmert, J.L., Garrow, T.A., and Baker, D.H. (1996). J. Nutr. 126, 2050.
Emmert, J.L., Garrow, T.A., and Baker, D.H. (1997). J. Anim. Sci. 74, 2738.
Ensminger, M.E., Bowland, J.P., and Cunha, T.J. (1947). J. Anim. Sci. 6, 409.
Erdman, R.A. (1992). Vitamins. Abstracted Proc. Large Dairy Herd Management Symposium, p. 24. Gainesville, Florida.
Erdman, R.A., and Sharma, B.K. (1991). J. Dairy Sci. 74, 1641.
Erdman, R.A., Shaver, R.D., and Vandersall, J.H. (1984). J. Dairy Sci. 67, 410.
Finkelstein, J.D., Martin, J.J., Harris, B.J., and Kyle, W.E. (1982). Arch,
Biochem. Biophys. 218, 169.
Firth, J., James, M., Chang, S., Mistry, P., and Johnson, B.C. (1953). J. Anim.
Sci. 12, 915(Abstr.).
Frye, T.M. (1978). In Proc. Roche Vitam. Nutr. Update Meet., Arkansas Nutr.
Conf., p. 54. Hoffmann-La Roche, Nutley, New Jersey.
Griffin, M.E., Wilson, K.A., White, M.R., and Brown, P.B. (1994). J. Nutr. 124,
1685.
Griffith, W.H., and Nyc, J.F. (1971). In The Vitamins (W.H. Sebrell Jr., and R.S.
Harris, eds.), Vol. 3, p. 3. Academic Press, New York.
Hall, C.E., and Bieri, J.G. (1953). Endocrinology 53, 661.
593

CHAPTER FOURTEEN

Handler, P., and Bernheim, F. (1949). Proc. Soc. Exp. Biol. Med. 72, 569.
Harms, R.H., and Miles, R.D. (1984). Poult. Sci. 63, 1464.
Harris, R.R., Yeates, H.F., and Barrett, J.E. Jr. (1966). J. Anim. Sci. 25, 248 (Abstr.).
Hartroft, W.S. (1955). Proc. Am. Soc. Exp.Biol. 14, 655.
Haydon, K.D., Campbell, R.G., and Prince, T.J. (1995). J. Anim. Sci. 73,(Suppl.
1), 83(Abstr.)
Hegazy, E., and Schwenk, M. (1984). J. Nutr. 114, 2217.
Hirsch, M.J., Growdon, J.H., and Wurtman, R.J. (1978) Metabolism 27, 953.
Johnson, B.C., Mitchell, H.H., Pinkos, J.A., and Merrill, C.C. (1951). J. Nutr.
43, 37.
Jukes, T.H. (1947). Annu. Rev. Biochem. 16, 193.
Ketola, H.G. (1976). J. Anim. Sci. 43, 474.
Ketola, H.G., and Nesheim, M.C. (1974). J. Nutr. 104, 1484.
Kim, Y., Miller, J.W., Da Costa, K., Nadeau, M. Smith, D. Selhub, J., Zeisel,
S.H., and Mason, J.B. (1994). J. Nutr. 124, 2197.
Kitamura, S., Suwa, T., Ohara, S., and Nakagawa, K. (1967). Bull. Jpn. Soc. Sci.
Fish. 33, 1126.
Kroening, G.H., and Pond, W.G. (1967). J. Anim. Sci. 26, 352.
Kuksis, A., and Mookerjea, S. (1984). In Present Knowledge in Nutrition (R.E.
Olson, H.P. Broquist, C.O. Chichester, W.J. Darby, A.C. Kolbye, and R.M.
Stalvey, eds.), p. 383. Nutrition Foundation, Washington, D.C.
Latshaw, J.D., and Jensen, L.S. (1972). J. Nutr 102, 749.
Lombardi, B., Ugazio, G., and Raick, A. (1966). Am. J. Physiol. 210, 31.
Lovett, T.D., Coffey, M.T., and Miles, R.D. (1985). J. Anim. Sci. 63, 467.
Lowry, K.R., Izquierdo, O.A., and Baker, D.H. (1987). Poult. Sci. 66(Suppl. 1),
135.
March, B.E., and MacMillan, C. (1980). Poult. Sci. 59, 611.
Maxwell, C.V., Johnson, R.K., and Luce, W.G. (1987). J. Anim. Sci. 64, 1044.
Meyers, S.P. (1990). Feed Manage. 41(8), 12.
Mikol, Y.G., Hoover, K.L., Creasia, D., and Poirier, L.A. (1983). Carcinogenesis. 4, 1619.
Miles, R.D., and Butcher, G.D. (1997). Industria Avicola 44(2), 29.
Miles, R.D., and Harms, R.H. (1983). In Proc. 1983 Florida Nutrition Conference, p. 75. Gainesville, Florida.
Miles, R.D., Ruiz, N., and Harms, H. (1983) Poult. Sci. 62, 495.
Miles, R.D., Ruiz, N., and Harms, R.H. (1986). Prof. Anim. Sci. 2, 33.
Molitoris, B.A., and Baker, D.H. (1976). J. Anim. Sci. 47, 481.
Mookerjea, S. (1971). Proc. Fed. Am. Soc. Exp. Biol. 30, 143.
Neill, A.R., Grime, D.W., Snoswell, A.M., Northrop, A.J., Lindsey, D.B., andDawson, R.M.C. (1979). Biochem. J. 180, 559.
Neumann, A.L., Krider, J.L., James, M.F., and Johnson, B.C. (1949). J. Nutr. 38,
195.
Norvell, M.J., and Nesheim, M.C. (1969). Proc. Cornell Nutr. Conf., p. 321.
Ithaca, New York.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
594

CHOLINE(1978). Nutrient Requirements of Nonhuman Primates.
(1981). Nutrient Requirements of Goats.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1985b). Nutrient Requirements of Sheep, 5th Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 4th Ed.
(1996). Nutrient Requirements of Beef Cattle, 7th Ed.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed. National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1976). Committee on Swine Nutrition. J. Anim. Sci. 42, 1211.
Odle, J. (1996). Feed Manage. 47(1), 25.
Ogino, C., Uki, N., Watanabe, T., Iida, Z., and Ando, K. (1970). Bull. Jpn. Soc.
Sci. Fish. 36, 1140.
Olson, R.E. (1959). Annu. Rev. Biochem. 28, 467.
Øverland, M., Røvik, K.A., and Skrede, A. (1999). J. Anim. Sci. 77, 2143.
Pesti, G.M., Benevenga, N.J., Harper, A.E., and Sunde, M.L. (1981). Poult. Sci.
60, 425.
Pesti, G.M., Harper, A.E., and Sunde, M.L. (1980). Poult. Sci. 59, 1073.
Pomfret, E.A., da Costa, K.A., Schurman, L.L., and Zeisel, S.H. (1989). Anal.
Biochem. 180, 85.
Pomfret, E.A., da Costa, K., and Zeisel, S.H. (1990). J. Nutr. Biochem. 1, 533.
RDA. (1989). Recommended Dietary Allowances, 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
Rohlfs, E.M., Garner, S.C., Mar, M., and Zeisel, S.H. (1993). J. Nutr. 123, 1762.
Ruiz, N., Miles, R.D., and Harms, R.H. (1983). WPSA J. 39, 185.
Rumsey, T.S. (1975). Feedstuffs 47, 30.
Sanford, P.A., and Smyth, D.H. (1971). J. Physiol. (Lond.) 215, 769.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Sharma, B.K., and Erdman, R.A. (1988). J. Dairy Sci. 71, 2670.
Shurson, J., Salzer, T., Johnston, L., White, M., Hathaway, M., Dayton, W., andWalker, B. (1994). In Proc. Minnesota Nutrition Conference and Roche
Technical Symposium, Bloomington, Minnesota.
Smith, C.M., Swase, M., Exton-Smith, A.N., Phillips, M.J., Overstall, P.W.,
Piper, M.E., and Bailey, M.R. (1978). Lancet. 2, 318.
Southern, L.L., Brown, D.R., Werner, D.D., and Fox, M.C. (1986). J. Anim. Sci.
62, 992.
Spires, H.R., Botts, R.L., and King, B.D. (1982). Syntex Research Report, Series
A, No. 1.
Swingle, R.S., and Dyer, I.A. (1970). J. Anim. Sci. 31, 404.
Tapia Romero, E., Rojas, R.E., Arias, L.E., and Avila, G.E. (1985). In Re595

CHAPTER FOURTEEN

sumenes ALPA 85 (C.F.Chicco, ed.), p. 49 (Abstr.). Acapulco, Mexico.
Wauben, P.M., and Wainwright, P.E. (1999). Nutr. Rev. 57, 35.
Welch, B.E., and Couch, J.R. (1955). Poult. Sci. 34, 217.
Wilson, R.B. (1978). In Handbook Series in Nutrition and Food, Section E: Nutritional Disorders (M. Rechcigl, Jr., ed.), Vol. 2, p. 95. CRC Press, Boca
Raton, Florida.
Wilson, R.P., and Poe, W.E. (1988). Aquaculture 68, 65.
Wise, M.B., Blumer, T.N., and Barrick, E.R. (1964). North Carolina Agri, Exp.
Sta. ANS Report 139. A.H. Series 10, 22.
Wood, J.L., and Allison, R.G. (1982). Fed. Proc. 41, 3015.
Zeisel, S.H. (1990). J. Nutr. Biochem. 1, 332.
Zeisel, S.H., and da Costa, K.A. (1990). J. Nutr. Biochem. 1, 55.
Zeisel, S.H., Char, D., and Sheard, N.F. (1986). J. Nutr. 116, 50.
Zeisel, S.H., Costa, K.A., Franklin, P.D., Alexander, E.A., Lamont, J.T., Sheard,
N.F., Beiser, A., and Costa, K.A. (1991). FASEB J. 5, 2093.

596

chapter fifteen

V I TA M I N C

INTRODUCTION
Scurvy, a potentially fatal condition resulting from inadequate vitamin C (ascorbic acid), has been known and feared since ancient times.
Its prevention and cure were associated with consumption of fresh
fruits, especially citrus, but it was not until 1928 that the antiscorbutic
factor was identified. Vitamin C is synthesized in almost all species, theexceptions being the primates, including humans, guinea pigs, fish, fruiteating bats, insects, and some birds. Animals that cannot synthesize thisvitamin need a dietary source for their normal maintenance.
The concept that the sole function of vitamin C is to prevent scurvy
has been revised in recent years. Small quantities of vitamin C are sufficient to prevent and cure scurvy; however, larger quantities may be required to maintain good health during conditions of adverse environment, physiological stress, and certain diseases. Antioxidant vitamins(vitamin C, vitamin E, and β-carotene) have received a great deal of attention in that they play important roles in animal and human health byinactivating harmful free radicals produced through normal cellular activity and from various stressors.

HISTORY
Several historical reviews of scurvy and vitamin C are available(Marks, 1975; Vilter, 1978; Moser and Bendich, 1991). Carpenter(1986) wrote a fascinating book, The History of Scurvy and Vitamin C,
that deals with the problems of scurvy throughout history. A historical
record is presented in Table 15.1 (Vilter, 1978).
If famine is excluded, scurvy is probably the nutritional deficiency
597

15

■ Table 15.1

Historical Record of Scurvy and Its Relationship to Vitamin C

1550 B.C.
600 B.C.
A.D. 1200
1492–1600

Scurvy described in Eber’s Papyrus (Thebes).
Hippocrates described soldiers afflicted with scurvy.
Crusaders weakened by scurvy.
World exploration threatened by scurvy.
—Magellan lost four-fifths of his crew.
—Vasco de Gama lost 100 of his 160 men.
Jacques Cartier’s expedition immobilized by scurvy; learned from Indiansthe curative value of pine needles and bark.
Captain James Lancaster prevented scurvy by giving crew members twojiggers of lemon juice daily.
Sir Richard Hawkins used oranges and lemons to treat scurvy in the
British Navy; Ponsseus referred to the therapeutic use of scurvy grass,watercress, and oranges.
Infantile scurvy described by Glisson but was confused with rickets.
Lime juice used experimentally on ships of the East India Company.
Backstrom related scurvy to a deficiency of fresh fruits and vegetables.
Lord Anson lost three-fifths of his crew of 1,950 men to scurvy.
James Lind performed controlled shipboard experiment on the preventive effect of oranges and lemons (published 1753).
Captain James Cook demonstrated that prolonged sea voyages were possible without ravages of scurvy.
William Stark induced scurvy in himself by a diet of bread and water for
60 days.
Lemon juice made a regular ration in British Navy.
Lemon juice made a regular ration in British Merchant Marines.
Scurvy epidemic occurred in the opposing armies during the American
Civil War.
Sir Thomas Barlow differentiated infantile scurvy from rickets.
Antiscorbutic ration became official in U.S. Army.
Boiling and pasteurization of infant formula increased incidence ofscurvy.
Hopkins suggested that infantile scurvy was a deficiency disease.
Hoist and Frolich produced experimental scurvy in guinea pigs by feeding a deficient diet, with pathological changes resembling those in humans.
Explorer Captain Scott and his team died of scurvy during their expedition to the South Pole.
Szent-Györgyi isolated hexuronic acid from orange juice, cabbage juice,and cattle adrenal glands.
Waugh and King isolated hexuronic acid from lemons and identified it asvitamin C.
Haworth determined structure of vitamin C.
Reichstein synthesized vitamin C.
Linus Pauling published book on relationship of vitamin C to the common cold, which stimulated research on therapeutic and prophylactic
uses of megadoses of vitamin C on the common cold, resistance, andvarious diseases.

1536
1570
1593
1650
17th century
1734
1740–1744
1747
1768–1771
1789
1795
1854
1863
1883
1895
1900
1906
1907
1912
1928
1932
1933
1933
1971

Source: Adapted and modified from Vilter (1978).

598

V I TA M I N C

disease that has caused the most suffering in recorded history (Carpenter, 1986). Scurvy was one of the earliest diseases known, with historical evidence of its existence in Egypt, Greece, and Rome. In the Middle
Ages scurvy was endemic in northern Europe during the late wintermonths and early spring, because at that time the foodstuffs that provided the chief source of vitamin C (green vegetables) had not been introduced. During this period scurvy was common among the seafaring
Vikings and the land-dwelling northern Europeans. Between 1556 and1857, 114 scurvy epidemics were reported in Europe, occurring during
winter when fruits and vegetables were not available. For hundreds ofyears it was known or suspected that a dietary factor would protect
against scurvy. Real progress was made toward identification of the dietary factor involved when in 1907 Holst and Frolich discovered thatguinea pigs could develop scurvy, thus providing an experimental animal.
World exploration and military operations were severely hampered
during the era prior to the seventeenth century by ravages of scurvy. On
the long sea voyages in explorations of the late 1400s and 1500s, scurvy
frequently occurred. Knowledge of the curative value of certain foods
had been known for a considerable time, and American Indians had
used an infusion of spruce or pine needles to prevent scurvy for at least
four centuries. As early as 1536, Jacques Cartier learned from these Indians that scurvy could be cured and prevented by consuming a drink
made from pine needles and bark. Previously, 107 cases of scurvy in 110
men resulted from Cartier’s expedition up the St. Lawrence River. Before
the relationship of scurvy to diet was found, there was a tendency to associate scurvy with venereal disease. Mercury was used as a treatment,with disastrous results.
In 1747 James Lind, a British fleet physician, carried out one of thefirst examples of a controlled clinical experiment, in which he showedthat patients consuming lemon juice recovered from scurvy, while others
failed to do so. Indeed, Lind was responsible for the relief of both scurvy
and typhus in the fleet and probably helped as much as Lord Nelson tobreak the power of Napoleon. By the eighteenth century, it was realizedthat fresh fruit and vegetables and lime or lemon juice would protect
sailors on long voyages. The term limey, colloquially applied to the English population, stems from lime juice given to sailors in the British
Navy. Lemon juice had become a routine part of the British Navy dietsby 1795. However, scurvy was an epidemic problem during the American Civil War, and it was not until 1895 that an antiscorbutic diet became official in the U.S. Army.
599

CHAPTER FIFTEENIn the nineteenth century, 104 country epidemics of scurvy were registered. Infantile scurvy (Barlow’s disease) was a severe problem in thelate 1800s and early 1900s; it came about from bottle-feeding, andbreast-fed babies were not affected. The problem arose because infantmilk was not pasteurized but rather sterilized by boiling, thus destroying vitamin C.
In 1928, Szent-Györgyi isolated a substance (which he called hexuronic acid) from orange juice, cabbage juice, and ox adrenal glands.
The isolated substance was acidic, with the formula C6H12O6, and wasstrongly reducing. The same year, King isolated an antiscorbutic substance in crystalline form from orange juice and demonstrated that it
cured scurvy; thus Szent-Györgyi tried his hexuronic acid, found it also
cured scurvy, and concluded it must be vitamin C. It was not readily accepted that it represented a vitamin because amounts required werelarge relative to amounts required of other vitamins (up to 100 mg). But
finally, in 1933, when hexuronic acid was synthesized by Richstein andshown to have the true activity of the natural product, ascorbic acid wasrecognized as a vitamin.

CHEMICAL STRUCTURE, PROPERTIES, AND
ANTAGONISTS
Vitamin C primarily occurs in two forms (Fig. 15.1), namely, the reduced ascorbic acid and the oxidized dehydroascorbic acid. The L-isomer of ascorbic acid has activity. Although most of the vitamin exists as
ascorbic acid, both forms are biologically active. In foods the reduced
form of vitamin C may reversibly oxidize to the dehydro form, with dehydroascorbic acid further oxidized to the inactive and irreversible compound of diketogulonic acid. This change takes place readily; thus vitamin C is very susceptible to destruction through oxidation, a change thatis accelerated by heat and light. Diketogulonic acid can be further oxidized to oxalic acid and L-threonic acid.
There are four stereoisomers of ascorbic acid; in addition to the Lisomer, only erythorbic acid (D-araborascorbic acid) has activity. Erythorbic acid has only 1/20 the activity of L-ascorbic acid and is often
used in the food industry for addition to meats or canning operations as
an antioxidant. This antioxidant property is used in the addition of thevitamin in canning of certain fruits (e.g., erythorbic acid) to prevent oxidation changes that cause darkening. Ascorbic acid is so readily oxidized to dehydroascorbic acid that other compounds may be protectedagainst oxidation. Reversible oxidation-reduction of ascorbic acid with600

V I TA M I N C

Fig. 15.1 Structures of vitamin C: L-ascorbic acid (reduced form) and dehydroascorbic acid (oxidized form).

dehydroascorbic acid is the most important chemical property of vitamin C and the basis for its known physiological activities and stabilities(Moser and Bendich, 1991). Vitamin C is the least stable, and therefore
most easily destroyed, of all vitamins.
Ascorbic acid is a white to yellow-tinged crystalline powder. It crystallizes out of water solution as square or oblong crystals and is slightly
soluble in acetone and lower alcohols. A 0.5% solution of ascorbic acidin water is strongly acid, with a pH of 3. The vitamin is more stable inan acid than an alkaline medium. It is not found in dry foods and ismarkedly destroyed by cooking, particularly when the pH is alkaline.
Cooking losses also result because of its solubility. A number of chemical substances—such as air pollutants, industrial toxins, heavy metals,
tobacco smoke, and several pharmacologically active compounds,
among them some antidepressants and diuretics—are antagonistic to vitamin C and can lead to increased requirements of the vitamin.
Crystalline ascorbic acid is relatively stable in air without moisture,and small concentrations of metal ions will accelerate destruction of
ascorbic acid. Various derivatives and analogs of vitamin C have been
prepared that have little if any antiscorbutic activity. Glycoascorbic acidacts as an antimetabolite for vitamin C, which is an ascorbic acid homolog, and contains an added CHOH group that has undergone optical
inversion.

ANALYTICAL PROCEDURES
Analysis of vitamin C includes biological, chemical, and physicalmethods. The biological method is specific for antiscorbutic activity
and, as such, can be accepted as the final standard of reference when it
is suspected that accuracy of chemical or physical procedures may be affected by presence of interfering substances. The biological test measures
601

CHAPTER FIFTEEN

total amount of vitamin C present, that is, in both the reduced form ofascorbic acid itself and the reversibly oxidized form of dehydroascorbic
acid. Applicability of the biological method may be limited only if potency of the test material is too low for it to be measured accurately. Ratscannot be used as test animals for vitamin C assay because of their ability to synthesize the vitamin, but guinea pigs have proved satisfactory.
Biological methods are based on prevention or cure of scurvy in guinea
pigs, in addition to dental histology, curative growth, and serum concentrations of alkaline phosphate associated with the deficiency. Procedures with guinea pigs require about 10 weeks for completion. Bioassays
based on microorganisms have not been developed because no organism
has been found that has an absolute requirement for L-ascorbic acid
(Moser and Bendich, 1991).
Biological analytical procedures have largely been replaced bychemical and physical methods, which provide precise, faster, and lessexpensive assays. Chemical and physical methods require precautions toprevent oxidation (e.g., homogenize under nitrogen and avoid copperand other metallic ions). Dye methods are widely used with the reagents,
2,6-dichlorophenolindophenol for reduced ascorbic acid and 2,4-dinitrophenylhydrazine for the oxidized form and for total ascorbic acid after oxidation. L-Ascorbic acid absorbs strongly in the ultraviolet, whichis the basis of spectrophotometric methods (Tono and Fujita, 1982).
Both gas-liquid chromatography and high-performance liquid chromatography (HPLC) have been developed for L-ascorbic acid determination (Rose and Nahrwold, 1982; Liau et al., 1993). The HPLC technology allows a rapid separation and detection of ascorbic acid as well
as of erythorbic acid.

METABOLISM
Vitamin C is absorbed in a manner similar to that of carbohydrates(monosaccharides). Intestinal absorption in vitamin C-dependent animals appears to require a Na+-dependent active transport system. It is
assumed that those species that are not prone to scurvy have an absorption mechanism by diffusion (Spencer et al., 1963). Ascorbic acid isreadily absorbed when quantities ingested are small, but limited intestinal absorption by active transport occurs when excessive amounts ofascorbic acid are ingested. However, uptake by passive diffusion occurs
at higher vitamin C intakes.
Bioavailability of vitamin C in foods is limited, but apparently 80 to

602

V I TA M I N C

90% appears to be absorbed (Kallner et al., 1977). The site of absorption in the guinea pig is in the duodenal and proximal small intestine,whereas the rat showed highest absorption in the ileum (Hornig et al.,1984). In humans, ascorbic acid is absorbed predominantly in the distal
portion of the small intestine and, to a lesser extent, in the mouth, stomach, and proximal intestine (Moser and Bendich, 1991).
In its metabolism, ascorbic acid is first converted to dehydroascorbate by a number of enzyme or nonenzymatic processes and is then reduced in cells (Rose et al., 1986). Vitamin C is transported in the plasmain association with the protein albumin. Absorbed vitamin C readilyequilibrates with the body pool of the vitamin.
Ascorbic acid is widely distributed throughout the tissues, both inanimals capable of synthesizing ascorbic acid as well as in those dependent on an adequate dietary amount of vitamin C. In experimental
animals, highest concentrations of vitamin C are found in the pituitary
and adrenal glands, with high levels also found in the liver, spleen, brain,
and pancreas. The vitamin tends to localize around healing wounds. Tissue levels are decreased by virtually all forms of stress, which also stimulates the biosynthesis of the vitamin in those animals capable of synthesis.
Humans receiving adequate intakes of vitamin C have a body poolof approximately 1.5 to 5 g of the vitamin, with 3 to 4% of the existing
body pool utilized daily. The major quantities of vitamin C are found in
the liver and muscles by virtue of their large masses. The half-life ofascorbic acid is inversely related to daily intake and is 13 to 40 days inhumans and 3 days in guinea pigs, which correlates with the longer timeneeded by a human to develop scurvy: 3 months for a human on a vitamin C-free diet compared with 3 weeks for the guinea pig.
Absorbed ascorbic acid is excreted in urine, sweat, and feces. Fecalloss is minimal, and even with large intakes in humans, only 6 to 10
mg/day is excreted by this route (Marks, 1975). Loss in sweat is also
probably low. In guinea pigs, rats, and rabbits, CO2 is the major excretory mechanism for vitamin C. Primates do not normally utilize the CO2
catabolic pathway, with the main loss occurring in the urine. However,
because of the limits on absorption, as dietary ascorbic acid increases
above 180 mg/day, it is broken down by intestinal bacteria, with 30%
or more recovered as carbon dioxide (Kallner et al., 1985). Urinary excretion of vitamin C depends on the body stores, intake, and renal function. Mechanism and mode of elimination are a function of glomerular
filtration rate of ascorbic acid and are dependent on plasma ascorbate603

CHAPTER FIFTEEN

concentration. Substantial quantities of L-ascorbic acid are excreted inurine after concentration in blood plasma exceeds its usual threshold ofapproximately 1.4 mg/100 ml.
Urine contains numerous metabolites of ascorbic acid, including dehydroascorbic acid, diketogulonic acid, ascorbate-2-sulfate, oxalate,methyl ascorbate, and 2-ketoascorbitol (Sauberlich, 1990). In humans,with physiological vitamin C doses of 60 to 100 mg/day, urinary oxalateis the major metabolite, with 30 to 50 mg/day being formed. But when
given in large doses, up to 10 g/day, urinary oxalate is increased by only
10 to 30 mg/day, and the vitamin is excreted largely unmetabolized inurine and feces (Moser and Bendich, 1991). Excretion of ascorbic acidin urine declines to undetectable levels with inadequate intakes of the vitamin or in the case of scurvy.

FUNCTIONS
Ascorbic acid has been found to be involved in a number of biochemical processes that involve donation of one or two electrons. Function of vitamin C is related to its reversible oxidation and reduction characteristics; however, the exact role of this vitamin in the living system is
not clearly known since a coenzyme form has not been reported. Nevertheless, vitamin C plays important roles in many biochemical reactions,such as mixed-function oxidation involving incorporation of oxygen into
the substrate. Biochemical and physiological functions of vitamin C havebeen reviewed (Chatterjee, 1978; Hornig et al., 1984; Sauberlich, 1990;
Moser and Bendich, 1991; Padh, 1991; Gershoff, 1993).

Collagen Synthesis
The most clearly established functional role for vitamin C involves
collagen biosynthesis. Collagens are the tough, fibrous, intercellular materials (proteins) that are principal components of skin and connective
tissue, the organic substances of bones and teeth, and the ground substances between cells. Impairment of collagen synthesis in vitamin C deficiency appears to be due to lowered ability to hydroxylate lysine andproline. Syntheses of collagens involve enzymatic hydroxylations of proline to form a stable extracellular matrix and of lysine for glycosylationand formation of cross-links in the fibers (Barnes and Kodicek, 1972).
Hydroxyproline residues contribute to the stiffness of the collagen triple
helix, and hydroxylysine residues bind (via their hydroxyl groups) carbohydrates and form intramolecular cross-links that give collagen structural integrity.
604

V I TA M I N C

In addition to the relationship of ascorbic acid to hydroxylase enzymes, Franceschi (1992) suggests that vitamin C is required for differentiation of mesenchyme (embryonic cells capable of developing into
connective tissue)-derived connective tissues such as muscle, cartilage,
and bone. It is proposed that the collagen matrix produced by ascorbic
acid-treated cells provides a permissive environment for tissue-specificgene expression. A common finding in all studies is that vitamin C canalter the expression of multiple genes as cells progress through specificdifferentiation programs (Ikeda et al., 1997).
The requirement for ascorbic acid is specific, probably protecting
the hydroxylase enzymes by oxidation of both the ferrous ions and thiolgroups present. It may be that ascorbic acid is not required for the hydroxylation reaction per se but is required to keep the enzyme-boundiron in the ferrous state. Hydroxyproline is found only in collagen(14%) and arises from hydroxylation of proline. In its absence, a nonfibrous collagen precursor is formed instead of fibrous collagen andwould result in scurvy.
Beneficial effects result from ascorbic acid in the synthesis of “repair” collagen. Alteration of basement membrane collagen synthesis andits integrity in mucosal epithelia during vitamin C restriction explains
the mechanism by which the capillary fragility is induced in scurvy andincreased incidences of periodontal disease under vitamin C deprivation(Chatterjee, 1978). Failure of wounds to heal and gum and bonechanges resulting from vitamin C undernutrition are direct consequencesof reduction of insoluble collagen fibers.

Antioxidant and Immunity Role
Free radicals can be extremely damaging to biological systems(Padh, 1991). Free radicals, including hydroxy, hypochorite, peroxy,
alkoxy, superoxide, hydrogen peroxide, and singlet oxygen are generated by auto-oxidation, radiation, or from activities of some oxidases,
dehydrogenases, and peroxidases. Also, phagocytic granuylocytes undergo respiratory burst to produce oxygen radicals to destroy the intracellular pathogens. However, these oxidative products can, in turn, damage healthy cells if they are not eliminated. Antioxidants serve tostabilize these highly reactive free radicals, thereby maintaining thestructural and functional integrity of cells (Chew, 1995). Therefore, antioxidants are very important to immune defense and health of humans
and animals.
Tissue defense mechanisms against free-radical damage generally include vitamin C, vitamin E, and β-carotene as the major vitamin an605

CHAPTER FIFTEEN

tioxidant sources. In addition, several metalloenzymes, including glutathione peroxidase (selenium), catalase (iron), and superoxide dismutase (copper, zinc, and manganese), are also critical in protecting the internal cellular constituents from oxidative damage. The dietary andtissue balance of all these nutrients are important in protecting tissue
against free-radical damage. Both in vitro and in vivo studies show thatthe antioxidant vitamins generally enhance different aspects of cellularand noncellular immunity. The antioxidant function of these vitamins
could, at least in part, enhance immunity by maintaining the functionaland structural integrity of important immune cells. A compromised immune system will affect human health and will result in reduced animal
production efficiency through increased susceptibility to diseases,
thereby leading to increased animal morbidity and mortality.
Vitamin C is the most important antioxidant in extracellular fluids(Stocker and Frei, 1991). Vitamin C can protect biomembranes against
lipid peroxidation damage by eliminating peroxyl radicals in the aqueous phase before the latter can initiate peroxidation (Frei et al., 1989;
Mukhopadhyay et al., 1995). In one study, vitamin C and E supplementation resulted in a 78% decrease in the susceptibility of lipoproteins tomononuclear cell-mediated oxidation (Rifici and Khachadurian, 1993).
Ascorbic acid is reported to have a stimulating effect on phagocytic
activity of leukocytes, on function of the reticuloendothelial system, andon formation of antibodies. Vitamin C can stimulate the production ofinterferons, the proteins that protect cells against viral attack (Siegel,1974). Some of the most controversial topics regarding the interactionsof vitamin C with immune functions have been the reduction of common cold symptoms and favorable responses to cancer treatment. In
guinea pigs, vitamin C was shown to be important in maintaining normal primary and secondary antibody responses and was important forneutrophil function (Anderson and Lukey, 1987). Ascorbic acid is very
high in phagocytic cells, with these cells using free radicals and otherhighly reactive oxygen containing molecules to help kill pathogens that
invade the body. In the process, however, cells and tissues may be damaged by these reactive species. Ascorbic acid helps to protect these cells
from oxidative damage.
One of the protective effects of vitamin C may partly be mediatedthrough its ability to reduce circulating glucocorticoids (Degkwitz,
1987). The suppressive effect of corticoids on neutrophil function in cattle is alleviated with vitamin C supplementation (Roth and Kaeberle,1985). In addition, ascorbate can regenerate the reduced form of α-tocopherol, perhaps accounting for observed sparing effects of these vita606

V I TA M I N C

mins (Jacob, 1995). In the process of sparing fatty acid oxidation, tocopherol is oxidized to the tocopheryl free radical. Ascorbic acid can donate an electron to the tocopheryl free radical, regenerating the reduced
antioxidant form of tocopherol.
Research findings in humans have suggested that “oxidative stress”
may be a causal factor in the etiology of diverse and important disordersof aging, such as cancer, cardiovascular disease, and cataract formation.
Cancers of the oral cavity, larynx, esophagus, stomach, colon, and rectum appear most related to low vitamin C intake compared to other cancer types (Gershoff, 1993). Several lines of evidence support a role foroxidized low-density lipoprotein (LDL) in the genesis of the atherosclerotic lesion. Vitamin C has been shown to inhibit LDL oxidation. Also,
there is evidence that vitamin C increases high-density lipoproteins(HDLs) and may lower total cholesterol. Ascorbic acid may also havebeneficial effects on blood pressure, and there are epidemiological studies associating subjects with cardiovascular disease with lower vitamin
C intakes than normal case controls (Simon, 1992). A number of reportssuggest that ascorbic acid will reduce the risk of cataracts (Gershoff,
1993; Jones and Hothersall, 1993).
The functional importance of vitamin C—other than the previously
mentioned roles in collagen synthesis and in immunity, and as an antioxidant—includes the following:
1. Because of the ease with which ascorbic acid can be oxidized and reversibly reduced, it is probable that it plays an important role in reactions
involving electron transfer in the cell. Almost all terminal oxidases in plant
and animal tissues are capable of directly or indirectly catalyzing the oxidation of L-ascorbic acid. Such enzymes include ascorbic acid oxidase, cytochrome oxidase, phenolase, and peroxidase. In addition, its oxidation isreadily induced under aerobic conditions by many metal ions, hemochromogens, and quinones.
2. Metabolic oxidation of certain amino acids, including tyrosine, occurs. Tyrosyluria is observed when high levels of tyrosine are being metabolized. Vitamin C appears to prevent the inhibition of the enzyme p-hydroxyphenylpyruvic acid oxidase by its substrate, p-hydroxyphenylpyruvic
acid, in the tyrosine metabolism sequence.
3. Ascorbic acid has a role in metal ion metabolism because of its reducing and chelating properties. The function of ascorbic acid is to provide
electrons to keep prosthetic metal ions in their reduced forms. This includes
cuprous ions in monooxygenases and ferrous ions in dioxygenases. Ascorbic acid can result in enhanced absorption of minerals from the diet and607

CHAPTER FIFTEEN

their mobilization and distribution throughout the body. Ascorbic acid promotes nonheme iron absorption from food (Olivares et al., 1997) and actsby reducing the ferric iron at the acid pH in the stomach and by forming
complexes with iron ions that stay in solution at alkaline conditions in the
duodenum. It appears to also function in the reduction and release of ferriciron from its tight linkage with plasma protein and its incorporation into
ferritin. There is a postabsorption role for ascorbate for cross-membranetransport of copper ions into cells (Harris and Percival, 1991). Ascorbic acidalso tends to alleviate toxic effects of transition metals in the body. Ascorbic acid, alone or in combination with chelators, lowers the concentrationof lead in the tissues of the body.
4. Carnitine is synthesized from lysine and methionine and is dependent
on two iron-containing hydroxylases for which ascorbic acid is a cofactor.
Vitamin C deficiency can reduce the formation of carnitine, which can result in accumulation of triglycerides in blood, and in the physical fatigue andlassitude associated with scurvy (Ha et al., 1994).
5. Interrelationships of vitamin C to B vitamins are known as tissue levels, and urinary excretion of vitamin C is affected in animals with deficiencies of thiamin, riboflavin, pantothenic acid, folacin, and biotin. Vitamin C
is active in changing the form of folacin to the tetrahydro derivative, a reduced form. It may also affect the ability of the body to store folacin. Whenvitamin C is deficient, utilization of folacin and vitamin B12 is impaired, resulting in anemia.
6. Vitamin C has a role in diabetes. In one study, the pancreas fromscorbutic guinea pigs contained 2.4 times more insulin than that from control guinea pigs, suggesting that the decreased insulin release from the scorbutic islets was not due to decreased insulin synthesis but due to abnormal
insulin secretion (Wells et al., 1995). From a different aspect of diabetes, excessive production of sorbitol from glucose is believed to play a role in thecausation of some of the complications of diabetes. It was reported that supplementation with vitamin C (100 mg/day) could correct the elevated sorbitol levels seen in diabetes (Cunningham et al., 1994). Vitamin C may exert its effect by inhibiting the activity of the enzyme aldose reductase.
7. Vitamin C has been demonstrated to be a natural inhibitor of nitrosamines, which are potent carcinogens. Action of ascorbate in preventing
formation of nitrosamines is reported to result from its direct reaction withnitrate.
8. Ascorbic acid is found in up to a 10-fold concentration in seminal
fluid (versus serum levels). Decreasing levels have caused nonspecific sperm
agglutination. It has been hypothesized that the vitamin may help to protect
sperm from effects of harmful oxidation. In a review of ascorbic acid and608

V I TA M I N C

fertility, Luck et al. (1995) suggested how three of ascorbic acid’s principal
functions—its promotion of collagen synthesis, its role in hormone production, and its ability to protect cells from free radicals—may explain its reproductive actions.
9. Other physiological effects of vitamin C include detoxification of histamine, metabolism of drugs (e.g., to detoxify), expression of acetylcholine
receptor, leukotriene biosynthesis, and a relationship to periodontal disease
and rheumatoid arthritis (Padh, 1991).

REQUIREMENTS
A wide variety of plant and animal species can synthesize vitamin Cfrom carbohydrate precursors, including glucose and galactose. The
missing step in the pathway of ascorbic acid biosynthesis in all vitamin
C-dependent species has been traced to inability to convert L-gulonolactone to 2-keto-L-gulonate, which is transformed by spontaneous isomerization into its tautomeric form, L-ascorbic acid. Vitamin C dietarydependent species, therefore, lack the enzyme L-gulonolactone oxidase.
Metabolic need for ascorbic acid is a general one among species, buta dietary need is limited to humans, subhuman primates, guinea pigs,
fruit-eating bats, some birds (including the red-vented bulbul and relatedPasseriformes species), insects, fish (such as coho salmon, rainbow
trout, and carp), and perhaps certain reptiles. Inherited ascorbic acid deficiency reported in pigs (Kristensen et al., 1986) and rats (Horio et al.,1985) was related to the lack of L-gulonolactone oxidase enzyme activity. Even for species that synthesize vitamin C, the synthesizing capacityof liver microsomal preparations was shown to vary strongly from animal to animal (Chatterjee, 1978), suggesting possible dietary need forthe vitamin for individuals within a species. Circumstances in which vitamin C deficiencies may occur in domestic animals are discussed in the
Deficiency and Supplementation sections.
In general, domestic animals such as poultry, ruminants, swine,
horses, dogs, and cats have the ability to biosynthesize ascorbic acidwithin their body, and hence there is no recommended requirement established by the National Research Council. However, Marks (1975)
proposed the following vitamin C requirements for poultry and swine,
per kilogram of diet: poultry, 50 to 60 mg; starting pigs, 300 mg; andfinishing pigs, 150 mg. Itze (1984) suggested 250 mg of ascorbic aciddaily for young calves. The ascorbic acid requirements of fish vary withspecies, size, environment, and health status of the animals. Rainbow
trout require 70 to 100 mg of L-ascorbic acid per kilogram of diet for609

CHAPTER FIFTEEN

normal growth and 500 mg/kg for wound repair (Halver, 1972). For
channel catfish, Li and Lovell (1985) determined a requirement of 30
mg/kg for maximum growth and 150 mg/kg for disease resistance. Likewise, increased vitamin C intakes are required for subjects exposed tocold or elevated temperatures and other acute stresses, including surgery
and trauma (Sauberlich, 1990). Others for whom dietary vitamin C requirement is increased include elderly individuals, alcoholics, and usersof oral contraceptives.
For humans, the RDA (1989) recommends 30 to 95 mg of vitamin
C depending on body weight and physiological function (Table 15.2).
The human requirement for vitamin C has been the subject of considerable debate, with diversity of opinion that seems to be irreconcilable.
This controversy is reflected in the wide range of recommended daily allowances established by different countries (20 to 200 mg). These values
are based on whether the allowance should prevent scurvy and permit amargin of safety, or whether more complete tissue saturation is preferable. Human scurvy can be prevented and cured with daily intake of 10
mg of vitamin C. This level of intake, however, permits little or no reserves. Approximately 18 to 25 mg/day keeps tissues half saturated.
Requirements are increased by pregnancy, lactation, thyrotoxicosis,
increased metabolism, or decreased absorption. Individuals may require
increased vitamin C as a result of stress or unfavorable environmental
situations. Cigarette smokers with vitamin C intakes comparable tothose of nonsmokers have serum vitamin C levels lower than those ofnonsmokers. Smoker requirements for ascorbic acid were estimated to
be increased by as much as 50%.
Research that previously established that the sole function of ascorbic acid was to prevent scurvy must be revised. There is evidence suggesting that ascorbic acid participates in extra “antiscorbutic” functions
for which a higher requirement than necessary to protect against overt■ Table 15.2

Recommended Vitamin C Daily Dietary
Allowances for Humans

Group

Amount (mg)

Under 1 year
1–10 years
11–14 years
15–51+ years
Pregnancy
Lactation
Source: RDA (1989).

610

30–35
40–45
50
60
70
90–95

V I TA M I N C

scurvy is needed. Pauling’s 1971 book stimulated widespread interest in
self-medication with megavitamin doses of ascorbic acid. There are reports in the literature concerning the beneficial effects of megadoses ofvitamin C on the common cold, resistance, and various diseases. Pauling(1971) suggested that for optimum health, daily intake of ascorbic acidfor an adult man should be 2.3 g, which could be increased to 9 to 10 g
in presence of some ailments. There is great individual variation in vitamin C requirements; some people in excellent health need 250 mg/day,most people need 4 to 5 g, and for many, 10 g is best. Ten grams is morethan 100 times the daily requirement suggested by the RDA (1989).
Intake of vitamin C far in excess of physiological requirements havebeen reported to have beneficial effects, including the following (Moser
and Bendich, 1991; Pauling, 1971; Ausman and Mayer, 1999):
1. Prevention and reduction of severity of the common cold.
2. Prevention of cancer and prolonged life of cancer patients.
3. Lowering of serum cholesterol and severity of atherosclerosis.
4. Increased wound repair and normal healing processes.
5. Increased immune response for prevention and treatment of infections.
6. Control of schizophrenia.
7. Inactivation of disease viruses.
8. Prevention of megaloblastic anemia of formula-fed infants.

The efficacy of pharmacological levels of the vitamin remains controversial because many of the claims are difficult to substantiate. Many
controlled studies will be required to establish which claims for megadoses of vitamin C are valid. In view of the controversy, some authorities suggest not taking large amounts of vitamin C without medical
counsel. However, in recent years, health benefits arising from using vitamin C as one of the antioxidant vitamins have become more accepted.

NATURAL SOURCES
The main sources of vitamin C are fruits and vegetables, but somefoods of animal origin contain more than traces of the vitamin (Table
15.3). Nearly 90% of vitamin C in the human diet is obtained fromfruits and vegetables as ascorbic and dehydroascorbic acids. Vitamin C
is present in relatively large amounts in fresh, canned, and frozen citrus
fruits and in smaller but important amounts in other fruits, tomatoes,
potatoes, and leafy vegetables (Nobile and Woodhill, 1981). Potatoes
611

CHAPTER FIFTEEN■ Table 15.3

Vitamin C Concentrations in Various Foods (mg/100 g, as-fed basis)

Vegetables
Asparagus, canned
Beans, runner
Brussels sprouts
Cabbage, redCarrots
Cauliflower, raw
Celery, raw
CornOats, whole
Onions, raw
ParsleyPeas, frozen
Peppers, raw
Potatoes, newRadishes
Rice
Rye, whole
SpinachWheat, whole

15
5
90
55
2–6
50–90
7
12
0
10
170
13
100
18
25
0
0
10–60
0

Fruit
Apples, unpeeled
Bananas

10–30
6–12

Blackberries
Cherries
Cranberries
Grapefruit
GuavasLemonsLimes
Melons (cantaloupe)
OlivesOrangesPeachesPineapple
Rose hips
Strawberries
Tangerines
Animal Products
Fish
Kidney, lamb
Kidney, pig
Liver, calf
Liver, pig
Milk, cow
Milk, human

20
5
12
34–45
300
80
250
25
0
40–60
7–14
25
1,000
40–90
30
5–30
9
11
13
15
1–2
3–6

Sources: Adapted from Nobile and Woodhill (1981) and Moser and Bendich (1991).

and cabbage are probably the most important sources of vitamin C for
the majority of the Western population, at least during winter (Moser
and Bendich, 1991). Vitamin C occurs in significant quantities in animalorgans such as liver and kidney, but in only small quantities in meat. In
untreated fruits and vegetables, content varies extremely in skin andpulp, and even between two leaves of the same vegetable or adjacent
plants of the same variety. Other factors affecting vitamin C content include variety, maturity, fertility, and season (Snehalatha Reddy and Lakshmi Kumari, 1988). Postharvest storage values vary with time, temperature, damage, and enzyme content (Zee, 1991). In Nigeria, significant
losses (between 21 and 83%) of ascorbic acid were observed after 4 and8 weeks storage of garri and cassava, respectively (Ukhun and Dibie,
1991).
Fresh tea leaves, some berries, guava, and rose hips are accumulators of ascorbic acid and consequently are rich sources. For practical
purposes, raw citrus fruits are good daily sources of ascorbic acid sinceappreciable amounts in other foods can be destroyed during processing.
Ascorbic acid in foods is easily destroyed by oxidation; therefore, undue
exposure to oxygen, copper, and iron, and prolonged cooking at high612

V I TA M I N C

temperatures in the presence of oxygen, should be avoided. However,quick heating methods can protect food vitamin C by inactivating plant
oxidases. Vitamin C is relatively stable to normal boiling, but losses aresubstantial with greater oxidation by steaming or pressure cooking. Often there is only a small vitamin C loss for foods during freezing or dehydration. However, in one report, frozen storage destroyed about 28%of the vitamin C content in black currants and 34% in strawberries(Hagg et al., 1995). No significant loss of vitamin C from microwave
heating of infant formula has been reported (Sigman-Grant et al., 1992).
Bioavailability of vitamin C for humans using a depletion-repletion
technique revealed that ascorbic acid from oranges, orange juice, andcooked broccoli is similar to that of synthetic ascorbic acid (Mangels etal., 1993). However, the bioavailability of ascorbic acid in raw broccoli
was 20% lower than the other sources. A study to simulate home conditions for vitamin C in orange juice showed that juice retained an average of 88% of the original ascorbic acid after 1 week and 67% after
2 weeks in opened containers stored at typical home refrigerator temperatures (Shaw and Moshonas, 1991).

DEFICIENCY
Effects of Deficiency
Under practical feeding situations, only humans, nonhuman primates, guinea pigs, and fish will develop vitamin C deficiency if diets arelacking in the vitamin. Farm livestock synthesize ascorbic acid from glucose in either the liver or the kidney, and vitamin C deficiency usually
does not occur in such animals. In the case of well-balanced nutrition,
their tissues receive endogenous ascorbate continuously, and the level inblood and tissues can only with difficulty be affected by exogenous vitamin C. However, with nutritionally unbalanced diets, relative vitamin
C deficiency may be induced in ascorbate-synthesizing animals as well
(Ginter, 1970). Low blood ascorbic acid can be caused by various typesof stress, including metabolic disorders, improper nutrition, insufficientvitamin A or β-carotene intake, and various infectious diseases. Under
such conditions, exogenous vitamin C can have a positive effect in ascorbic acid-synthesizing species.

Ruminants
All known ruminants can synthesize ascorbic acid; however, clinicalcases of scurvy in ruminants have been described. Death of cows and613

CHAPTER FIFTEEN

calves due to scurvy was characterized by changes in the oral cavity mucosa, muzzle, and skin, accompanied by weight loss and general unthriftiness (Cole et al., 1944; Duncan, 1944). In calves there was an extensive dermatosis accompanied by hair loss and thickening of skin inanimals receiving insufficient milk. Blood ascorbic acid was low, and thecondition was successfully treated with vitamin C injections. Scurvy andlowered blood ascorbic acid content in weaned calves were reported by
Martynjuk (1952). Studies of blood vitamin C concentrations in calvesreceiving the same diet have revealed great individual differences, withvariations related to genetic background (Palludan and Wegger, 1984).
Positive effects of ascorbic acid supplementation on milk yield andmilk quality were reported by Kuemyj (1955). Studies with bulls housed
differently showed a lowering of vitamin C stores as a result of cold
stress (Hidiroglou et al., 1977). Ruminants can actually be consideredmore prone to vitamin C deficiency because of impaired synthesis thanmonogastric animals since they cannot rely on exogenous supplies ofthis vitamin, which is rapidly destroyed by ruminal microflora (Cappa,
1958; Itze, 1984).
Hypovitaminosis C is most often observed in winter and spring andtends to reduce general resistance of the animals, causing infertility, highincidence of retained placenta, low viability of progeny, and several
other pathological conditions that can result in economic losses, particularly with calves (Soldatenkov and Suganova, 1966). Some data fromthe literature indicate that ascorbic acid-synthesizing capacity of calvesduring the first phases of life is insufficient to satisfy their requirements.
Some researchers even suggested that calves do not synthesize vitamin C
before 2 to 3 weeks after birth (Itze, 1984). However, Bouda et al.
(1980) established that adult levels of vitamin C were not present in thecalf until after 3 months of age. Evidence suggests that serum vitamin C
content is dependent on nutrition of the dam but also on age and healthof the calves. Yashin (1985) diagnosed marked scurvy clinical signs in
neonatal calves whose mothers had low ascorbic acid levels in the bloodand colostrum.
Jagos et al. (1977) found considerably lower plasma vitamin C content in calves with bronchopneumonia than in healthy animals. A relationship has been reported between hypovitaminosis C and skeletal
muscle pain and subcutaneous hemorrhages in calves (Pribyl, 1963).
Dobsinska et al. (1981) studied the relationship between ascorbemia
and body weight gains of calves in a large-capacity calf-house facility
and found a negative correlation between the two parameters in 2- to
22-week-old bull calves.
614

V I TA M I N C

Studies tend to show that low plasma ascorbic acid levels are likely
in animals experiencing gastrointestinal, respiratory, or other health ailments. Calves supplemented with vitamin C had lower incidence ofscouring (Cummins and Brunner, 1989). In calves with respiratory disease signs, Bouda et al. (1980) found significantly lower plasma ascorbic acid levels than in healthy calves. Podgornova and Donskova (1972)
found markedly reduced ascorbic acid concentrations in the organs of
cattle infested with tapeworm. Scott (1981) described a skin disorder incalves that had low plasma ascorbic acid levels and could be cured byadministration of vitamin C.
Calves from herds characterized by poor health status generally also
had reduced ascorbic acid status during the critical period from birth to
2 weeks of age. When calves were given doses of 1.25 to 2.5 g/day, infectious disease resistance appeared to increase, and respiratory diseases
were almost totally eliminated (Itze, 1984; Palludan and Wegger, 1984).
In conclusion, it is generally accepted that healthy adult ruminants
under normal dietary and environmental conditions are able to meettheir vitamin C requirements by body synthesis. However, young ruminants are susceptible to deficiency during the first few weeks of life, particularly when subjected to stress conditions, including cold, damp environments and disease, and/or if limited in colostrum consumption.

Swine
The signs of vitamin C deficiency in swine include weakness; fatigue; dyspnea; pain in the bones; and hemorrhages of the skin, musculature, adipose tissue, and certain organs (Zintzen, 1975). Swine nutritionists have generally formulated diets without vitamin C because theyoung pig can synthesize ascorbic acid within a week of birth, and both
sow colostrum and milk provide a plentiful source of the vitamin to thenursing pig (Wegger and Palludan, 1984). After it is weaned, the pig becomes dependent on its own tissue synthesis of the vitamin to meet its
metabolic needs. Although most farm species synthesize adequate quantities of ascorbic acid, some evidence suggests that the rate of synthesis
in swine may be inadequate during adverse environmental conditions,after a disease insult, and during other periods of stress. Mahan et al.
(1994) reported a growth response the initial 2 weeks postweaning but
not thereafter.
Swine researchers have indicated that under certain situations, pigs
may need supplemental vitamin C for maximum weight gain and feeduse (Mahan et al., 1966; Yen and Pond, 1981); however, nearly an equal
number of reports are negative (Brown et al., 1970; Yen and Pond,
615

CHAPTER FIFTEEN

1984, 1987; NRC, 1998). Reasons for this inconsistency may be that
unpredictable environmental and psychological stresses imposed onswine may increase requirements for ascorbic acid.
The level of available dietary energy is a major factor in determining the amount of ascorbic acid available to the pig (Brown et al., 1975;
Brown, 1984). Serum ascorbic acid concentrations as well as urinary
output are directly related to the level of energy in the diet. It was also
found that a minor stress such as individual penning will evoke a positive growth response from supplementary ascorbic acid, especially in animals fed a “low-energy” diet. Dietary energy is able to cause a shift inascorbic acid synthesis because of restrictions on amount of free glucose
available for this synthesis.
If a need for dietary vitamin C exists in swine, the newly weaned pig
would seem to be the class of swine most likely to be deficient. Sow’s
milk contains a high concentration of vitamin C at parturition, but thelevel drops dramatically with time toward weaning. For the baby pig,
the general consensus is that ascorbic acid blood level increases withcolostrum intake, drops at weaning, and slowly increases after 7 weeks
to the mature level (Wegger and Palludan, 1984).
A specific clinical leg-weakness syndrome in growing pigs manifests
itself mainly as crooked and/or deviated forelegs. These signs are indicated by contracted flexor tendons and weak joint ligaments that become apparent in pigs weighing 30 to 45 kg and seem to indicate impaired development in growing connective tissues (Nielsen and Vinther,1984). Vitamin C administered to boars during the growing period from39 to 105 kg body weight resulted in straightness of front legs compared
to controls (Cleveland et al., 1987).
Ivos et al. (1971) reported an inverse relationship between ambient
temperature and conception rate in sows. Additionally, these authors reported that average conception rate in sows increased when boars were
supplemented with either 1 or 2 g daily of ascorbic acid compared tocontrols. Lin et al. (1985) observed increased sperm concentration per
ejaculate in heat-stressed working boars that received 300 mg of ascorbic acid per day compared to unsupplemented boars. Boars that received
the supplemental ascorbic acid also had fewer abnormal sperm cells per
ejaculate.
Handling practices at weaning (especially early weaning)—which
are generally considered to be stressful and include transport and mixing with unfamiliar pigs—have been shown to deplete body ascorbate.
In view of decreased plasma vitamin C concentration and dramaticchanges in nutritional, social, and other environmental factors associ616

V I TA M I N C

ated with weaning, it was suggested that beneficial response from supplemental vitamin C with weanling pigs may be related to suppressionof postweaning subclinical disease (Yen and Pond, 1981). In a study
with growing pigs between the ages of 4 and 7 weeks, Park and Harrison (1990) reported an improvement in nursing pig performance (6%improvement in daily gain, 5% improvement in gain/feed) resulting
from vitamin C supplementation in tap drinking water.
Spontaneous scurvy as a result of a genetic defect was observed in aswine production herd among 2- to 3-week-old piglets (Jensen andBasse, 1984). Closer observation revealed that all pigs were from thesame boar. Analysis of their blood and tissues revealed only a very small
concentration of vitamin C. The 3:1 ratio between normal and affected
pigs was characteristic of simple autosomal recessive inheritance in matings between nonaffected carriers. Liver microsomes were shown to beincapable of synthesizing ascorbic acid in vitro even with L-gulonolactone as substrate (Jensen and Basse, 1984).
The influence of maternal vitamin C deficiency on fetal development
was studied in swine with a hereditary lack of ability to synthesize ascorbic acid (Wegger and Palludan, 1994). Severe pathological changes wereseen in the uterus and fetuses. Characteristic findings were hemorrhages
and hematomas in both fetal and maternal placenta, and general edema
and subcutaneous hemorrhages in the fetuses. Ossification of the fetal
skeleton was severely deranged.

Poultry
Like swine, poultry are able to synthesize vitamin C, and thus it isassumed they do not require dietary sources of the vitamin. However, in
newly hatched poultry, there is a slow rate of ascorbate synthesis, andthis, combined with encountered stress, increases probability of vitamin
C deficiency. The chick is subject to considerable stress conditions, suchas rapid growth, exposure to hot or cold temperatures, starvation, vaccination, and disease conditions such as coccidiosis. Pardue andWilliams (1990) reported that plasma ascorbic acid levels in poults weredepressed significantly by cold stress, beak trimming, and injection at 1
and 14 days of age. Supplemental vitamin C (150 ppm) enhanced performance of broiler chicks exposed to multiple concurrent environmental stressors (McKee and Harrison, 1995).
For both stressed mature and newly hatched poultry, several reports
have documented a beneficial effect of supplementing the feed withascorbic acid on growth rate, egg production, eggshell strength andthickness, fertility and spermatozoa production, counteracting unfavor617

CHAPTER FIFTEEN

able climate and housing conditions, and intoxication or disease (McDonald et al., 1981; Pardue, 1987). On the contrary, many researchers
have found no beneficial effect of vitamin C supplementation under any
conditions.
For heat-stressed chickens, supplemental vitamin C provided definite improvements in egg production, eggshell strength, and interior egg
quality (El-Boushy and Van Albada, 1970; Cheng et al., 1988). Peebles
and Brake (1985) also reported that supplemental ascorbic acid holds
promise for increased production during high environmental temperatures or for nutritionally marginal diets. When ascorbic acid was used at
levels of 100 ppm or less for commercial layers, there was improvement
in livability, egg production, and eggshell quality. Perek and Kendler(1963) carried out experiments in the Jordan Valley, where hens were
subjected to hot temperatures, and reported increases in egg productionof 23 and 11.2% in two experiments in which the birds were given supplemental ascorbic acid. They also reported increased egg weights, decreased culls and mortality, and no shell quality differences. Other researchers were not able to confirm the positive effects of ascorbic acidsupplementation.
Male reproduction is favored by vitamin C supplementation. Monsi
and Onitchi (1991) supplemented the feed of heat-stressed broiler breeders with 0, 125, 250, or 500 ppm of ascorbic acid. Semen volume, total
sperm per ejaculate, and motile sperm per ejaculate were significantly increased due to the addition of ascorbic acid. Semen volume and sperm
concentration of turkey toms were found to be increased by 28% by the
supplementation of 150 ppm of ascorbic acid to the breeder ration (Dobrescu, 1987). When Noll (1993) supplemented the feed of male breeder
turkeys with 200 ppm of ascorbic acid for 8 weeks, ascorbic acid supplementation increased semen volume 16% and sperm concentration
18%.
Vitamin C is necessary for bone development and eggshell quality.
Supplementing ascorbic acid to molted laying hens was beneficial to egg
production and eggshell quality (Zapata and Gernat, 1995). Orban et al.
(1993) reported that large doses of ascorbic acid (2,000 ppm) in the dietinfluenced calcium metabolism, affecting bone and eggshell mineralization in chickens. Vitamin C is a necessary cofactor for the bioconversionof vitamin D3 to its active form, 1,25(OH)2D3. Weiser et al. (1990) reported that 100 ppm of ascorbic acid in the diet of chicks increased
plasma concentrations of 1,25(OH)2D3, which led to elevated activitiesof duodenal calcium-binding protein and greater weights and breaking

618

V I TA M I N C

strength of bones. It is possible that the many cases of “field rickets” inpoults may be due to stress-induced deficiency of vitamin C.
Njoku (1986) concluded that during periods of heat stress in thetropics, dietary supplementation of broiler diets with 200 ppm of ascorbic acid was necessary and economically advantageous as body weight
and feed; gain responses were improved. Other researchers have notbeen able to confirm the positive effects of vitamin C supplementation.
In a study supplementing 2,600 ppm of ascorbic acid, egg production,
eggshell thickness, egg weight, and mortality were not affected, but interior quality was improved (Nockels, 1984).
Pardue et al. (1985) were unable to find significant vitamin C effectsof heat stress in broiler performance, except that heat-associated mortality was markedly reduced in supplemented females. When acute heatstress was imposed on broilers, vitamin C reduced adrenal corticosteroid
concentration in the plasma of birds associated with the stress. Apparently, high levels of ascorbic acid in the adrenal gland regulates glucocorticoid synthesis, thus limiting some of the deleterious responses associated with stress and delaying the depletion of steroid hormone
precursors.
Disease conditions have been found to affect vitamin C metabolism
in poultry. When chicks were infected with fowl typhoid, their plasma
vitamin C concentrations were reduced (Hill and Garren, 1958). The vitamin C concentrations in plasma and tissue were also reduced in chicks
infected with intestinal coccidiosis (Kechik and Sykes, 1979). Dietary
ascorbate was shown to prevent this and contributed to intestinal repair.
In addition to performance, evidence also suggests an association
between vitamin C and the animal’s ability to tolerate or resist bacterial
infection. In early work, chickens infected with fowl typhoid had reduced blood ascorbic acid, and the administration of vitamin C at 1,000
ppm in feed resulted in reduced early mortality from typhoid infection(Satterfield et al., 1940). Chickens fed a diet containing supplemental
ascorbic acid showed increased resistance to a combined Newcastle disease virus–Mycoplasma gallissepticum infection and to a secondary E.
coli infection, as well as to a primary Escherichia coli challenge infection(Takahashi et al., 1991).
Leghorn-type chickens aged 6 weeks, supplemented with 330 ppm
of vitamin C and exposed to air-sac challenge with E. coli, had a 19%
incidence of E. coli infection versus 76% in controls (Gross et al., 1988).
The authors hypothesized that the response to vitamin C may be attributed to the vitamin increasing the synthesis of superoxide anion that619

CHAPTER FIFTEEN

kills phagocytized bacteria. Dietary level of ascorbic acid appears to beimportant since too little vitamin C results in too little superoxide anion
production, and excessive vitamin C may result in reduction of superoxide anion in the phagocytic cells.
Since ascorbic acid can be synthesized at the tissue level by domestic fowl, it has been held by many nutritionists that exogenous supplementation of vitamin C to poultry would be senseless. Over the past several decades, the relationship between stress and vitamin C in poultry
was recognized; however, research data have been inconsistent and conflicting, making it difficult to establish requirements for this nutrient under all conditions. Some data suggest that supplementation with vitamin
C should be considered as a management alternative to prevent vitamin
C deficiency when poultry are stressed (Quarles and Adrian, 1989;
Quarles et al., 1989).

HorsesHorses, like other farm species, synthesize vitamin C, but stress situations such as bacterial and viral infection (e.g., influenza, rhinopneumonia) have been reported to lower vitamin C serum levels (Jaeschke,1984). This is further associated with blood parameters, indicating delayed and/or disturbed collagen metabolism in young horses. Studies
have shown that horses of all ages that suddenly show poor performance
often have a reduced ascorbic acid serum level (Jaeschke, 1984). Performance of these horses improved after intravenous administration of
ascorbic acid. Despite early suggestions that supplementary ascorbic
acid (vitamin C) improved the sperm quality of stallions and the breeding performance of mares (Davis and Cole, 1943), other workers havebeen unable to repeat those results.

Other Animal Species
DOGS AND CATS
The dog is able to synthesize vitamin C. Naismith (1958) showedthat this synthetic ability is present in puppies during the first weeks ofpostnatal life. Chatterjee et al. (1975) demonstrated that dogs synthesized ascorbic acid in the liver at an hourly rate of 5 μg/mg of protein.
However, this rate is low compared with the hourly rate in other mammals, such as cows (68 μg/mg of protein), rats (39 μg/mg of protein),and rabbits (23 μg/mg of protein).
There is controversy about the therapeutic use of ascorbic acid incanine diseases. Vitamin C has been used in treatment of canine viral in620

V I TA M I N C

fections. Some clinicians stated that 1,000 to 2,500 mg of ascorbic acidgiven intravenously once daily for at least 3 days was beneficial in thetreatment of canine distemper (Belfield, 1967; Leveque, 1969).
Vitamin C deficiency also has been reported to be associated withcanine hypertrophic osteodystrophy (Grondalen, 1976). Meier et al.
(1957) found that dogs with hyperbaric oxygen drenching had low
plasma ascorbic acid concentration, and large doses of vitamin C (100to 200 mg) given orally or intramuscularly enhanced healing. Vitamin C
supplementation of 50 to 100 mg/day may be beneficial in combating
arthritis in older dogs (Lewis and Morris, 1983).
Some trials failed to demonstrate a need for dietary ascorbic acid incats (Carvalho da Silva, 1950). Successful growth and reproduction areroutinely obtained with commercial and purified (NRC, 1986) diets
containing no supplemental ascorbic acid.
As in the dog, vitamin C synthesis in the cat is lower than in otherspecies, including the cow, sheep, rat, and rabbit (Rucker et al., 1980).
Pietronigro et al. (1983) reported that central nervous system function
following spinal cord injury in the cat is associated with large losses ofascorbic acid from the region of the injury. Treatment of injury with two
drugs (naloxone or methyl-prednisolone) preserved neurologic function
and prevented ascorbate loss.

FISH
Deficiency of vitamin C in most aquaculture species results in poor
formation of connective tissue. Intensively fed, caged channel catfish
grew slowly and exhibited scoliosis and lordosis (Fig. 15.2), brokenback syndrome, and elevated mortality from bacterial infection while receiving an ascorbic acid-deficient diet (Lovell, 1973). Other signs ofascorbic acid deficiency in channel catfish are internal and external hemorrhage, fin erosion, dark skin color, and reduced formation of bone collagen. Similar structural deformities such as scoliosis and lordosis due tovitamin C deficiency have been observed in Indian major carp, common
carp, roach, blue tilapia, Nile tilapia, and yellowtail (NRC, 1993). Vitamin C-deficient salmon and trout exhibited structural deformities (scoliosis, lordosis, and abnormal support cartilage of the eye, gill, and fins)and internal hemorrhaging, usually preceded by nonspecific signs suchas anorexia and lethargy (NRC, 1993).
Japanese eels fed a vitamin C-deficient diet had reduced growth after 10 weeks and hemorrhage in the head and fins after 14 weeks (Araiet al., 1972). Vitamin C-deficient shrimp exhibited black death syndrome, a condition characterized by melanized lesions in connective tis621

CHAPTER FIFTEEN

A

B

Fig. 15.2 Vitamin C deficiency in catfish. (A) Fingerling channel catfish fed adiet devoid of vitamin C for 8 weeks. Note scoliosis and lordosis. (B) Channel
catfish from commercial cage culture where the regular diet was devoid of vitamin C. Fish at left shows lateral curvature of the spine (scoliosis); fish atright shows vertical curvature (lordosis) and a vertical depigmented band at
the point of spinal injury, which is characteristic. (Courtesy of R.T. Lovell andAuburn University, Alabama.)
622

V I TA M I N C

sue under the exoskeleton as well as on the gills, abdomen, and gut (Magarelli et al., 1979).
The immune system and reproduction are dependent on vitamin C.
Vitamin C appears to protect phagocytic cells and surrounding tissues
from oxidative damage. An increased immune response due to high concentrations of vitamin C supplementation has been demonstrated inchannel catfish (Li and Lovell, 1985) and rainbow trout (Navarre andHalver, 1989). Reduced reproductive performance has been reported intilapia and rainbow trout (NRC, 1993).

FOXES AND MINK
Helgebostad (1984) indicated that the fox and mink are able to synthesize sufficient vitamin C.

LABORATORY ANIMALS
Because of body synthesis, the rat, mouse, and hamster do not require dietary vitamin C, and classic studies on scurvy have been conducted with the guinea pig (NRC, 1995). Early signs of vitamin C deficiency in guinea pigs are reduced feed intake and weight loss, followedby anemia and widespread hemorrhages. Additional signs include enlarged costochondral junction, disturbed epiphyseal growth centers oflong bones, bone demineralization, altered dentine, and gingivitis (NRC,
1995; Kip et al., 1996). Vitamin C deficiency in guinea pigs also resultsin hypertriglyceridemia, hypercholesterolemia, and decreased vitamin E
concentrations in liver and lungs.
A number of studies using guinea pigs as a laboratory animal requiring vitamin C are examining new functional roles of the vitamin. As
examples, ascorbic acid has been shown to prevent lipid peroxidationand prevent damage of proteins in guinea pig extrahepatic tissue microsomes (Mukhopadhyay et al., 1995), and ascorbic acid was found to be
essential for the release of insulin from scorbutic guinea pig pancreatic
islets (Wells et al., 1995).

NONHUMAN PRIMATES
Unless dietary vitamin C content is adequate, scurvy is apt to occur
in all species of nonhuman primates. In fact, along with vitamin D3 deficiency in New World monkeys, scurvy is the most frequently diagnosed
specific nutrient deficiency in captive primates (NRC, 1978). Lesions ofvitamin C deficiency states include gingival hemorrhage, loose teeth,
subperiosteal hemorrhage, normocytic normochromic anemia, and epiphyseal fractures (Banerjee and Bal, 1959).
623

CHAPTER FIFTEENRABBITSThe rabbit does not require dietary vitamin C (NRC, 1977). Harris
et al. (1956) demonstrated that young rabbits kept for periods as longas 25 weeks on vitamin C-free diets gained weight normally and continued to excrete considerable amounts of ascorbic acid in their urine.

HumansIn humans, gross vitamin C deficiency results in scurvy, a diseasecharacterized by multiple hemorrhages (Fig. 15.3). In adults, manifest
scurvy is often preceded by lassitude, fatigue, anorexia, muscle pain, andgreater susceptibility to infection and stress. Scurvy is characterized byanemia and alteration of protein metabolism; weakening of collagenous
structures in bone, cartilage, teeth, and connective tissue; swollen, bleeding gums, with loss of teeth; fatigue and lethargy; rheumatic pain in thelegs; degeneration of muscles; massive “sheet” hematomas in the thighs;
and skin lesions. Bleeding gums gingivitis and loosening of the teeth areusually the earliest objective signs (Marks, 1975; Vilter, 1978; Moser
and Bendich, 1991). Structural defects characteristic of scurvy include
the following:
1. Bones and cartilage—the cartilage cells cease to form matrix.
2. Teeth—odontoblasts, predentine, dentine, and enamel are notformed; interrelated with bone and collagen formation.
3. Muscles—atrophy and necrosis occur, with calcium deposits.
4. Connective tissues—collagen fibers are not formed by fibroblasts.
5. Capillaries and vascular system—capillary walls are fragile.
6. Blood—hemorrhage causes anemia because red blood cells are rapidly destroyed.
7. Liver and other organs—liver atrophies and is infiltrated with fat;bile secretion is impaired; kidney atrophies; and spleen enlarges.
8. Reproductive organs—degeneration of ovaries or germinal epithelium of testes occurs but may be nonspecific.
9. Other endocrines—thyroid shows hyperemia, hypersecretion, and irregularity of structure; adrenals are abnormal.

Scurvy may be fatal, particularly in infants and otherwise debilitatedadults. Well-defined scurvy is not common in more developed countries,
and occurs chiefly in infants fed diets deficient in ascorbic acid. Scurvy
is an epidemic disease in underdeveloped areas of the world, where
poverty, wars, and great migrations produce situations of malnutrition.
624

V I TA M I N C

ABFig. 15.3 Scurvy in a vitamin C-deficient human. (A) Severe scurvy, withswelling, bleeding, and receding gums. (B) Capillary fragility, with small hemorrhagic areas (ecchymoses). (Courtesy of Howerde E. Sauberlich, Universityof Alabama.)

Dietary levels were found to be deficient for 41 to 48% of children in aregion of Russia (Krasnopevtsev et al., 1993) and for 30 to 41% of
adults in Warsaw, Poland (Pardo et al., 1991). In urban and rural regions of Turkey, 43% of school-age children had deficient plasma vitamin C concentrations.
Infantile scurvy (Barlow’s disease), which is usually due to lack ofvitamin C in artificial foods, generally occurs between the ages of 6 and625

CHAPTER FIFTEEN

18 months. In the United States in the late 1800s and early 1900s, many
children died from infantile scurvy due to the practice of boiling or sterilizing milk, which completely destroyed the vitamin C. As a rule, infantile scurvy is first noticed when the infant cries on being handled, isirritable, and loses appetite and weight. Tenderness of extremities andpain on movement are almost invariably present (Marks, 1975). Manifestations of scurvy appear insidiously, usually after 5 to 6 months of severe deprivation of vitamin C.
Scurvy is a rare disease in developed countries and appears sporadically in elderly individuals who live alone, in smokers, in alcoholics, ininfants with prolonged artificial feeding without a complementary diet,and in a miscellaneous group of patients with unusual diets, some ofwhom have psychiatric problems (Moser and Bendich, 1991; GomezCarrasco et al., 1994). Other risk groups include diabetics, rheumatoid
arthritis patients, oral contraceptive users, and individuals exposed topetrochemicals in the workplace (Moser and Bendich, 1991).
Vitamin C and the other antioxidant vitamins (vitamin E and βcarotene) convert highly active radicals to less active species and thereby
reduce certain disease incidence. A large body of evidence has been accumulated, including the results of recently published epidemiologicalstudies that consistently report an association of dietary intake of vitamins C and E and of β-carotene and a risk reduction for coronary heartdisease and cancer (Weber, 1994). Vitamin C alone has been related todramatic reduction of coronary heart disease risk factors (Kranowski,
1991; Jacques, 1992; Singh et al., 1995; Duell, 1996; Toohey et al.,
1996; Jacob, 1998) and cancer (Block, 1991; Gershoff, 1993; Kune etal., 1993). In addition to reduction of cancer and heart disease, vitamin
C is reported beneficial in eliminating cataracts, reducing complicationsof diabetes, maintaining normal sperm, lowering hypertension, and reducing infectious disease by maintaining a normal immune response (seeFunctions).

Assessment of Status
Completely satisfactory and reliable procedures to assess vitamin C
nutritional status have not been developed because of limited knowledge
concerning the vitamin’s metabolic functions. However, information
concerning adequacy has been determined by an analysis of vitamin C
concentrations in serum, leukocytes, whole blood, or urine. Leukocyte
vitamin C concentrations provide information concerning body stores ofascorbic acid (Turnbull et al., 1981). Precautions need to be taken to626

V I TA M I N C

protect the vitamin in solution, and to select an assay that measures thevitamin itself and not other substances present.
Methods of biochemical detection of deficiency include the following:
1. Tissue content—28 mg/100 g of liver is the saturation level for humans and guinea pigs; lower concentrations will reveal reduced intake.
2. Leukocytes—ascorbate concentration in leukocytes with adequate diets is about 25 mg/100 ml, with less than 20 mg/100 ml in deficiencies.
3. Serum—the normal range of ascorbate (0.5 to 2.2 mg/100 ml) is often too variable to permit reliable estimation of deficiencies.
4. Urinary excretion—the load test and saturation test generally indicate immediate past intake and not overall nutritional status; nevertheless,amount of a given dose that is excreted is indicative of the tissue stores.
Amount of ascorbate excreted during 3 hours following a 100-mg dose will
be 50% for a normal, saturated person; 15% for a depleted person; and 5%
for a scorbutic patient.
5. Other tests include (a) intradermal test, which involves rate of decoloration of 2,6-dichlorophenolindophenol injected intradermally; (b) decolorization of dye on the tongue; and (c) serum alkaline phosphatase, excretion of tyrosine metabolism products, and urinary creatine, which are notspecific.

SUPPLEMENTATION
Supplementation with vitamin C is not normally recommended forcommon livestock species (ruminants, poultry, swine, and horses) undernormal management and feeding regimens. As previously mentioned,stress conditions affect vitamin C synthesis, and supplementation considerations must take this into account. Kolb (1984) summarized various types of stress that apparently increase demands while reducing animals’ capability to synthesize vitamin C, as follows:
1. Dietary conditions—deficiencies of energy, protein, vitamin E, selenium, iron, etc.
2. Production or performance stress—high production or performance(e.g., rapid growth rates, high milk production, racehorse running).
3. Transportation, animal handling, and new environmental locationstress—animals being driven or transported to market, animals placed in
new surroundings (e.g., weaned pigs from different litters placed together),627

CHAPTER FIFTEEN

and stressful management practices (e.g., castration, vaccination).
4. Temperature—high ambient temperature or cold trauma.
5. Diseases and parasites—fever and infection reduce blood ascorbic
acid, while parasites, particularly of the liver, disturb ascorbic acid synthesis and increase requirements for the vitamin.

The stress associated with confinement calf housing decreased immune response to a specific antigen and decreased plasma ascorbate
concentrations (Cummins and Brunner, 1991). The health of young
calves supplemented with vitamin C improved, as reflected by decreased
navel infections, peritonitis, pneumonia, enteritis, respiratory disease,scouring, and mortality (Blair and Cummins, 1984; Itze, 1984; Cummins and Brunner, 1989).
During the first weeks of life, the calf’s requirement for ascorbic acidmust be covered by colostrum and milk concentrations and from the inborn storages of the vitamin. One calf study in which reared calves weredenied colostrum found that all but one of the experimental animals
died of umbilical infections and peritonitis (Palludan and Wegger, 1984).
That survivor had a high content of ascorbic acid in its blood at birth.
Further investigations showed positive results from supplementation of
calves with ascorbic acid. Calves fed 1.75 g/day of vitamin C that werecolostrum-deprived had lower clinical scores for diarrhea, and at 14days of age, plasma IgG concentrations were higher than those of controls not fed ascorbate (Cummins and Brunner, 1989).
It is important to note that reserves of ascorbic acid are high at birth
but decline rapidly afterward unless exogenous supply is furnished until
synthesis can handle the load (Itzeova, 1984; Palludan and Wegger,
1984). Plasma ascorbic acid level in calves fed fresh colostrum twice a
day and frozen colostrum of the first milking once a day was high, withindications that the decrease in ascorbic acid content usually seen after
birth can be avoided by this practice (Itzeova, 1984). Itze (1984) recommended supplementation with vitamin C for calves reared on milk
diets. The initial days of calf rearing are critical, for the calf must adapt
itself to a new environment, feeding practice, and housing at an age
when its resistance is minimal. Of various programs tested, the author
recommends daily oral supplementation with 2.5 g of ascorbic acid in
combination with parenteral application of 500 mg of ascorbic acid in
two doses immediately after moving animals into their new rearing facilities. Lehocky (1981), in his work with dairy calf supplementation,found that only 50% of a calf’s requirement for vitamin C is covered
when feeding various milk replacers.
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Various studies have demonstrated beneficial effects of low doses of
50 to 100 mg of ascorbic acid per kilogram to diets of broilers or laying
hens exposed to heat stress (Kolb, 1984). Njoku (1984) reported that
200 mg of ascorbic acid per kilogram fed to broilers helped alleviate
heat stress. Eggshell thickness increased for hens (El-Boushy et al.,1968), while livability, weight gain, and immune response improved in
broilers (Pardue and Thaxton, 1982) when heat-stressed birds received
supplemental vitamin C. Peebles and Brake (1985) fed vitamin C tobroiler breeders throughout a complete production cycle. They foundfertility improved at dietary levels of 50 mg/kg, with improvement in
hatch of fertile eggs due to a decrease in early embryonic mortality.
There was no further benefit at 100 mg/kg. In turkeys, toms given 150
mg/kg increased semen volume by 31% per ejaculate. It is thought thatvitamin C stimulates testicular activity by its involvement in the synthesis of steroid hormones (Dobrescu, 1987).
Since vitamin C can be synthesized at the tissue level by swine, it has
been held by many nutritionists that supplemental vitamin C to swine
would be senseless. Data suggest that supplementation with ascorbic
acid should be considered as a management alternative to prevent vitamin C deficiencies when swine are stressed.
The literature reports concerning efficacy of supplementation ofswine diets with ascorbic acid is conflicting. Young pigs seem to be more
likely than adults to respond to supplementation, much the same as wasreported for ruminants. Perhaps the inconsistency of results is due to uncontrolled stress or genetic differences (Brown, 1984).
Early weaning (0 to 3 weeks) of piglets has been shown to decrease
ascorbic acid levels in liver, and tests seem to indicate that maximal synthesizing capacity is not developed until about 8 weeks, thus indicating
a possible advantage in supplementing milk replacer products with vitamin C (Wegger and Palludan, 1984). Sandholm et al. (1979) reported
that umbilical hemorrhages (Fig. 15.4) occurring in piglets immediately
after birth can be prevented by supplementing the sows’ feed with 1 g ofascorbic acid per day during the last week of gestation.
The intensive selection that has taken place for several decades inthe swine industry may have altered the enzymatic constitution of animals so that ability to synthesize vitamin C has changed. Furthermore,
modern intensive production systems and continuous demand for higher
productivity may have increased the requirement of ascorbic acid forswine. Feeding practice in pig production has also changed; the tendency
is to use more processed feedstuffs that, practically speaking, contain noascorbic acid.
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CHAPTER FIFTEEN

Fig. 15.4 Navel bleeding syndrome. Umbilical cords of a bleeding
piglet (left) and a normal piglet (right), age
10 hours. Navel bleeding has been prevented
by preparturient administration of ascorbic
acid. (Courtesy ofMarkus Sandholm,
College of Veterinary
Medicine, Helsinki,
Finland.)

Therapeutic, as distinguished from nutritional, use of ascorbic acidhas been useful in treating infectious diseases of the horse and dog(Kolb, 1984). The importance of ascorbic acid for defense against infections and for phagocytosis has been stressed. Beneficial effects of supplemental vitamin C have been reported for puppies with disorders inbone development accompanied by painful swelling of joints when thevitamin is deficient (Kolb, 1984).
L-Ascorbic acid is the most important of the several compounds thathave vitamin C activity. Ascorbic acid is commercially available as
100% crystalline, 50% fat-coated, and 97.5% ethylcellulose-coatedproducts and their dilutions. The more soluble sodium salt of ascorbic
acid (sodium ascorbate) is also commercially available. Various derivatives of ascorbic acid, which are more stable than the parent compound,have been shown to provide antiscorbutic activity. These include Lascorbate-2-sulfate, L-ascorbyl-2-monophosphate, magnesium-L-ascor630

V I TA M I N C

byl-2-phosphate, and L-ascorbyl-2-polyphosphate. When providing supplemental ascorbic acid, it is advisable to use a stabilized form, and coating ascorbic acid crystals with ethylcellulose is a suitable stabilization
method. In storage experiments, ascorbic acid protected in this manner
was found to be four times more stable than untreated ascorbic acidcrystals (Kolb, 1984).
Adams (1982) reported that coated (ethylcellulose) ascorbic acidshowed a higher retention after processing than the crystalline form (84versus 48%). Retention of ascorbic acid in mash feed was fairly good,
but with elevated storage time and temperature, stability was poor in
crumbled feeds. Although retention of vitamin C activity in feed containing the ethylcellulose-coated product was low, it was 19 to 32% better than that of the crystalline form. Cows dosed orally with vitamin C
coated with ethylcellulose had higher plasma ascorbic acid concentrations than did cows that received vitamin C in the form of fine powder(Hidiroglou, 1999).
Reports have evaluated polyphosphorylated L-ascorbic acid in fish
(Chen and Chang, 1994; Matusiewicz et al., 1995), and found it to be
more stable against oxidation and extrusion. Approximately 50% of the
supplemental ascorbic acid is destroyed during the manufacture of extruded catfish feeds (Lovell and Lim, 1978), and excess ascorbic acid is
added to commercial formulations to ensure that an adequate concentration of the vitamin is retained during processing. The form of the vitamin selected depends on how the fish feed is to be manufactured andhow long it is to be stored before being fed to the fish. It may be more
economical to overfortify channel catfish feeds with the ethycellulosecoated product than to use the phosphate derivatives of ascorbic acid;price-to-benefit relationships must be considered. Crystalline L-ascorbic
acid and L-ascorbyl-2-polyphosphate were of similar bioavailability inbroiler chicks (Pardue et al., 1993). Magnesium-L-ascorbyl-2-phosphate
is a stable form of vitamin C that was shown to be available in swine diets (Mahan et al., 1994).
Supplementation of vitamins E (see Chapter 4, Supplementation)
and C have been shown to improve oxidation stability of beef, improving lipid and color stability. Jugular infusion of sodium ascorbate 10
minutes prior to slaughter extended color display of fresh beef by 2 to 4
days (Schaefer et al., 1995). Contrary to feeding supplemental vitaminE, providing supplemental vitamin C to stabilize lipid and beef color will
likely not be practical until a ruminally stable vitamin C product is developed.
The food industry uses ascorbic acid as a natural antioxidant. For
631

CHAPTER FIFTEEN

certain human foods, enrichment or fortification with vitamin C is practiced, and vitamin C is also used in the food industry for its antioxidant
activity. The antioxidant—that is, the reducing property of ascorbic
acid—is exploited for preservation of flavor, color, and appearance of
certain foods (Nobile and Woodhill, 1981). It is used in commercial
preparation of beer, fruit juices, and canned and frozen vegetables andfruits; in meat curing; and in the flour industry to enhance baking qualities and appearance of bread. In meat processing, ascorbic acid makes
it possible to reduce both the amount of added nitrite and the residual
nitrite content in the product. In the stomach, nitrites are transformed
into potentially carcinogenic nitrosamines. The addition of ascorbic acidto fresh flour improves its baking qualities, thus saving the 4 to 8 weeks
of maturation flour would normally have to undergo after milling. As aresult of food fortification and antioxidant uses in the food industry,food that is characteristically low in vitamin C will have higher levels
than expected, provided processing methods have not destroyed theadded vitamin.
Scurvy remains an important consideration, especially for underdeveloped countries, where crop failure and malnutrition remain real concerns due to droughts, floods, pestilence, wars, and general poverty. In
more developed countries, cases of scurvy still occur occasionally (despite improved standards of living) due to inadequate diets and food faddism. Infants receiving artificial diets, alcoholics, and elderly individuals—especially elderly men living alone with low incomes—are at risk ofdeveloping overt signs of vitamin C deficiency. Vitamin C deficiency islikely to develop in individuals who do not consume optimum quantities
of fruits (particularly citrus) and vegetables.
For humans in developed countries, large quantities of supplemental vitamin C are taken as part of multivitamin pills and separately,
sometimes in megavitamin doses. There are significant new views on thehealth benefits of megavitamin doses, particularly the antioxidant vitamins (C, E, and β-carotene). Megadoses (e.g., more than 0.5 g/day) of
vitamin C have been reported beneficial for preventing or modifying
heart disease, cancer, cataracts, diabetes, and other disease conditions(see Deficiency, Humans). The beneficial effects of megadoses of vitamin
C seem obvious; however, more studies are needed to further clarify
these effects. For some subjects, excess supplemental vitamin C may be
unwise (e.g., individuals with hemochromatosis). Of lesser considerationis the method of administering large doses of vitamin C (e.g., 0.5 g or
greater) in tablets that are swallowed or chewable. Chewing large doses

632

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of the vitamin regularly can raise the acidity level in the mouth. This
may dissolve the enamel and lead to tooth damage.

TOXICITY
The reported benefits of gram doses of vitamin C have led to widespread ingestion of vitamin C supplements by many individuals, thus
raising issues of the safety of such practices. In general, high intakes ofvitamin C are considered to be of low toxicity. Safety and tolerance of
ascorbic acid in humans at levels as high as 10 g/day have been demonstrated (Koemer and Weber, 1972). Reported toxic effects include possible acidosis, gastrointestinal complaints, glycosuria or sensitivity reactions, mutagenic activity, and adverse effects concerning the metabolism
of some minerals. Barness (1977) reports that toxic effects of megadosesof vitamin C are insignificant, rare, or troublesome but of little consequence. Effects demonstrated by in vitro experiments have often later
been found to be nonexistent once definitive in vivo studies were conducted. Danger for vitamin C toxicity is minimized and unlikely in humans because of limited intestinal absorption capacity and efficient renal elimination. Nevertheless, prolonged megadose intakes of vitamin C
should be avoided as adverse effects may result, particularly for patientswith inborn errors of metabolism (e.g., cytinuria, oxalosis, and hyperuricemia).
The two greatest concerns with excess vitamin C for humans relateto (1) elevated oxalate production, which increases the risk of urinary
calculi formation, and (2) ascorbic acid-enhancing iron absorption,
which may lead to iron accumulation (e.g., hemochromatosis). In actuality, with excessive intake, the absorbed ascorbic acid is largely excreted
into the urine. Only a small amount is metabolized to oxalate regardlessof the level of dietary vitamin C. Methodological pitfalls account for
many discrepancies in findings of varying oxalate levels after vitamin C
intake. Inappropriate handling and storage of the samples favor an invitro conversion of ascorbic acid to oxalic acid. Wandzilak et al. (1994)
fed diets of 1, 5, and 10 g/day of ascorbic acid to healthy subjects andconcluded that increasingly large doses of vitamin C did not increase urinary oxalates excretion, and thus do not increase the risk of kidneystone formation in healthy individuals. Contrary to the majority of scientists, Herbert (1996) believes that most free radical damage is produced by catalytic iron and that free radical release from high body ironslowly and insidiously promotes heart disease, cancer, and premature633

CHAPTER FIFTEEN

aging in individuals with hemochromatosis. Such an effect is not to be
expected, however, as optimal iron absorption is effected with rather
low doses of vitamin C (25 to 50 mg of ascorbic acid per meal). It canbe concluded that vitamin C cannot be considered a risk factor for oxalate stones or excess iron absorption in healthy persons. Nevertheless,prudence dictates the avoidance of megadoses of vitamin C for individuals with a history of forming renal stones or patients with hemochromatosis or other forms of excess iron accumulation.
Few studies on potential vitamin C toxicity in domestic animal
species have been carried out (NRC, 1987). Chronic toxicity studies
generally indicate that ascorbic acid is well tolerated in animals. Oral
ascorbic acid may be administered to most laboratory animals at dosesof several grams per kilogram of body weight without appearance of any
obvious general effect on health. Rabbits showed only transient subconjunctival hemorrhages without other manifestations after 4 months ofdaily parenteral injections of 200 mg/kg body weight. Guinea pigs tolerated daily doses of 8.9 g/kg body weight, equivalent to 1,800 times thenormal requirement of 4 to 5 mg/kg body weight per day. Male guinea
pigs fed 8.7% ascorbic acid for 6 weeks had decreased bone density anddecreased urinary hydroxyproline compared to controls (Bray andBriggs, 1984). Helgebostad (1984) reported that high doses of 100 to
200 mg/kg body weight daily were harmful to mink with pronounced
anemia in pregnant females and reduced the number and size of kits. A
dietary ascorbic acid concentration of 1 g/kg of feed appears to pose nohazard to chickens, pigs, dogs, cats, or, probably, horses. Dietary supplementation of vitamin C even at levels as high as 3% had no appreciable effects on body weight gain, feed intake, or feed efficiency ofgrowing chicks (Nakaya et al., 1986).

■ REFERENCES
Adams, C.R. (1982). In Vitamins–The Life Essentials. Nutrition Institute, National Feed Ingredient Association. NI-82, 1-9, Des Moines, Iowa.
Anderson, R., and Lukey, P.T. (1987). Ann. N. Y. Acad. Sci. 49, 229.
Arai, S., Nose, T., and Hashimoto, Y. (1972). Bull. Freshwater Res. Lab. Tokyo22, 69.
Ausman, L.M., and Mayer, J. (1999). Nutr. Rev. 57, 222.
Banerjee, S., and Bal, H. (1959). Indian J. Med. Res. 47, 646.
Barnes, M.J., and Kodicek, K. (1972). Vitam. Horm. 30, 1.
Barness, L.A. (1977). In Re-evaluation of Vitamin C (A. Hanck and G. Ritzel,eds.), p. 23. Huber, Bern.
634

V I TA M I N C

Belfield, W.O. (1967). Vet. Med. Small Anim. Clin. 62, 345.
Blair, L., and Cummins, K.A. (1984). J. Dairy Sci. 67(Suppl. 1), 138(Abstr.).
Block, G. (1991). Am. J. Clin. Nutr. 54, 1310S.
Bouda, J., Jagos, P., Dvorak, R., and Ondrova, J. (1980). Acta Vet. Brno. 49, 53.
Bray, D.L., and Briggs, G.M. (1984). J. Nutr. 114, 920.
Brown, R.G. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J.
Tagwerker, and J. Moustgaard, eds.), p. 60. Danish Agriculture Society,
Copenhagen.
Brown, R.G., Sharma, V.D., and Young, L.G. (1970). Can. J. Anim. Sci. 50, 605.
Brown, R.G., Buchanan-Smith, J.G., and Sharma, V.D. (1975). Can. J. Anim.
Sci. 55, 353.
Cappa, C. (1958). Rev. Zooiatr. 31, 299.
Carpenter, K.J. (1986). The History of Scurvy and Vitamin C. Cambridge University Press, Cambridge.
Carvalho da Silva, A. (1950). Acta Physiol. Lat. Am. 1, 26.
Chatterjee, G.C. (1978). In Handbook Series in Nutrition and Food, Section E:
Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 2, p. 149. CRC Press, Boca
Raton, Florida.
Chatterjee, I.B., Majumder, A.K., and Nandi, B.K. (1975). Ann. N.Y. Acad. Sci.
258, 24.
Chen, H., and Chang, C. (1994). J. Nutr. 124, 2033.
Cheng, T.C., Coon, C.N., and Hamre, M.L. (1988). Poult. Sci. 67(Suppl. 1),
67(Abstr.).
Chew, B.P. (1995). J. Nutr. 125, 1804S.
Cleveland, E.R., Bondari, K., and Newton, G.L. (1987). Livestock Prod. Sci. 17,277.
Cole, C.L., Rasmussen, R.A., and Thorp, F. (1944). Vet. Med. 39, 204.
Cummins, K.A., and Brunner, C.J. (1989). J. Dairy Sci. 72, 129.
Cummins, K.A., and Brunner, C.J. (1991). J. Dairy Sci. 74, 1582.
Cunningham, J.J., Mearkle, P.L., and Brown, R.G. (1994). J. Am. Coll. Nutr. 13,
344.
Davis, G.K., and Cole, C.L. (1943). J. Anim. Sci. 2, 53.
Degkwitz, E. (1987). Ann. N.Y. Acad. Sci. 498, 470.
Dobrescu, O. (1987). Feedstuffs 54(9), 18.
Dobsinska, E., Sova, Z., Kopak, V., and Trhon, M. (1981). Vet. Med. Praha. 26,
203.
Duell, P.B. (1996). J. Nutr. 126, 1067S.
Duncan, C.W. (1944). J. Dairy Sci. 27, 636.
El-Boushy, A.R., and Van Albada, M. (1970). Neth. J. Agric. Sci. 18, 62.
El-Boushy, A.R., Simons, P.C.M., and Wiert, G. (1968). Poult. Sci. 47, 456(Abstr.).
Franceschi, R.T. (1992). Nutr. Rev. 50, 65.
Frei, B., England, L., and Ames, B.N. (1989). Proc. Natl. Acad. Sci. 86, 6377.
Fumihiko, H., Shibata, T., Makino, S., Machina, S., Hayashi, Y., Hattori, T., andYoshida, A. (1993). J. Nutr. 123, 2075.
Gershoff, S.N. (1993). Nutr. Rev. 51, 313.
Ginter, E. (1970). Acta Med. Acad. Sci. Hung. 27, 23.
Gomez-Carrasco, J.A., Lopez-Herce, C.J., and Bernabe de Frutos, C. (1994). J.
635

CHAPTER FIFTEEN

Pediatr. Gastroenterol. Nutr. 19. 118.
Grondalen, J. (1976). J. Small Anim. Prac. 17, 721.
Gross, W.G., Jones, D., and Cherry, J. (1988). Avian Dis. 32, 407.
Ha, T.Y., Otsuka, M., and Arakawa, N. (1994). J. Nutr. 124, 732.
Hagg, M., Ylikoski, S., and Kumpulainen, J. (1995). J. Food Compos. Anal. 8,
12.
Halver, J.E. (1972). Bull. Jpn. Soc. Sci. Fish 38, 79.
Harris, E.D., and Percival, S.S. (1991). Am. J. Clin. Nutr. 54, 1193S.
Harris, L., Constable, B.J., Howard, A.N., and Leader, A. (1956). Br. J. Nutr.
10, 373.
Helgebostad, A. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger,
F.J. Tagwerker, and J. Moustgaard, eds.) p. 169. Danish Agriculture Society,
Copenhagen.
Herbert, V. (1996). J. Nutr. 126, 1197S.
Hidiroglou, M. (1999). J. Dairy Sci. 82, 1831.
Hidiroglou, M., Ivan, M., and Lessard, J.R. (1977). Can. J. Anim. Sci. 57, 519.
Hill, C.H., and Garren, H.W. (1958). Poult. Sci. 37, 236.
Horio, F., Ozaki, K., Yoshida, A., Makino, S., and Hayashi, Y. (1985). J. Nutr.
115, 1630.
Hornig, D., Glatthaar, B., and Moser, U. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.) p. 3.
Danish Agriculture Society, Copenhagen.
Ikeda, S., Tahasu, M., Matsuda, T., Kakinuma, A., and Horio, F. (1997). J. Nutr.
127, 2173.
Itze, L. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.) p. 120. Danish Agriculture Society,
Copenhagen.
Itzeova, V. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J.
Tagwerker, and J. Moustgaard, eds.) p. 139. Danish Agriculture Society,
Copenhagen.
Ivos, J., Doplihar, C., and Muhaxhiri, G. (1971). Veteriwarski. Arhiv. Zagreb.
Knjiga. 41(7-8), 202.
Jacob, R.A. (1998). Nutr. Rev. 56, 334.
Jaeschke, G. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J.
Tagwerker, and J. Moustgaard, eds.) p. 153. Danish Agriculture Society,
Copenhagen.
Jacob, R.A. (1995). Nutr. Res. 15, 755.
Jacques, P.F. (1992). J. An. Coll. Nutr. 11, 139.
Jagos, P., Bouda, J., and Dvorak, R. (1977). Vet. Med. Praha. 22, 133.
Jensen, P.T., and Basse, A. (1984). In Proc. Ascorbic Acid in Domestic Animals(I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.) p. 87. Danish Agriculture Society, Copenhagen.
Jones, R.H.V., and Hothersall, J.S. (1993). Biochem. Med. Metab. Biol. 50, 197.
Kallner, A., Hartman, D., and Hornig, D. (1977). Int. Vitam. Nutr. Res. 47, 383.
Kallner, A., Hornig, D., and Pelikha, R. (1985). Am. J. Clin. Nutr. 41, 609.
Kechik, I.T., and Sykes, A.H. (1979). Br. J. Nutr. 42, 97.
Kip, D.E., Grey, C.E., McElvain, M.E., Kimmel, D.B., Robinson, R.G., and Lukert, B.P. (1996). J. Nutr. 126, 2044.
636

V I TA M I N C

Koemer, W.F., and Weber, F. (1972). Int. J. Vitam. Nutr. Res. 42, 528.
Kolb, E. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J.
Tagwerker, and J. Moustgaard, eds.) p. 162. Danish Agriculture Society,
Copenhagen.
Kranowski, J.J. (1991). J. Fla. Med. Assoc. 78, 435.
Krasnopevtsev, V.M., Istomin, A.V., Grishina, T.I., and Chizhov, S.S. (1993).
Gigiena. I. Sanitariya 6, 34.
Kristensen, B., Thomsen, P.D., Palludan, B., and Wegger, I. (1986). Acta Vet.
Scand. 27, 486.
Kuemyj, A. (1955). Vopr. Pitan. 14, 3.
Kune, G.A., Kune, S., Field, B., Watson, L.F., Cleland, H., Merenstein, D., andVitetta, L. (1993). Nutr. Cancer 20, 61.
Lehocky, J. (1981). Veterinarstvi 31, 388.
Leveque, J.I. (1969). Vet. Med. Small Anim. Clin. 64, 997.
Lewis, L.D., and Morris, M.R. (1983). Small Animal Clinical Nutrition. MarkMorris Associates, Topeka, Kansas.
Li, Y., and Lovell, R.T. (1985). J. Nutr. 115, 123.
Liau, L.S., Lee, B.L., New, A.L., and Ong, C.N. (1993). J. Chromatogr. 612, 63.
Lin, H.K., Chen, S.Y., Huang, C.Y., Kuo, M.H., and Wung, L.C. (1985). Ann.
Res. Rep. Anim. Ind. Res. Inst. TSC, 59.
Lovell, R.T. (1973). J. Nutr. 103, 134.
Lovell, R.T., and Lim, C. (1978). Trans. Am. Fish Soc. 107, 321.
Luck, M.R., Jeyaseelan, I., and Scholes, R.A. (1995). Biol. Reprod. 52, 262.
Magarelli, P.C., Hunter, B., Lightner, D.V., and Colvin, L.B. (1979). Comp.
Biochem. Physiol. 63, 103.
Mahan, D.C., Lepine, A.J., and Dabrowski, K. (1994). J. Anim. Sci. 72, 2354.
Mahan, D.C., Pickett, R.A., Perry, T.W., Curtin, T.M., Featherson, W.R., andBeeson, W.M. (1966). J. Anim. Sci. 25, 1019.
Mangels, A.R., Block, G., Frey, C.M., Patterson, B.H., Taylor, P.R., Norkus,
E.P., and Levander, O.A. (1993). J.Nutr. 123, 1054.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
Martynjuk, B.F. (1952). Med. Veter. 29, 21.
Matusiewicz, M., Dabrowski, K., Volker, L., and Matusiewicz, K. (1995). J.
Nutr. 125, 3055.
McDonald, P., Edwards, R.A.,and Greenhalgh, J.F.D. (1981). Animal Nutrition,
3rd Ed., p. 83. Longman, New York.
McKee, J.S., and Harrison, P.C. (1995). Poult. Sci. 74, 1772.
Meier, H., Clark, S.T., and Schnelle, G.B. (1957). Am. Vet. Med. Assoc. 130,
483.
Monsi, A., and Onitchi, D.O. (1991). Anim. Feed Sci. Tech. 34, 141.
Moser, U., and Bendich, A. (1991). In Handbook of Vitamins, 2nd Ed. (L.J.
Machlin, ed.) p. 195. Dekker, New York.
Mukhopadhyay, C.K., Ghosh, M.K., and Chatterjee, I.B. (1995). Mol. Cell
Biochem. 142, 71.
Naismith, D.H. (1958). Proc. Nutr. Soc. 17, xlii.
Nakaya, T., Suzuki, S., and Watanabe, K. (1986). Jpn. Poult. Sci. 23, 276.
Navarre, O., and Halver, J.E. (1989). Aquaculture 79, 207.
637

CHAPTER FIFTEEN

Nielsen, N.C., and Vinther, K. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.) p. 39. Danish
Agriculture Society, Copenhagen.
Njoku, P.C. (1984). Feedstuffs 56(52), 23.
Njoku, P.C. (1986). Anim. Feed Sci. Tech. 16, 17.
Nobile, S., and Woodhill, J.M. (1981). Vitamin C. MTP Press (International
Medical Publ.), Boston.
Nockels, C.F. (1984). In Proc. Ascorbic Acid in Domestic Animals (I. Wegger,
F.J. Tagwerker, and J. Moustgaard, eds.) p. 175. Danish Agriculture Society,
Copenhagen.
Noll, S. (1993). Personal communication to Tillman, P.B. Feed Manage. 44(10),31.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1977). Nutrient Requirements of Rabbits, 2nd Ed.
(1978). Nutrient Requirements of Nonhuman Primates.
(1985). Nutrient Requirements of Dogs, 2nd Ed.
(1986). Nutrient Requirements of Cats.
(1993). Nutrient Requirements of Fish.
(1995). Nutrient Requirements of Laboratory Animals.
(1998). Nutrient Requirements of Swine, 10th Ed.
NRC. (1987). Vitamin Tolerance of Animals. National Academy of Sciences-National Research Council, Washington, D.C.
Olivares, M., Pizarro, F., Pineda, O., Name, J.J., Hertrampf, E., and Walter, T.
(1997). J. Nutr. 127, 1407.
Orban, J.I., Roland, D.A., Cummins, K., and Lovell, R.T. (1993). Poult. Sci. 72,
691.
Padh, H. (1991). Nutr. Rev. 49, 65.
Palludan, B., and Wegger, I. (1984). In Proc. Ascorbic Acid in Domestic Animals(I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.) p. 131. Danish Agriculture Society, Copenhagen.
Pardo, B., Sygnowska, E., Rywik, S., Kulesza, W., and Waskiewicz, A. (1991).
Appetite 16, 1.
Pardue, S.L. (1987). Proc. The Role of Vitamins on Animal Performance and Immune Response, p. 18. Hoffmann-La Roche, Nutley, New Jersey.
Pardue, S.L., and Thaxton, J.P. (1982). Poult. Sci. 61, 1522(Abstr.).
Pardue, S.L., and Williams, S.H. (1990). In Proc. 2nd Symp. Ascorbic Acid in
Domestic Animals. Kartause Ittingen, Switzerland, Oct. 9.
Pardue, S.L., Thaxton, J.P., and Brake, J. (1985). Poult. Sci. 64, 1334.
Pardue, S.L., Brake, J., Seib, P.A., and Wang, X.Y. (1993). Poult. Sci. 72, 1330.
Park, T.F., and Harrison, P.C. (1990). Growth performance of nursery pigs provided tap and carbonated drinking water sources supplemented withmonopotassium ascorbate. Personal communication. Dept. of Anim. Sci.,
Univ. of Illinois, Urbana, Illinois.
Pauling, L. (1971). Vitamin C and the Common Cold. Freeman, San Francisco.
Peebles, E.D., and Brake, J. (1985). Poultry Sci. 64, 2041.
Perek, M., and Kendler, J. (1963). Br. Poult. Sci. 4, 191.
Pietronigro, D.D., Hovsepian, M., Demopoulos, H.B., and Flamm, E.S. (1983).
638

V I TA M I N C

J. Neurochem. 41, 1072.
Podgornova, G.P., and Donskova, T.J. (1972). Voprosy Mofologii Ekologii i
Parazitologii Zhivotnykh 1, 137.
Pribyl, E. (1963). Diseases of Young Cattle, 230 p. SZN, Praha.
Quarles, C.L., and Adrian, W.J. (1989). In The Role of Vitamin C in Poultry
Stress Managment, p. 37. RCD 7839. Hoffmann-La Roche, Inc., Nutley,
New Jersey.
Quarles, C.L., Adrian, W.J., and Krautmann, B.A. (1989). Tenth Ann. Mtg.
South. Poult. Sci. Soc. p. 34(Abstr.).
RDA. (1989). Recommended Dietary Allowances 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
Rifici, V.A., and Khachadurian, A.K. (1993). J. Am. Coll. Nutr. 12, 631.
Rose, R.C., and Nahrwold, D.L. (1982). Anal. Biochem. 123, 389.
Rose, R.C., McCormick, D.B., Li, T.K., Lumeng, L., Haddad, J.G., and Spector,
R. (1986). Fed. Proc. Fed. Am. Soc. Exp. Biol. 45, 30.
Roth, J.A., and Kaeberle, M.L. (1985). Am. J. Vet. Res. 46, 2434.
Rucker, R.B., Dubick, M.A., and Mouritsen, J. (1980). Am. J. CLin. Nutr. 33,
961.
Sandholm, M., Honkanen-Buzalski, R., and Rasi, V. (1979). Vet. Rec. 104, 337.
Satterfield, G.H., Mosley, M.A., Gauger, H.C., Holmes, A.D., and Tripp, F.
(1940). Poult. Sci. 19, 337.
Sauberlich, H.E. (1990). In Nutrition Reviews, Present Knowledge in Nutrition:(R.E. Olson, ed.), p. 132. Nutrition Foundation, Washington, D.C.
Schaefer, D.M., Liu, Q., Faustman, C., and Yin, M. (1995). J. Nutr. 125, 1792S.
Scott, D.W. (1981). Bov. Practice 2, 22.
Shaw, P.E., and Moshonas, M.G. (1991). J. Food Sci. 56, 867.
Siegel, B.V. (1974). Infect. Immunol. 10, 409.
Sigman-Grant, M., Bush, G., and Anantheswaran, R. (1992). Pediatrics 90, 412.
Simon, J.A. (1992). J. Am. Coll. Nutr. 11, 107.
Singh, R.B., Niaz, M.A., Agarwal, P., Begom, R., and Rastogi, S.S. (1995). J.
Am. Diet. Assoc. 95, 775.
Snehalatha Reddy, N., and Lakshmi Kumari, R. (1988). Nutr. Rep. Int. 37, 77.
Soldatenkov, P.F., and Suganova, N.M. (1966). Selskochoz. Biol. 1, 446.
Spencer, R.P., Purdy, S., Hoeldtke, R., Bow, T.M., and Markulis, M.A. (1963).
Gastroenterology 44 768.
Stocker, R., and Frei, B. (1991). In Oxidative Stress (H. Sies, ed.), p. 213. Academic Press, United Kingdom.
Takahashi, K., Akiba, Y., and Horiguchi, M. (1991). Br. Poult. Sci. 32, 545.
Toohey, L., Harris, M.A., Allen, K.G.D., and Melby, C.L. (1996). J. Nutr. 126,
121.
Tono, T., and Fujita, S. (1982). Agric. Biol. Chem. 46, 2953.
Turnbull, J.D., Sudduth, J.H., Saberlich, H.E., and Omaye, S.T. (1981). Int. J.
Vitam. Nutr. Res. 51, 47.
Ukhun, M.E., and Dibie, E.N. (1991). Food-Chem Essex 41, 277.
Vilter, R.W. (1978). In Handbook Series in Nutrition and Food. Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 3, p. 91. CRC Press, Boca Raton, Florida.
Wandzilak, T.R., D’Andre, S.D., Davis, P.A., and Williams, H.E. (1994). J. Urol.
639

CHAPTER FIFTEEN

151, 834.
Weber, P. (1994). In Proc. Roche Technical Seminar, Antioxidant Vitamins, p.
134. Hoffmann-La Roche Inc., Nutley, New Jersey.
Wegger, I., and Palludan, B. (1984). In Proc. Ascorbic Acid in Domestic Animals(I. Wegger, F.J. Tagwerker, and J. Moustgaard, eds.), p. 68. Danish Agriculture Society, Copenhagen.
Wegger, I., and Palludan, B. (1994). J. Nutr. 124, 241.
Weiser, H., Schlachter, M., Probst, H.P., and Kormann, A.W. (1990). In Proc.
2nd Symp. Ascorbic Acid in Domestic Animals. Kartause Ittingen, Switzerland, Oct. 9.
Wells, W.W., Dou, C.Z., Dybas, L.N., Jung, C.H., Kalbach, H.L., and Xu, D.P.
(1995). Proc. Natl. Acad. Sci. 92, 11869.
Wetherilt, H., Ackurt, F., Brubacker, G., Okan, B., Aktas, S., and Turdu, S.
(1992). Int. J. Vitam. Nutr. Res. 62, 21.
Yashin, A.V. (1985). Veterinariya (Moscow) 1, 57.
Yen, J.T., and Pond, W.G. (1981). J. Anim. Sci. 53, 1291.
Yen, J.T., and Pond, W.G. (1984). J. Anim. Sci. 58, 132.
Yen, J.T., and Pond, W.G. (1987). J. Anim. Sci. 64, 1672.
Zapata, L.F., and Gernat, A.G. (1995). Poult. Sci. 74, 1049.
Zee, J.A., Carmichael, L., Codere, D., Poirier, D., and Fournier, M. (1991). J.
Food Compos. Anal. 4, 77.
Zintzen, H. (1975). A Guide to the Nutritional Management of Breeding Sows
and Pigs No. 1465. Hoffmann-La Roche & Co. Ltd., Basel, Switzerland.

640

chapter sixteen

CARNITINE

INTRODUCTION
Under most conditions for the majority of species, carnitine would
not be considered a vitamin as it is adequately synthesized in body tissues. However, the need for supplemental carnitine has been demonstrated in mammals in circumstances in which the biosynthesis is limited
by nutritional deprivation of the precursor amino acids lysine and methionine. Dietary carnitine is essential for some insect species, including
beetles of the family Tenebrionidai (mealworms), the beetle Oryzaephilus surinamensis, and the fly Drosophila melanogaster. For these
species it is appropriate to refer to carnitine as a vitamin.

HISTORY
Carnitine was isolated from meat extracts and identified in 1905. In
1948, Fraenkel’s research on dietary requirements of the mealworm(Tenebrio molitor) led to recognition of a new B vitamin, which in 1932
was identified as carnitine (Friedman and Fraenkel, 1972). Since it wasa small water-soluble compound required in the diet of T. molitor, it wasgiven the name vitamin BT (sometimes referred to as vitamin B7). It was
not until the 1960s that carnitine was recognized as a biologically active
substance (Borum, 1991).

CHEMICAL STRUCTURE AND PROPERTIES
Carnitine is a quaternary amine, β-hydroxy-γ-trimethylaminobutyrate(Fig. 16.1). It is a very hygroscopic compound, easily soluble in water,and has a molecular weight of 161.2. Carnitine is found in biological641

16

CHAPTER SIXTEEN

Fig. 16.1 Chemical
structure of carnitine.

samples both as the free carnitine and as the ester of a wide variety ofacyl compounds. Of the two types of carnitine, L- and D-carnitine, only
L-carnitine is biologically active.

ANALYTICAL PROCEDURES
Methods of analysis first utilized the bioassay technique using T.
molitor. Other methods developed for carnitine determination include
chemical, enzymatic, gas chromatographic, and radioisotopic procedures (Chan, 1984). The most commonly used methodology to determine the carnitine concentration in biological samples is the radioenzymatic assay (Rossle et al., 1985). Methods such as radioisotopic assays
are sensitive; the radioactive acylcarnitine formed is separated fromresidual acetyl CoA by cation exchange filtration, and radioactivity is
counted (Deufel, 1990).
Development and use of high-performance liquid chromatography(HPLC) is common (Minkler and Hoppel, 1993) for carnitine analysis,
as is the use of radioisotopic exchange/HPLC methods (Schmidt-Sommerfeld et al., 1992). Recently, enzyme spectrophotometer assays (Indyk
and Woollard, 1995) and a modified enzyme-spectrophotometry
method (Tuan and Chou, 1995) have been used to determine carnitine
in milk and infant formulas.

METABOLISM
Under normal conditions in omnivores, about 70 to 80% of dietarycarnitine is absorbed (Rebouche and Chenard, 1991). Carnitine appears
to be absorbed across the proximal small intestine by an active processdependent on Na+ as well as by a passive diffusion, which may be important for the absorption of large doses of the factor. The uptake ofcarnitine from the intestinal lumen into the mucosa is rapid, and about
one-half of the carnitine taken up is acetylated in that tissue. Carnitine
is not carried in blood in any tightly bound forms, in contrast to many
water-soluble vitamins. Tissues such as cardiac and skeletal muscle re642

CARNITINE

quire carnitine for normal fuel metabolism but cannot synthesize carnitine and are totally dependent on the transport of carnitine from othertissues. Cantrell and Borum (1982) reported that carnitine uptake by theheart is facilitated by a cardiac carnitine-binding protein.
Carnitine is synthesized in liver and kidney and stored in skeletal
muscle; free carnitine is excreted mainly in the urine (Tanphaichitr andLeelahagul, 1993). The product is trimethylamine oxide (Mitchell,1978). Carnitine is highly conserved by the human kidney, which reabsorbs more than 90% of filtered carnitine, thus playing an importantrole in the regulation of carnitine concentration in blood.
Carnitine synthesis depends on two precursors, L-lysine and methionine, as well as ascorbic acid, nicotinamide, vitamin B6, and iron
(Borum, 1991). Deficiency in any cofactor will cause L-carnitine deficiency. In rats, total acid-soluble carnitine and free carnitine in plasma
and tissues were reduced in a vitamin B6 deficiency but increased whenvitamin B6 was provided in a repletion diet (Cho and LeKlem, 1990; Haet al., 1994). It has been suggested that early features of scurvy (fatigue
and weakness) may be attributed to carnitine deficiency. Vitamin C is a
cofactor for two α-ketoglutarate-requiring dioxygenase reactions (epsilon-N-trimethyllysine hydroxylase and γ-butyrobetaine hydroxylase)in the pathway of carnitine biosynthesis. Carnitine concentrations arevariably low in some tissues of vitamin C-deficient guinea pigs (Rebouche, 1991). The results of studies of enzyme preparations and perfused liver in vitro, and of scorbutic guinea pigs in vivo, provide compelling evidence for participation of ascorbic acid in carnitine
biosynthesis (Rebouche, 1991). Results reported by Ha et al. (1991)
suggest that ascorbic acid is specifically required for the hydroxylationof γ-butyrobetaine and, furthermore, that ascorbic acid can regulate carnitine synthesis in primary cultured liver cells from guinea pigs.
Choline has also been shown to affect carnitine homeostasis in humans and guinea pigs (Daily and Sachan, 1995). Choline supplementation resulted in decreased urinary excretion of carnitine in young adultwomen, and choline resulted in a conservation of carnitine in guinea
pigs. In choline-deficient rats, a single injection of choline raised the concentration of liver carnitine within 1.5 hours (Carter and Frenkel, 1978).
This suggests that choline was capable of facilitating carnitine releasefrom some storage pool, as de novo synthesis would require more time.
In the rat, about 1/15 to 1/20 of the body pool turns over each day,consistent with the slow rate of turnover in muscle, where most of the
body carnitine is stored (Bremer, 1983). For the dog, 95 to 98% of thecarnitine body pool is in skeletal muscle and heart (Rebouche and En643

CHAPTER SIXTEEN

gel, 1983). Flores et al. (1996) reported that the small intestine in rats is
a considerable and previously unrecognized proportion of the carnitine
pool of suckling animals.

FUNCTIONS
Carnitine is an important cofactor for normal cellular metabolism.
Optimal utilization of fuel substrates for adenosine triphosphate (ATP)generation by skeletal muscle during exercise is dependent on adequatecarnitine stores. Carnitine is required for transport of long-chain fatty
acids into the matrix compartment of mitochondria from cytoplasm for
subsequent oxidation by the fatty acid oxidase complex for energy production. The oxidation of long-chain fatty acids in animal tissues is dependent on carnitine because it allows long-chain acyl-CoA esters tocross the mitochondrial membrane, which is otherwise impermeable to
CoA compounds. Carnitine facilitates the β-oxidation of long-chain
fatty acids in the mitochondria by transporting the substrate into the mitochondria. Carnitine acyltransferase is the enzyme responsible for thisshuttle mechanism. It exists in two forms, carnitine acyltransferase I andcarnitine acyltransferase II. After the long-chain fatty acid is activated toacyl-CoA, it is converted to acylcarnitine by the enzyme carnitine acyltransferase I and crosses to the matrix side of the inner mitochondrial
membrane. Carnitine acyltransferase II then releases carnitine and theacyl-CoA into the mitochondrial matrix. Acyl-CoA is then catabolized
via β-oxidation (Mitchell, 1978; Borum, 1991). Thus, utilization of
long-chain fatty acids as a fuel source depends on adequate concentrations of carnitine.
The role of carnitine in the transport of long-chain fatty acids acrossthe inner mitochondrial membrane is known. However, liver mediumchain fatty acid (MCFA) metabolism has been considered carnitine independent because of their passive diffusion through the inner mitochondrial membrane and intramitochondrial activation (Bremer, 1990).
However, evidence suggests that MCFA metabolism may be affected bysupplemental carnitine (Rebouche et al., 1990; Van Kempen and Odle,
1993, 1995).
Another role of carnitine may be to protect cells against toxic accumulation of acyl-CoA compounds of either endogenous or exogenous
origin by trapping acyl groups such as carnitine esters, which may thenbe transported to the liver for catabolism or to the kidney for excretionin the urine. It has been reported that nicotinamide and L-carnitine protect human cells from oxygen free radical-induced damage and that this644

CARNITINE

makes these compounds possible antiaging substances (Monti et al.,1992). Carnitine also has functions in other physiological processes critical to survival, such as lipolysis, thermogenesis, ketogenesis, and possibly regulation of certain aspects of nitrogen metabolism (Borum, 1985).
Carnitine administration has significant benefits in patients with disorders of ammonia metabolism, including urea cycle defects, chronic valproic acid therapy, liver failure, organic acidemias, and Reye’s syndrome(Rebouche, 1992). Carnitine plays a role in ammonia detoxification(Sakemi et al., 1992; Freeman et al., 1994; Melegh et al., 1994). Propionyl-L-carnitine protects the ischemic heart from reperfusion injury, perhaps by scavenging free radicals or preventing their formation by chelating iron necessary for generation of hydroxyl radicals. Its function as anantiarrhythmic agent for the heart has been reported (Pande andMurthy, 1989; Reznic et al., 1992). An additional function of carnitine
is as a memory- and alertness-enhancing agent in Alzheimer’s disease(Kendler, 1986; Corrigan et al., 1995).

REQUIREMENTS
Carnitine is an essential growth factor for some insects, such as themealworm. However, most insects and higher animals, as well as mammals, can synthesize carnitine. For common species there are no established nutritional requirements for carnitine. Studies have indicated thatthe biosynthesis of carnitine may be limited or inadequate in certainclasses of humans and animals. Because the young of several species, including humans, have been found to have low tissue carnitine when fed
low-carnitine diets, it is likely that the total carnitine biosynthetic capacity may be immature in newborns, thus they are dependent on theirdiets for preformed carnitine.
In omnivores, carnitine synthesis normally provides only about oneeighth to one-half of total carnitine available to the organism, whereas
in strict vegetarians, endogenous carnitine synthesis provides more than
90% of total available carnitine. Normal adult humans both synthesize
and ingest carnitine in amounts totaling 100 mg/day. Healthy children
and adults can synthesize up to 20 mg of endogenous carnitine daily(Anonymous, 1985). In mammals, γ-butyrobetaine (YBB), the immediate precursor of carnitine, can be synthesized from the essential aminoacids lysine and methionine in most tissues. The four-carbon chain
comes from lysine; the methyl groups come from methionine. The ultimate conversion of YBB to carnitine occurs in the liver (Olson and Rebouche, 1987).
645

CHAPTER SIXTEEN

NATURAL SOURCES
In general, foods of plant origin are low in carnitine, whereas animal-derived foods are rich in carnitine (Table 16.1) (Mitchell, 1978).
Red meats and dairy products are particularly rich sources. The carnitine concentration increases in the order of fish, poultry, pork, and beef.
In general, the redder the meat, the higher the concentration of carnitine.
Typical concentrations of carnitine could be 600 μg/kg in beef, 45 to 90
μg/kg in chicken, and 75 μg/kg in lamb (Mitchell, 1978). On the contrary, grains such as barley, corn, and wheat have undetectable or negligible concentrations.
Carnitine is located principally in skeletal muscle, which has about
40 times the concentration of carnitine in blood. Average muscle content
of total carnitine ranges from 10.75 to 19.06 nmol/mg of noncollagenprotein. Most plant foods that are low in carnitine are also likely to be
low in lysine and methionine, the precursors of carnitine. Therefore, apure vegetarian diet may lack both preformed carnitine and its precursors. Strict vegetarian diets contain less than 10% as much carnitine as
typical omnivorous diets of the developed nations. Nevertheless, vegetarian diets furnishing adequate lysine, methionine, and micronutrientsfor carnitine biosynthesis should theoretically maintain normal carnitine
nutrition in the healthy individual.
Milk is essential for the nursing mammal to supply carnitine. Although carnitine is synthesized in growing young and adult animals, previous studies in humans provide evidence that exogenous carnitine is necessary to maintain normal fat metabolism during infancy. Healthy
full-term infants fed formulas devoid of carnitine show reduction in the
products of ketogenesis and accumulation of fatty acid precursors inplasma compared with infants provided carnitine in either cow’s milk formula or breast milk (Novak et al., 1983). Studies by Davis (1989) indicated that up to 50% of tissue carnitine in suckling rats is derived fromthe mother’s milk. Studies with neonatal rabbits (Penn and Schmidt-Sommerfeld, 1988) and rats (Flores et al., 1996) demonstrated that body tissues of carnitine are greatly diminished in newborns deprived of milk
during early life. Coffey et al. (1991) showed that diminished dietary intake is associated with decreased levels of carnitine in the liver, but notheart or muscle, in neonatal piglets receiving low levels of dietary carnitine compared with piglets receiving carnitine supplementation. Theseobservations indicate the importance of milk. The demand for carnitine
during the suckling period may exceed the capacity for its synthesis.
646

CARNITINE■ Table 16.1

Typical Carnitine Concentrations in Various Foods and Feedstuffs (μg/100 g)

Plant Deriveda
Alfalfa concentrate
AvocadoCauliflower
PeanutTorula yeast

2.00
1.25
0.13
0.76
1.60–3.29

Animal Derived
Beef, muscle
Beef heart
Beef kidney
Beef liver
Chicken, muscle
Cow’s milk
Lamb, muscle

59.8–67.4
19.3
1.8
2.6
4.6–9.1
0.53–3.91
78.0

Source: Mitchell (1978).
aCarnitine was not detected in analyses of cabbage,orange juice, spinach, corn, or egg.

DEFICIENCY
In carnitine deficiency, fatty acid oxidation is reduced, and fatty
acids are diverted into triglyceride synthesis, particularly in the liver. Mitochondrial failure develops in carnitine deficiency when there is insufficient tissue carnitine available to buffer toxic acyl-coenzyme (acylCoA) metabolites. Toxic amounts of acyl-CoA impair the citrate cycle,
gluconeogenesis, the urea cycle, and fatty acid oxidation. Carnitine replacement treatment is safe and induces excretion of toxic acyl groups
in the urine (Stumpf et al., 1985).
If carnitine deficiency involves the liver, the supply of ketones andthe utilization of long-chain fatty acids during starvation are cut off; alltissues become glucose dependent. When liver carnitine is depleted, starvation tends to cause nonketotic, insulinopenic hypoglycemia. Becauseliver hepatocytes depend on fatty acids for their energy requirements
during fasting, carnitine depletion may also cause clinical liver dysfunction, shown by hyperammonemia, encephalopathy, and hyperbilirubinemia (Feller and Rudman, 1988). Skeletal muscles are generally involved,
with weakness, lipid myopathy, and myoglobinuria often aggravated or
precipitated by fasting or exercise. The heart, like skeletal muscle, is dependent on fatty acids for energy during fasting, and heart failure andarrhythmias are frequent manifestations of systemic carnitine deficiency.
647

CHAPTER SIXTEENThe heart derives approximately 60% of its ATP supply from β-oxidation of fatty acids. Carnitine concentrations in the heart are normallyvery high in many species (Rebouche and Paulson, 1986).

Effects of Deficiency
Ruminants
Research conducted with both steers and heifers fed high-roughage
diets supplemented with soybean meal resulted in increased weight gains
when supplemented with carnitine, while other studies have showed noeffect (Hill et al., 1995). Diets high in forage or roughage content generally shift the ruminal volatile fatty acid profile in the direction of moreacetate production and less propionate production, and carnitine is reported to be important in the regulation of liver and blood acetate levels.
Feeding veal male Holstein calves from 140 days with or without
supplement of 1, 3, or 5 g/day of L-carnitine resulted in mean daily
weight gains of 1.12, 1.14, 1.19, and 1.23 kg, while carcass yield was
64.1, 64.7, 65.5, and 66.4%, respectively. The auxinic effect of carnitine
is attributed to its hyperglycemic activity and favorable effect on lipid
metabolism (Bonomi et al., 1991). Intravenous L-carnitine significantly
lowered plasma ammonia N levels in ewes given oral urea (Chapa et al.,1998). Addition of L-carnitine to ruminant feeds could alleviate hyperammonemia experienced by ruminants that consume a high level of nonprotein N.
The most popular topic of carnitine research in ruminants has beenthat dealing with ketonemia, an economically important problem commonly known as pregnancy toxemia in sheep, and ketosis, acetonemia,
and ketoacidosis in cattle. Ketosis occurs when body reserves are mobilized for productive functions; the condition is exacerbated by restriction of energy or protein intake. It is hypothesized that ketosis is the result of the derepression of carnitine acyltransferase, which allows large,
uncontrolled amounts of fatty acids to enter the liver mitochondria,
where they are metabolized to acetoacetate and subsequently form β-hydroxybutyrate, both of which accumulate.
In lactating dairy cows, carnitine supplementation increased concentrations of carnitine in plasma and liver and improved lipid digestibility (La Count et al., 1995). However, in three dairy cattle experiments, supplemental carnitine (6 to 12 g/day) did not benefit milk yieldor milk composition but was effective at increasing the concentrationsof carnitine in liver, plasma, and milk of dairy cows during early lactation (La Count et al., 1995, 1996a,b).
648

CARNITINESwine
The capacity of swine to synthesize carnitine has not been directly
examined. However, plasma and tissue concentrations of carnitine are
reduced in neonatal pigs reared on formulas devoid of carnitine (Baltzell
et al., 1987). Supplementation with carnitine in 4-kg mini-pigs during
total parenteral nutrition was shown to increase their energy gain fromexogenous fat and to increase their nitrogen retention fourfold (Bohles
et al., 1984). Supplemental carnitine resulted in increased growth rate atmoderately high lysine intakes, but tended to depress growth rate whenlysine levels were near, below, or in considerable excess of published requirements (Newton and Burtle, 1992). Additionally, it was found thatpigs that were lightweight at weaning (generally 5 kg or less) were more
likely to have a positive growth response to supplemental carnitine overthe 4-week nursery period than heavier pigs. Hoffman et al. (1993)
found no benefit from supplemental L-carnitine to young pigs. Likewise,
addition of L-carnitine to the lactation diet had little effect on the performance of the first parity sow. However, decreased feed intake during
the first week of lactation and a tendency toward fewer days to estrus
were observed (Musser et al., 1997).
Dietary L-carnitine has been found to improve the gain:feed ratio
and reduces carcass lipid accretion in early weaned pigs fed 1,000 ppm
of L-carnitine (Owen et al., 1996). In newborn pigs, medium-chain fatty
acid (MCFA) oxidation accounted for 40% of the MCFA infused, andcarnitine, independent of the level, increased the fatty acid oxidation by
as much as 20% if the energy provided as MCFA exceeded 50% of themetabolic needs of the pig (Van Kempen et al., 1993).
From results of trials with finishing pigs, there was a curvilinear response in growth rate for the first 14 days, and in back-fat thickness at
slaughter weight, due to carnitine intake. Growth rate and back fat wereboth increased at higher carnitine intakes (greater than 0.4 mg/kg ofbody weight per day). There was also a differential response due to sex,
with female pigs responding to a much greater degree than male castrates (Newton and Haydon, 1989).

Poultry
The increases in plasma and hepatic acylcarnitines in broilers fed
0.5% L-carnitine indicated that supplementary carnitine lessens the loadof free acyl groups in the liver by eventual oxidation or excretion (Smithet al., 1994). In another study, Barker and Sell (1994) found that carnitine intake (0, 50, and 100 mg/kg) did not affect body weight, feed con649

CHAPTER SIXTEEN

version efficiency, or proximate composition at 21 days in turkeys andat 45 days in broilers. L-Carnitine significantly increased cholesterol in
egg yolk when added with or without nicotinic acid at 500 mg/kg. With
50 and 100 mg/kg in the feed of layer hens, hatchability increased by 4
and 2.9%, respectively (Leibetseder, 1995).

HorsesWorking with horses in combined training, the animals were tested
in exercise trials and were given 5 g/day of L-carnitine with oats. It was
found that carnitine did not improve performance, but there was a lower
increase of lactate in the carnitine group during exercise and a lower increase of enzyme activity of lactate dehydrogenase. It was concluded
that carnitine, besides its function in mitochondria energy-linked
processes, also improves membrane stability (Iben et al., 1992). In broodmares supplemented with 10 g/day of L-carnitine, it was found that the
decline in milk carnitine after foaling was reversed in supplemented animals but not in unsupplemented ones. The drop in foal plasma carnitine was also reversed in foals from supplemented mares, and there were
no apparent side effects of carnitine supplementation (Benamou andHarris, 1993).

Other Animal Species
DOGS
In dogs with dilated cardiomyopathy (DCM), myocardial concentrations of L-carnitine are sometimes very low. Carnitine deficiency associated with DCM was documented in Doberman pinschers (Keene etal., 1989) and a family of boxers (Keene et al., 1986). A few years later,
Keene (1992) reported DCM in a wide range of dog breeds and foundthat myocardial free L-carnitine deficiency occurred in 50 to 90% of
dogs with DCM. In these dogs, the myocardial concentrations of carnitine were very low, and substantial clinical improvement after intravenous or oral therapy with L-carnitine was observed. The possible explanation is that these dogs have a membrane transport defect thatprevents adequate quantities of carnitine from moving into the heartfrom the plasma (Keene, 1991; Keene et al., 1991). McEntee et al.
(1995) reported DCM in a female Labrador, whose diet was exclusively
from vegetables and cereals because of a presumed allergy to animal
proteins. The dog had a poor appetite, coughing, abdominal distention,
exercise intolerance, and a peculiar body odor. After 4 days of carnitine650

CARNITINE

supplementation, the dog showed spectacular improvement in appetite,
increased tolerance to exercise, and reduction of the body odor. The peculiar body odor was also described in children with carnitine deficiency(Waber et al., 1982).
Grandjean et al. (1993) demonstrated the need of supplemental carnitine for working husky dogs. After exercise, carnitine-supplemented
dogs showed better utilization of lipids, lower free fatty acid blood levels, more stable blood glucose levels, and less accumulation of lactate
residues.
Systemic and myopathic forms of L-carnitine deficiency are wellknown etiologies of DCM in human medicine. Evidence suggests thatcongestive heart failure caused by rapid ventricular pacing in dogs is also
associated with myocardial carnitine deficiency (Keene, 1994).

FISH
Accelerated growth and reduced body fat have been reported for
hatchery-reared sea bass (Santulli and D’Amelio, 1986; Santulli et al.,1988) and African catfish fed carnitine (Torreele et al., 1993). Atlantic
salmon fed L-carnitine exhibited altered intermediary metabolism andreduced tissue lipid, but no change in growth rate (Ji et al., 1996). These
results implicated induction of pyruvate carboxylase (or a reduction in
turnover) and enhanced protein synthesis in the mechanism for carnitine-induced changes in gluconeogenesis and nitrogen metabolism.
Supplementation of carnitine (500 mg/kg of diet) in catfish fingerlings produced a nonsignificant increase in growth rate; 1,000 mg/kg
produced a significant increase in growth rate and feed efficiency, andthe response to 2,000 mg/kg was approximately equal to that observedwith 500 mg/kg. Fish fed 2,000 mg/kg, however, had significantly lesslipid in their fillets than fish fed the control diet (Newton and Burtle,
1992).

LABORATORY ANIMALS
Rats have not been reported to be deficient in carnitine. However,rats fed diets deficient in lysine or methionine were shown to develop
mild depressions in tissue carnitine concentrations and to suffer growthdepression and fatty liver, both of which are at least partially alleviatedby feeding carnitine. In guinea pigs, vitamin C deficiency was linked to
impairment in carnitine synthesis, increased urinary carnitine excretion,and prolonged survival time with carnitine supplementation (Alkonyi etal., 1990).
651

CHAPTER SIXTEEN

Humans
It was formerly assumed that because humans have the ability tosynthesize carnitine, this compound is not an essential nutrient. However, since 1973 it has been described as “conditionally essential” because it was discovered that certain segments of the humanpopulation—mainly preterm infants, normal infants, adults on total parenteral nutrition, and adults and children with a variety of genetic, infectious, and injury-related illnesses—have a need for carnitine.
Borum (1981) presented evidence that newborns have a critical needfor carnitine since they have not attained the full biosynthetic capacity
for carnitine, and their plasma and tissue concentrations are low. Newborn infants depend heavily on lipids as a concentrated source of fuel toachieve rapid growth during the first months of life (Shenai and Borum,
1984; Nakano et al., 1989). It has been estimated that the amount ofcarnitine in skeletal muscle of very premature infants, indexed to wholebody weight, is about 10 times less than that of adults (Schmidt-Sommerfeld and Penn, 1990). In premature infants, Helms et al. (1990) observed increased nitrogen balance and weight gain in intravenously fed
infants supplemented with carnitine.
Bonner et al. (1995) concluded that very low birth weight infants requiring prolonged parenteral nutrition have carnitine deficiency with
impaired ketogenesis. Parenteral administration of carnitine appears toalleviate this metabolic disturbance. In low birth weight infants, lowered
plasma-free carnitine produced a higher blood ammonia (Nakamura etal., 1990).
Human carnitine deficiency can be hereditary or acquired. Hereditary carnitine deficiency can be grouped into three clinical entities: myopathic carnitine deficiency, systemic carnitine deficiency, and organic
acidurias. Acquired carnitine deficiency is due to inadequate intake, increased requirement, and increased loss of carnitine (Tanphaichitr andLeelahagul, 1993).
Myopathic and systemic abnormalities are classified as autosomal
recessive disorders (DiMauro, 1979). A defective transport system forcarnitine uptake has been proposed as the cause of the myopathic disorder (Engel, 1980; Bremer, 1983). It is not known what biochemical defect causes the systemic disorder. In both cases, the muscle is infiltrated
with fat, and there is general weakness. The organic acidemias are inborn errors of metabolism in which fatty and organic acids accumulate,causing growth retardation, muscle hypotonia, protein intolerance, hyperammonemia, and ketoacidosis. Metabolic improvement has been ob652

CARNITINE

served with oral L-carnitine (Roe and Bohan, 1982; Seccombe et al.,1982).

Assessment of Status
Most carnitine status assessment relates to plasma or serum concentrations compared with normal values for a particular species. Assessment of the carnitine status in humans is difficult because plasma
carnitine concentrations and urinary carnitine excretion are not good indicators of tissue carnitine status (Borum, 1991). An individual with low
carnitine concentrations in plasma may have normal concentrations inmuscle or liver. It is also difficult to know how low a carnitine concentration must be before it is pathological.

SUPPLEMENTATION
Although carnitine has been studied in humans and under laboratory conditions for many years, its effectiveness in promoting the performance and well-being of domestic animals has only recently received
attention. A role for carnitine in swine and fish diets has been established, and continued research may find that it has a place in the production of poultry, horses, and ruminants. Some studies have found production responses for supplemental carnitine in poultry, cattle, andhorse diets, while other reports have found no benefits.
Supplementation of weanling pigs with carnitine appears to havecommercial potential. The major response may be improved growth ratewith lower-energy diets and improved feed efficiency with added-fat diets. Optimum dietary carnitine levels would appear to be between 800
and 1,200 mg/kg of feed for the first 1 or 2 weeks postweaning, with thelevel dropped to 50 to 500 mg/kg after that (Newton and Burtle, 1992).
Supplemental levels of carnitine for fish to accelerate growth and reduce
body fat have ranged from 500 to 2,000 mg/kg of feed.
Carnitine has been observed to be beneficial in the treatment of dilated cardiomyopathy in certain families of dogs (Keene et al., 1986,
1988), but ineffective in others (Costa and Labuc, 1994). A carnitinelevel of 50 mg/kg of body weight has been used as preventive therapy.
Working dogs have been shown to have a need for supplemental carnitine (Grandjean et al., 1993).
In the dog, maintenance requirements of endogenous L-carnitine appear sufficient, while in certain physiological states (exercise, exposure
to cold, reproduction) or deficiencies (myopathies, muscle damage), in
which mobilization of fatty acids is desirable, L-carnitine supplementa653

CHAPTER SIXTEEN

tion has been suggested at a level of 50 mg/kg of body weight (Pelletier,1992). On the basis of physiology tests (e.g., cardiac frequency, free fatty
acids, and lactic acid) after exercising Alaskan husky dogs, Grandjean etal. (1993) concluded that supplemental carnitine was beneficial.
Normal adult humans both synthesize and ingest carnitine inamounts totaling 100 mg/day. Under a variety of circumstances, however, carnitine deficiency may become manifest. Individuals particularly
vulnerable are newborn infants, premature infants, persons with inborn
metabolic defects, individuals receiving peritoneal dialysis or hemodialysis, alcoholics with liver disease, malnourished infants, and persons receiving total parenteral nutrition (Anonymous, 1985; Rebouche, 1986).
Hemodialysis patients can also be depleted of carnitine due to the lossof carnitine in the dialysate, which greatly exceeds the amount normallylost in the urine. This can be prevented if carnitine is added to thedialysate. Oral carnitine has also been found effective in correcting thehypertriglyceridemia that is frequent in uremic patients undergoing hemodialysis. The most economical and practical method of supplementing L-carnitine is ingestion of 660 to 990 mg/day or 2 to 3 g just prior
to hemodialysis (Golper and Ahmad, 1992). Hypocarnitinemia and tissue carnitine depletion are common in patients with advanced cirrhosis,
who not only tend to have marginal intakes of carnitine and its precursors, but also have loss of liver function, including the capacity to synthesize carnitine.
Some segments of the human population, such as vegetarians andThird World rural Asians, who consume diets high in grains and low inanimal products, were found to have slightly lower serum or plasma carnitine concentrations than their omnivorous counterparts, but no clinical abnormalities (Rebouche, 1986; Lombard et al., 1989). The need forsupplemental carnitine for this population group has yet to be established; however, if intake of lysine and methionine are inadequate, supplemental carnitine may be beneficial.
Acetyl-L-carnitine has been shown to be effective in slowing the
progression of mental deterioration in Alzheimer’s disease by improving
the ability of patients to handle tasks requiring attention and concentration (Spagnoli et al., 1991). In a double-blind, randomized, controlled
clinical trial, progression of Alzheimer’s disease was significantly reduced in patients who received acetyl-carnitine (2 g/day) for 1 year
(Bowman, 1992).
Dietary carnitine deficiency may be a cause of smooth muscle dysmotility of the gastrointestinal tract. After dietary supplementation ofcarnitine, gastrointestinal symptoms resolved, oesophageal manometry654

CARNITINE

returned to normal, and serum carnitine increased to 37.2 mmol/L(Weaver et al., 1992).
The consequences of suboptimal carnitine status would appear to begreat for the infant who, at birth, changes from a pattern of energy metabolism based on glucose as the major fuel to one based on utilizationof fats. Infants have compromised endogenous carnitine synthesis; theircarnitine status is dependent on that of the mother, on the placental
transfer of carnitine in utero, and on the availability of exogenous
sources after birth. However, milk or infant formula prepared fromcow’s milk contains appreciable amounts of carnitine, which is adequate
to meet daily requirements. Soybeans, on the other hand, lack carnitine,
hence infants fed an infant formula prepared from soy protein receive no
carnitine unless it is added by the manufacturers. Many infant formulas
contain low concentrations of carnitine, and supplemental carnitine isrecommended to be added to these formulas (Sugiyama et al., 1984).
Likewise, preterm infants have often been maintained on total parenteral nutrition containing no carnitine.

TOXICITY
Studies to determine maximum tolerance for carnitine in animals
are lacking. Carnitine supplementation at dosages that far exceed theusual dietary intake of carnitine have been administered in humans (Goaand Brogden, 1987). Oral dosages such as 100 mg/kg body weight per
day for infants and children with failure to thrive and 1 to 3 g of carnitine per day for adults with muscle weakness have been frequently used.
Some patients have experienced diarrhea, but not if they started withsmaller dosages and then increased gradually (Borum, 1991). Administration of the D-isomer may interfere with the normal functions of the Lisomer.

■ REFERENCES
Alkonyi, I., Cseko, J., and Sandor, A. (1990). J. Clin. Chem. Clin. Biochem. 28,
319.
Anonymous (1985). Nutr. Rev. 43, 23.
Baltzell, J.K., Bazer, F.W., Miguel, S.G., and Borum, P.R. (1987). J. Nutr. 117,
754.
Barker, D.L., and Sell, J.L. (1994). Poult. Sci. 73, 281.
Benamou, A.E., and Harris, R.C. (1993). Equine Vet. J. 25, 49.
Bohles, H., Segerer, H., and Fekl, W. (1984). JPEN 8, 9.
655

CHAPTER SIXTEEN

Bonner, C.M., DeBrie, K.L., Hug, G., Landrigan, E., and Taylor, B.J. (1995). J.
Pediatr. 126, 287.
Bonomi, A., Quantarelli, A., Sabbioni, A., Mazzali, I., Cabassi, E., Corradi, A.,
and Cantoni, A.M. (1991). Riv. Soc. Ital. Sci. Alim. 20, 401.
Borum, P.R. (1981). Nutr. Rev. 39, 385.
Borum, P.R. (1985). Can. J. Physiol. Pharmacol. 63, 571.
Borum, P.R. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 557. Marcel Dekker, New York.
Bowman, B.A.B. (1992). Nutr. Rev. 50, 142.
Bremer, J. (1983). Physiol. Rev. 63, 1420.
Bremer, J. (1990). J. Clin. Chem. Clin. Biochem. 28, 297.
Cantrell, C.R., and Borum, P.R. (1982). J. Biol. Chem. 257, 10599.
Carter, A.L., and Frenkel, R. (1978). J. Nutr. 108, 1748.
Chan, M.M. (1984). In Handbook of Vitamins (L.J. Machlin, ed.), p. 549. Marcel Dekker, New York.
Chapa, A.M., Fernandez, J.M, White, T.W., Bunting, L.D., Gentry, L.R., Ward,
T.L., and Blum, S.A. (1998). J. Anim. Sci. 76, 2930.
Cho, Y.O., and LeKlem, J.F. (1990). J. Nutr. 120, 258.
Coffey, M.T., Shireman, R.B., Herman, D.L., and Jones, E.E. (1991). J. Nutr.
121, 1047.
Corrigan, F.M., Rhijn, A.G., Best, P.V., Besson, J.A., Cooper, M.B., and Van
Rhijn, A.G. (1995) J. Nutr. Med. 5, 35.
Costa, N.D., and Labuc, R.H. (1994). J. Nutr. 124, 2687S.
Daily, J.W., and Sachan, D.S. (1995). J. Nutr. 125, 1938.
Davis, A. (1989). J. Nutr. 119, 262.
Deufel, T. (1990). J. Clin. Chem. Clin. Biochem. 28, 307.
DiMauro, G. (1979). In Handbook of Clinical Neurology (P.J. Vinken and G.W.
Bruyn, eds.), p. 271. Academic, New York.
Engel, A.G. (1980). In Carnitine Biosynthesis, Metabolism and Functions (R.A.
Frenkel and J.D. McGarry, eds.), p. 271. Academic Press, New York.
Feller, A.G., and Rudman, D. (1988). J. Nutr. 118, 541.
Flores, C.A., Hu, C., Edmond, J., and Koldovsky, O. (1996). J. Nutr. 126, 1673.
Freeman, J.M., Vining, E.P., Cost, S., and Singhi, P. (1994). Pediatrics 93, 893.
Friedman, S., and Fraenkel, G.S. (1972). In The Vitamins (W.H. Sebrell Jr. and
R.S. Harris, eds.), Vol. 5, 2nd Ed., p. 329. Academic Press, New York.
Goa, K.L., and Brogden, R.N. (1987). Drugs, 34, 1.
Golper, T.A., and Ahmad, S. (1992). Semin. Dialysis, 5, 94.
Grandjean, D., Valette, J.P., Jougln, M., Gabillard, C., Barque, H., Bene, M.,
and Guillaud, J.P. (1993). Recueil de Medecine Veterinaire, 169, 543.
Ha, T.Y., Otsuka, M., and Arakawa, N. (1991). J. Nutr. Sci. Vitaminol. 37, 371.
Ha, T.Y., Otsuka, M., and Arakawa, N. (1994). J. Nutr. 124, 732.
Helms, R.A., Mauer, E.C., Hay, W.W., Christensen, M.L., and Storm, M.C.
(1990). JPEN 14, 448.
Hill, G.M., Newton, G.L., and Mathis, M.J. (1995). The University of Georgia,
College of Agriculture, Department of Animal and Dairy Science 1995 Annual Report, p. 52.
Hoffman, L.A., Ivers, D.J., Ellersieck, M.R., and Veum, T.L. (1993). J. Anim.
Sci. 71, 132.
656

CARNITINE

Iben, C., Bergmeister, G., Sadila, E., and Leibetseder, J. (1992). In European
Conference on the Nutrition of the Horse. Physiology and Pathology of theDigestive Tract, p. 150. Hanover, Germany.
Indyk, H.E., and Woollard, D.C. (1995). J. Assoc. Off. Anal. Chem. Int. 78, 69.
Ji, H., Bradley, T.M., and Tremblay, G.C. (1996). J. Nutr. 126, 1937.
Keene, B.W. (1991). Vet. Clin. North Am. Small Anim. Pract. 21(5), 1005.
Keene, B.W. (1992). In Current Veterinary Therapy XI: Small Animal Practice(R.W. Kirk and J.D. Bonagura, eds.). W.B. Saunders, Philadelphia.
Keene, B.W. (1994). Abstract of 4th Annual ESVIM, p. 20. Congress, Brussels,
Belgium.
Keene, B.W., Atkins, C.E., Kittleson, M.D., Rush, J.E., and Shug, A.L. (1988).
ACVIM (Abstr.), p. 26. American College of Veterinary Internal Medicine,
Washington, D.C.
Keene, B.W., Kittleson, M.D., Rush, J.E., Plan, P.D., Atkins, C.E., DeLellis, L.D.,
Meurs, K.M., and Shug, A.L. (1989). J. Vet. Intern. Med. 3, 126.
Keene, B.W., Panciera, D.P., Atkins, C.E., Regitz, V., Schmidt, M.J., and Shug,
A.L. (1991). J. Am. Vet. Med. Assoc. 198(4), 647.
Keene, B.W., Panciera, D.L., Regitz, V., Noonan, J.J., Subramanian, R., andShug, A.L. (1986). ACVIM (Abstr.), p. 22. American College of Veterinary
Internal Medicine, Washington, D.C.
Kendler, E. (1986). Prev. Med. 15, 373.
La Count, D.W., Drackley, J.K., and Weigel, D.J. (1995). J. Dairy Sci. 78, 1824.
La Count, D.W., Emmert, L.S., and Drackley, J.K. (1996a). J. Dairy Sci. 79, 591.
La Count, D.W., Ruppert, L.D., and Drackley, J.K. (1996b). J. Dairy Sci. 79,
260.
Leibetseder, J. (1995). Arch. Anim. Nutr. 48, 97.
Lombard, K.A., Olson, A.L., Nelson, S.E., and Rebouche, C.J. (1989). Am. J.
Clin. Nutr. 50, 301.
McEntee, K., Clercx, C., Snaps, F., and Henroteaux, M. (1995). Canine Pract.
20(2), 12.
Melegh, B., Pap, M., Morava, E., Molnar, D., Dani, M., and Kurucz, J. (1994).
J. Pediatr. 125, 317.
Minkler, P.E., and Hoppel, C.L. (1993). J. Chromatogr. 613, 203.
Mitchell, M. (1978). Am. J. Clin. Nutr. 31, 293.
Monti, D., Troiano, L., Tropea, F., Grassilli, E., Cossarizza, A., Barozzi, D., Pelloni, M., Tamassia, M.G., Bellomo, G., and Franceschi, C. (1992). Am. J.
Clin. Nutr. 55 (6, Suppl.), 1208.
Musser, R.E., Goodbund, R.D., Tokack, M.D., Neissen, J.L., Owens, K.Q., andBlum, S.A. (1997). J. Anim. Sci. 75(Suppl. 1), 194.
Nakamura, T., Nakamura, S., Kondo, J., Ikeda, T., Ogata, T., Endo, F., andMatsuda, I. (1990). J. Pediatr. Gastroenterol. Nutr. 10, 66.
Nakano, C., Takashima, S., and Takeshita, K. (1989). Early Hum. Dev. 19, 21.
Newton, G.L., and Burtle, G.J. (1992). In Current Concepts in Carnitine Research (A.L. Carter, ed.), p. 59. CRC Press, Boca Raton, Florida.
Newton, G.L., and Haydon, K.D. (1989). J. Anim. Sci. 67(Suppl. 1), 267.
Novak, M., Monkus, E., Buch, M., Lesmes, H., and Silverio, J. (1983). Acta
Chir. Scand. 517(Suppl), 149.
Olson, A.L., and Rebouche, C.J. (1987). J. Nutr. 117, 1024.
657

CHAPTER SIXTEEN

Owen, K.Q., Nelssen, J.L., Goodband, R.D., Weeden, T.L., and Blum, S.A.
(1996) J. Anim. Sci. 74, 1612.
Pande, S.V., and Murthy, M.S. (1989). Biochem. Cell Biol. 67, 671.
Pelletier, B. (1992). Action Veterinaire 1210, 19.
Penn, D., and Schmidt-Sommerfeld, E. (1988). J. Nutr. 118, 1535.
Rebouche, C.J. (1986). Ann. Rev. Nutr. 61, 41.
Rebouche, C.J. (1991). Am. J. Clin. Nutr. 54(6 Suppl.), 1147S.
Rebouche, C.J. (1992). FASEB J. 6, 3379.
Rebouche, C.J., and Chenard, C.A. (1991). J. Nutr. 121, 539.
Rebouche, C.J., and Engel, A.G. (1983). Arch. Biochem. Biophys. 220, 60.
Rebouche, C.J., and Paulson, D.J. (1986). Ann. Rev. Nutr. 6, 41.
Rebouche, C.J., Panagides, D.D., and Nelson, S.E. (1990). Am. J. Clin. Nutr. 52,
820.
Reznic, A.Z., Kagan, V.E., Ramsey, R., Tsuchiya, M., Khwaja, S., Serbinova,
E.A., and Packer, L. (1992). Arch. Biochem. Biophys. 296, 394.
Roe, C.R., and Bohan, T.P. (1982). Lancet 1, 1411.
Rossle, C., Kohse, K.P., Franz, H.E., and Furst, P. (1985). Clinica Chimica Acta,
149, 263.
Sakemi, K., Hayasaka, K., Tahara, M., Sanada, Y., and Takada, G. (1992). Tohoku J. Exp. Med. 167, 89.
Santulli, A., and D’Amelio, V. (1986). Aquaculture 59, 177.
Santulli, A., Modica, A., Curatolo, A., and D’Amelio, V. (1988). Aquaculture
68, 345.
Schmidt-Sommerfeld, E., and Penn, D. (1990). Biol. Neonate 58(Suppl. 1), 81.
Schmidt-Sommerfeld, E., Penn, D., Rinaldo, P., Kossack, B.D., Li, B.U.K.,
Huang, Z.H., and Gage, D.A. (1992). Pediatr. Res. 31, 545.
Seccombe, D.W., Snyder, F., and Parsons, H.G. (1982). Lancet 2, 1401.
Shenai, J.P., and Borum, P.R. (1984). Pediatr. Res. 18, 679.
Smith, M.O., Cha, Y.S., and Sachan, D.S. (1994). Comp. Biochem. Physiol. 109,
177.
Spagnoli, A., Lucca, U., Menasce, G., and Bandera. L. (1991). Neurology 41,
1726.
Stumpf, D.A., Parker, W.D., and Angelini, C. (1985). Neurology 35, 1041.
Sugiyama, N., Suzuki, K., and Wada, Y. (1984). Jpn. Pediatr. Soc. 88, 1968.
Tanphaichitr, V., and Leelahagul, P. (1993). Nutrition 9, 246.
Torreele, E., Van Der Sluisgen, A., and Verreth, J. (1993). Br. J. Nutr. 69, 289.
Tuan, S., and Chou, C.C. (1995). J. Chi. Nutr. Soc. 20, 73.
Van Kempen, T., and Odle, J. (1993). J. Nutr. 123, 1531.
Van Kempen, T., and Odle, J. (1995). J. Nutr. 125, 238.
Waber, I.J., Valle, D., Neill, C., Dimauro, S., and Shug, A. (1982). J. Pediatr.
191(5), 700.
Weaver, L.T., Rosenthal, S.R., Gladstone, W., and Winter, H.S. (1992). Acta Pediatr. 81, 79.

658

CHAPTER SEVENTEENVITAMIN-LIKE SUBSTANCESINTRODUCTION
In addition to the 15 vitamins discussed in previous chapters, othersubstances have been classed with the vitamins although their true vitamin character has not been established. For various reasons, the term vitamin has been applied to many substances that do not meet criteria forvitamin status. Vitamins \are regarded as essential organic micronutrientsthat must be supplied in the diet. What is a vitamin for some species may
only be an essential metabolite for others, as dietary sources are notneeded because of tissue synthesis (e.g., vitamin C). myo-Inositol fits thiscategory, but apparently for only a few species, including fish, gerbils,
and perhaps other species.
Many compounds described as vitamins in the scientific literature of
the 1930s and 1940s have since proven to be identical to other essential
nutrients or to be mixtures of various compounds. Many of the substances referred to as vitamins, growth factors, or accessory factors inearly research literature are no longer considered to be vitamins (Cody,
1991). Some compounds, such as myo-inositol, lipoic acid, coenzyme Q,
and polyphenols, exhibit biological activity without being dietary essentials (or vitamins) for most species. The term vitamin denotes dietaryneed. Pyrroloquinoline quinone is a potential new vitamin, with additional evidence needed for its establishment as a vitamin. Another groupof substances is called vitamins by potential promoters for profit; thesesubstances, including pangamate, laetrile, gerovital, and cabagin, are notdietary essentials and are more properly called pseudovitamins (Cody,
1991). The present chapter will emphasize myo-inositol since this substance is a vitamin to some species, with only brief mention allotted toother vitamin-like substances.
659

CHAPTER SEVENTEEN

MYO-INOSITOL (INOSITOL)
Introduction
myo-Inositol, also referred to as inositol, is a water-soluble growth
factor for which no coenzyme function is known. It was first isolated
from muscle in 1850 and was identified as a growth factor for yeast andmolds, though not for bacteria. myo-Inositol deficiency in mice, characterized by inadequate growth, alopecia, and death, was reported. However, these results were challenged as the diets were ill defined and wereapparently deficient in some of the B-complex vitamins (Kukis andMookerjea, 1978). Most evidence suggests that myo-inositol is not atrue vitamin for most species. Nevertheless, signs of myo-inositol deficiency have been demonstrated in fish and gerbils (Kroes, 1978). Difficulty in demonstrating deficiency is related to endogenous synthesis,
highly variable turnover rates, and interactions of myo-inositol with certain vitamins or other nutrients.

Chemical Structure and Properties
Inositol exists in nine forms. It is a cyclohexane compound, with
only myo-inositol (Fig. 17.1) demonstrating any biological activity.
myo-Inositol is an alcohol, similar to a hexose sugar. It is a white, crystalline, water-soluble compound with a sweet taste, and is stable inacids, alkalines, and heat up to about 250oC. Because of hydroxyl
groups, it forms various ester, ethers, and acetals. The hexaphosphoricacid ester (combined with six phosphate molecules) of myo-inositol isphytic acid, a compound that complexes with phosphorus and otherminerals, making them less available for absorption (Fig. 17.1). Methods of analysis include the traditional microbiological method, which is
being replaced by more rapid methods of gas-liquid, paper, and thinlayer chromatography.

Metabolism and Functions
Ingested myo-inositol is absorbed by rats and humans at a rate of over
99%, with myo-inositol from phytate absorbed at less than 50% in thepresence of high dietary calcium (Cody, 1991). myo-Inositol is absorbedby active transport from dietary sources, or it may be synthesized de novo
from glucose. Based on animal studies, myo-inositol may also be converted to glucose. myo-Inositol appears to have three metabolic fates: oxidation to CO2, use in gluconeogenesis, and synthesis of phospholipids.
The function of myo-inositol is not completely understood, al660

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Fig. 17.1 Chemical structures of myo-inositol, phytic acid, and phytic acidcombined with zinc.

though its biochemical functions probably relate to its role as a phospholipid component of membranes and lipoproteins. Assuming that
myo-inositol is bound as lipositol, it makes up over 25% of total lipid
and almost 50% of phospholipid in microsomes of the liver cell. myoinositol is a structural component of phosphoinositides, with many ofthe possible functions associated with its role as a membrane component. In addition to the general role of maintaining selective permeability of plasma membranes, phosphatidylinositol and its highly charged
phosphorylated forms are hypothesized to regulate cell-surface phenomena, such as binding of hormones and transfer of nervous impulses(Michell, 1975). Inositol 1,4,5-triphosphate has been shown to releaseintracellular Ca2+ from the endoplasmic reticulum, providing a link between receptor activation and cellular Ca2+ mobilization (Putney et al.,1986). Examples of cellular processes controlled by the phosphoinositides include amylase secretion, insulin release, smooth muscle contraction, liver glycogenolysis, platelet aggregation, histamine secretion, andDNA synthesis in fibroblasts and lymphoblasts. Under certain conditions, myo-inositol is lipotropic, with the lipotropic activity usually synergistic with that of choline. myo-Inositol reduces liver lipids for diets
low in protein and fat, while choline is lipotropic also for diets containing lipids.

Sources, Requirements, and Deficiency
All plants and animals contain measurable amounts of myo-inositol(Clements and Darnell, 1980; Cody, 1991). The most concentrated dietary sources of myo-inositol are foods that consist of seeds, such as661

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beans, grains, and nuts; cantaloupe and citrus fruits are also good plantsources. Most myo-inositol from plant sources occurs bound to phosphate as phytate. The best animal sources are the organ meats, whichcontain myo-inositol in free form or as a component of phospholipids(primarily phosphatidyl inositol). High concentrations are found inheart, kidney, liver, spleen, and thyroid.
There are no known dietary requirements for myo-inositol in humans or most species of animals studied. This is most likely due to theavailability of myo-inositol from dietary sources, from endogenous synthesis, and from bacterial synthesis. Early work suggested that myo-inositol was an essential nutrient for several animal species. Effects of
myo-inositol deficiency included alopecia in rats and mice; fatty liverand “spectacle eye” in rats; and retarded growth in guinea pigs, hamsters, and chickens. Subsequent work using more complete and defineddiets with these species failed to show any myo-inositol requirement.
Early reports of deficiency were in fact deficiencies of biotin, choline, vitamin E, or other nutrients.
For ruminants, synthesis of myo-inositol by microorganisms in thedigestive tract, in addition to dietary sources, is presumably sufficient tomeet the animals’ needs. Since myo-inositol deficiency has been associated with failure of lipid transport in a number of species, it was suggested that myo-inositol supplementation might be beneficial in alleviating fatty liver syndrome (hepatic lipidoses) in dairy cattle. However,
myo-inositol supplementation (17 g daily) to dairy cows was ineffective
in altering liver myo-inositol concentrations or in improving liver
lipoprotein output (Gerloff et al., 1986).
From a review of the myo-inositol literature, Kroes (1978) reported
that there is little or inconclusive evidence that myo-inositol is requiredfor cats, poultry, ruminants, dogs, guinea pigs, hamsters, mice, pigs, orrats. For rats and other species, however, reports show relative requirements for myo-inositol when diets are deficient in single other nutrients,
such as choline, or in multiple vitamin and mineral deficiencies (Yagi etal., 1965). The deficiency signs were lipid accumulation for simultaneous choline deficiency and depressed growth if vitamin and mineral imbalance existed.
Under typical feeding conditions, most evidence indicates that pigs
and poultry have no requirement for dietary myo-inositol. Lindley andCunha (1946) demonstrated that addition of 100 mg of myo-inositol per
100 g of a diet deficient in biotin and supplemented with sulfathalidine
could partially relieve signs of biotin deficiency. Presumably, the added
myo-inositol stimulated intestinal synthesis of biotin. In some studies,662

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myo-inositol has promoted growth in chicks, decreased liver fat, and increased egg production. These results have been refuted by other investigators (Kroes, 1978).
Animal species that have shown a dietary need for myo-inositol arefish and gerbils. In fish, deficiency of myo-inositol results in anorexia,
dark skin coloration, fin degeneration, edema, anemia, reduced growth,and inefficient feed conversion, as well as a decreased rate of gastric
emptying and activity of cholinesterase and certain transaminases(Halver, 1982). Decreased cholinesterase and certain aminotransferase
activities were observed in trout, red sea bream, Japanese eel, Japaneseparrot fish, and yellowtail with myo-inositol deficiency (NRC, 1993).
Rainbow trout fed a diet deficient in myo-inositol had large accumulations of neutral lipids in the liver and increased levels of cholesterol andtriglycerides, but decreased amounts of total phospholipid, phosphotidylcholine, phosphotidylethanolamine, and phosphotidylinositol(Holub et al., 1982).
myo-Inositol is synthesized in common carp intestine (Aoe and Masuda, 1967), but not in amounts sufficient to sustain normal growth ofyoung fish without dietary source of this vitamin, because younger carp
require a higher level of myo-inositol than older fish. Burtle and Lovell(1989) demonstrated de novo synthesis of myo-inositol in the liver ofchannel catfish, as well as intestinal synthesis. A tentative recommended
allowance for young salmon and trout is 350 to 500 mg/kg of feed
(McLaren et al., 1947).
Female gerbils with myo-inositol deficiency were characterized withintestinal lipodystrophy, with a resulting hypocholesterolemia, debilitation, and eventual death (Hegsted et al., 1973). Male gerbils were apparently protected by testicular synthesis of myo-inositol. Requirementof myo-inositol for female gerbils varied from 20 mg/kg of diet when thediet contained predominantly unsaturated fats (20% safflower oil) to
120 mg/kg of diet with saturated fats (20% coconut oil) (Kroes et al.,1973).
A number of factors affect myo-inositol status, and it is apparent
that under certain conditions, a need for myo-inositol can be shown forvarious species. The reasons for the need under those conditions is notexactly known; however, administration of antibiotics, dietary stress,
and physiological stress may influence the need for myo-inositol(Cunha, 1971; Cody, 1991). Antibiotics kill myo-inositol-producing intestinal flora, thus reducing the exogenous supply of myo-inositol to thebody. Increasing levels of dietary saturated fatty acids may stress myoinositol-requiring lipid transport systems. For humans, impaired myo663

CHAPTER SEVENTEEN

inositol metabolism occurs in diabetics, uremic individuals, and premature infants (Cody, 1991).

Detrimental Effects of myo-Inositol as Phytates
myo-Inositol is of greatest significance to livestock and human nutrition because of its hexaphosphate ester phytic acid (Fig. 17.1). Phyticacid, which is formed from six phosphate molecules combined with
myo-inositol, hinders intestinal absorption of phosphorus, calcium, andother minerals, including zinc, copper, cobalt, iron, manganese, andmagnesium. In grains and plant protein supplements, about two-thirdsof the phosphorus is in the less available phytate form. Utilization ofphytate phosphorus is influenced by phytase present in plant materials
or synthesized by rumen microflora, and by intake of vitamin D, calcium, and zinc, as well as by such factors as alimentary tract pH and dietary ratio of calcium to phosphorus. Calcium exaggerates the inhibitionof zinc absorption by phytate, while vitamin D needs are higher to counteract high phytate intakes.
Phytin is especially high in bran of cereal grains and oilseed meals.
About 20 to 50% of phytin phosphorus is available to the pig. A good
guide is to assume that no more than about 50% of the phosphorus inplant feeds is available to the pig (Cunha, 1977). There is disagreement
concerning the ability of poultry to utilize phytin phosphorus (NRC,
1994). Most data, however, indicate that the utilization of phytin phosphorus by young or adult poultry is negligible if dietary calcium concentrations are sufficient to meet the birds’ requirements. Some reportssuggest, however, that the older bird has ability to use most of the phytin
phosphorus. Many cereal grains contain the enzyme phytase, which iscapable of splitting phosphorus from myo-inositol and leaving the phosphorus, calcium, and other minerals attached to it available for absorption. Rye in particular, and also wheat, contain enough phytase to leadto considerable destruction of phytic acid. Thus, although 50% or moreof the phosphorus in the original whole grain may be in the form ofphytic acid, the amount in the final product may be very much less. At
the other extreme, oats contain little phytase. Phytin phosphorus may bealmost totally unavailable to the pig unless the phytase of grains or othersources is present in the diet. Ruminants utilize phytin phosphorus quitesatisfactorily because of consumption of dietary phytases and phytase
production by rumen microorganisms (Morse et al., 1992).
Low availability of phytate phosphorus poses two problems: (1) theneed to add inorganic phosphorus supplements to diets, and (2) the excretion of large amounts of phosphorus in manure. Land application of664

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manure has been the primary method of disposal of livestock and poultry waste due to its fertilizing value. There is a potential environmental
impact from excessive phosphorus buildup in soil, with surface runoff
and erosion of soil phosphorus into rivers, lakes, and streams. Excess
phosphorus causes algae and other aquatic plants to have unchecked
growth, which reduces and/or changes the oxygen and carbon dioxidelevels in water and causes other aquatic life to suffer. Because of this,many state and local governments have enacted or are enacting legislation to reduce phosphorus pollution. Providing phytases to monogastricdiets makes plant sources of phosphorus more available, with less inorganic phosphorus required and less of the mineral in feces to be detrimental to the environment.
Great strides have been made to develop microbial phytases tobreak down phytate in natural feed sources. The addition of microbial
phytase produced from a genetically modified Aspergillus niger strain tohigh phytate corn-soybean meal diets fed to poultry (Denbow et al.,
1995; Yi et al., 1996) and swine (Simons et al., 1990; Cromwell et al.,
1993; Lei et al., 1993) have been beneficial. Some studies have shown
the bioavailability of phytate phosphorus to be improved by 20 to 30%.
Lei et al. (1993) reported that supplemental microbial phytase improved
phytate phosphorus utilization by pigs and reduced fecal phosphorus excretion. Cereal phytase from wheat bran fed to pigs from weaning
through finishing was shown to be almost as effective as microbial phytase in improving phytate phosphorus utilization for body weight gain,
but not for bone mineralization (Han et al., 1997).
Many human nutritionists feel that the two minerals most likely to
be deficient in typical diets are calcium and iron, both of which are affected by plant phytates. In human nutrition, the amount of calcium absorbed is dependent on bread type. Calcium is much less freely absorbed
from diets consisting largely of brown bread than from those consistinglargely of white, because whole wheat flour contains much higher levelsof phytate. Inhibitory effects of soybean-derived food products and cereal grains on iron absorption as a result of phytates have caused concern among nutritionists and food scientists (Morris, 1983). For pigs,
phytase was shown to improve iron bioavailability from soybean mealfor hemoglobin synthesis (Stahl et al., 1999).
The high use of soy protein or soybean meal in human and monogastric diets has led to zinc deficiency. During extraction of soy protein,phytic acid forms a complex with zinc to form zinc phytate (Fig. 17.1),
which is insoluble in the intestinal tract. The zinc requirement of growing pigs receiving semipurified diets containing isolated soybean protein
665

CHAPTER SEVENTEEN

or natural corn-soybean meal diets containing the recommended calcium level is about 50 ppm. However, in the absence of plant phytates,
pigs receiving a casein-glucose diet require only 15 ppm zinc (NRC,1988). Higher levels of calcium are known to further exaggerate the inhibition of zinc absorption by phytate, resulting in the formation of zinccalcium-phytate complexes (Forbes et al., 1983).

PYRROLOQUINOLINE QUINONE (PQQ)
In 1979 studies of specialized bacteria, the methylotrophs, resulted
in the discovery of a new enzyme cofactor, pyrroloquinoline quinone(PQQ) (Salisbury et al., 1979). Structural analysis showed that PQQ
had a molecular weight of 424 and an empirical formula of
C17H12N2O9 . 2H2O.
Pyrroloquinoline quinone has been identified in several other important enzymes (now collectively called quinoproteins) in yeasts,
plants, and animals. However, three laboratories were unable to confirm
PQQ as a cofactor in three mammalian enzymes previously shown tocontain it (Harris, 1992). Nevertheless, PQQ is reported as part of the
enzyme lysyl oxidase (Williamson et al., 1986).
Pyrroloquinoline quinone functions as the redox center in thequinoprotein enzymes (Bishop et al., 1998) The redox behavior of PQQ
involves its ability to facilitate both one- and two-electron transfers.
There is substrate oxidation by a two-electron transfer to PQQ, followed by single-electron transfer to acceptors such as copper-containing
proteins and cytochromes. The two forms of the cofactor—the semiquinone PQQH and the catechole PQQH2 (Fig. 17.2)—have been foundin the bacterial quinoproteins.
Pyrroloquinoline quinone and copper are cofactors for lysyl oxidase, an enzyme required for cross-linking of collagen and elastin (Killgore et al., 1989). Mice deficient in PQQ showed friable skin, mildalopecia, and hunched posture. About one-fifth of the PQQ-deprived
mice died within 8 weeks of feeding, with aortic aneurysms or abdominal hemorrhages. Pregnant mice fed chemically defined diets apparently
devoid of PQQ exhibited reproductive failure.
Little information is available on dietary sources of PQQ. Some reports indicate PQQ to be present in egg yolk, adrenal tissue, and many
citrus fruits in the range of 500 to 20,000 ppb. It is ubiquitous in dietarycomponents, including water, and may be produced in mammalian cells
at less than optimal levels, making it difficult to determine if it is an es-

666

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Fig. 17.2 Chemical structures off the semiquinone PQQH, the catecholesential nutrient. Further studies with PQQ may result in its establishment as a vitamin.

p-AMINOBENZOIC ACID (PABA)
p-Aminobenzoic acid (PABA) (Fig. 17.2) was originally identified as a
growth factor for many species of bacteria and as a required nutrient for
lactation in rats and growth for the chick. The only role of PABA in higher
animals would appear to be as part of the folacin molecule (see Chapter
12). When sufficient dietary folacin is available, there is little evidence that667

CHAPTER SEVENTEENPABA plays a direct role in the nutrition of higher animals or humans, andtherefore it cannot be classified as a vitamin. If folacin is lacking, PABA
may have its main effect by providing a building block for intestinal synthesis of the vitamin. One of the better recognized properties of PABA is
its ability to counteract the bacteriostatic effects of sulfonamides. The
chemical structure of PABA is very similar to that of some sulfonamides,
which explains why it can counteract inhibition of microbial growth bythese drugs. Therefore, ingestion of PABA can worsen infection. Of interest, PABA is the protective component in sunscreen.

FLAVONOIDS (POLYPHENOLS)
In 1936, Rusznyak and Szent-Györgyi reported the presence of a
substance in citrus fruits, different from vitamin C, that is essential toprevent fragility of capillaries. The substance was designated as vitamin
P. Several reports have shown that quercetin, catechol, rutin (Fig. 17.2),
hesperidin, chalcone, and other nonspecific polyphenols, or flavonoids,
can provide some protection against capillary fragility under certain
conditions. This effect, which would appear to spare vitamin C, may be
due to the ability of flavonoids to chelate divalent metal cations (e.g.,
Cu++, Fe++), thus performing antioxidant functions. Some of these compounds may have value as a supplement to limited vitamin C intake,particularly under conditions of stress (Maynard et al., 1979).
Flavonoids are colored phenolic substances found in all higher plants;
more than 3,000 different flavonoids have been isolated (Cody, 1991).
They are the major sources of red, blue, and yellow pigments (except for
carotenoids) in the plant kingdom. Flavonoids are ubiquitous throughout
the plant kingdom, with higher flavonoid concentrations found in coloredexterior tissues, such as peels and skins, than in interior tissues (Herrmann, 1976). They are widely distributed in fruits and vegetables, and inbeverages such as tea and wine (Cao et al., 1998). The greatest source offlavonoids in the diet is fruit juice, with orange juice supplying approximately 22 mg of hesperidin per capita per day and grapefruit juice supplying approximately 5.6 mg of naringin per capita per day (Cody, 1991).
Hertog et al. (1993b) reported in a food-consumption survey in the
Netherlands that the most important flavonoid was quercetin (mean intake 16 mg/day). Important sources of flavonoids were tea (48% of total
intake), onions (29%), and apples (7%). On a milligram-per-day basis, theintake of the antioxidant flavonoids exceeded that of the antioxidants βcarotene and vitamin E. Thus flavonoids represent an important source ofantioxidants in the human diet.
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In addition to reducing capillary fragility, numerous other biologicaleffects of flavonoids have been claimed (Vlietinck et al., 1988; Bravo,
1998; Yang et al., 1998). They are believed to lower blood cholesterol
and to prevent cancer, hypertension, viral infection, and allergic reactions. Other potentially important reports of the flavonoid activities include the in vitro inhibition of aldose reductase, which converts glucose
and galactose to their polyols (Varma and Kinoshita, 1976). These metabolic products have been implicated in the neuropathy of diabetes andin cataract formation.
Although a number of polyphenols exhibit biological activities, including reduction of capillary fragility and protection of biologically important compounds through antioxidant activity, none of the polyphenols has been demonstrated to be essential or to be capable of causing
deficiency signs when removed from the diet. However, epidemiological
data from the Netherlands (Hertog et al., 1993a) indicated a reduced
risk of coronary heart disease in men associated with increased ingestionof flavonoids. Relative risk of mortality from coronary heart disease andincidence of a first myocardial infarction were about 50% lower for thehigher flavonoid intake groups. Red wine is a significant source offlavonoids in the diet of some cultures, and its antioxidant properties
may contribute to a possible explanation of reduced risk of coronary
heart disease in wine drinkers—the so-called French paradox (Frankel etal., 1993).

LIPOIC ACID (THIOCTIC ACID)
Lipoic acid (thioctic acid) (Fig. 17.2) plays an important role in thegrowth of certain microorganisms. It also is essential in oxidative decarboxylations of α-keto acids, such as pyruvic acid, in carbohydrate metabolism. There is, however, no clear evidence for an established need inanimal nutrition that enables it to be classed as a vitamin, despite several experiments with rats and chicks (Maynard et al., 1979).

COENZYME Q (UBIQUINONES)Coenzyme Q is a collective name for a number of ubiquinones, suchas Q4 and Q10, that play an established role in the respiratory chain in
mitochondrial systems. It is an electron carrier between flavoproteinsand cytochromes. In addition, the coenzyme appears to have beneficial
effects in certain disease states, including muscular dystrophy, periodontal disease, hypertension, and congestive heart failure (Basu and Dicker669

CHAPTER SEVENTEEN

son, 1996). The importance of coenzyme Q as a ubiquitous catalyst for
respiration ensures its status as an essential metabolite. There is evidence
that specific ubiquinones have a sparing effect on vitamin E, resulting inremission of some clinical signs of the vitamin deficiency. Dietaryubiquinone seems on the whole to be unimportant unless it provides thearomatic nucleus for endogenous synthesis. There is no proof that justifies classification of coenzyme Q as a separate vitamin.

VITAMIN B13 (OROTIC ACID)
Vitamin B13 (orotic acid) was isolated from distiller’s solubles, withthe purified compound orotic acid, which is an intermediate in pyrimidine metabolism. It has been found to stimulate the growth of rats,
chicks, and pigs under certain conditions, but evidence remains uncertain whether it plays an essential role in an otherwise adequate diet
(Cody, 1991). It has no known coenzyme function.

VITAMIN B15 (PANGAMIC ACID)
Vitamin B15 (pangamic acid) is found in rice bran, yeast, blood
meal, and other feeds. It is not a chemically defined substance, and there
is no evidence that pangamic acid preparations have vitamin activity oroffer therapeutic benefit.

VITAMIN B17 (LAETRILE)
Vitamin B17, or laetrile (amygdalin), is a β-cyanogenic glucoside occurring naturally in the kernels or seeds of most fruits (e.g., apricot). Although many unsupported claims have been made for the therapeutic
benefit of laetrile treatment, most publicized claims are for its use intreating cancer. In these claims, the two major lines of argument advanced are that (1) the cyanide in laetrile acts specifically to destroy cancer cells and (2) cancer is a nutritional deficiency disease requiring
laetrile treatment for dietary control (Cody, 1991). In a clinical study involving 178 terminally ill cancer patients conducted at four U.S. cancer
centers, no differences were observed in terms of cure, disease progression, improvement of symptoms, or extension of life span in patientswith or without laetrile therapy (Moertel et al., 1982). Most nutritionists do not consider laetrile a vitamin.

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VITAMIN H3 (GEROVITAL)
Vitamin H3 (gerovital) is a buffered solution of procaine hydrochloride better known as novocaine, which is used as a pain killer by dentists. It is promoted as a nutritional substance that alleviates symptoms
of diseases associated with aging. These claims have not been supported
by scientific studies, and it is therefore not recognized as a vitamin.

VITAMIN U (CABAGIN)
Vitamin U (cabagin) is claimed to be an antiulcer factor and occurs
naturally in cabbage and other green vegetables. The actual active substance is a methylsulfonium salt of methionine. The claims as an antiulcer factor have not been supported by some studies.

GLUCOSE TOLERANCE FACTOR
Glucose tolerance factor contains the element chromium, which has
been reported to be involved in maintaining normal serum cholesterol
and in regulating glucose metabolism (Williams and McDowell, 1985).
The glucose tolerance factor qualifies as a vitamin since it contains
chromium, organic components of nicotinic acid, glycine, glutamic acid,and cysteine and has much greater biological activity than inorganic
sources of chromium alone. This would be comparable to vitamin B12being more metabolically effective than the element cobalt.

OTHER VITAMIN-LIKE FACTORSIn 1972, Cheldelin and Baich listed unidentified growth factors(UGFs) and the organisms utilizing them. A total of 255 references werenoted. In particular, many references involved unidentified factors present in fish products, alfalfa meal, liver, and whey. Thus, we have suchterms as whey factor, fish solubles factor, grass juice factor, and manyothers.
With the discovery of folacin in the early 1940s, many believed that
all of the unidentified factors had been discovered and that no more factors were needed for optimum nutrition. One reason for this supposition
was the fact that completely synthetic diets of known nutrient composition, containing all recognized vitamins and mineral nutrients and ade-671

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quate in the essential amino acids, would support growth and development in young weanling rats (Scott et al., 1982). However, in 1948 thediscovery of vitamin B12 as the unknown activity—termed the animal
protein factor—responsible for special growth-promoting effects was an
excellent example of the fallacy of assuming that no more factors exist
simply because animals can survive on synthetic diets.
With the discovery of vitamin B12 in 1948, the period of active identification and isolation of the major vitamins appeared to be ending.
However, even since 1948 many field reports have suggested that practical diets containing sources of UGF are superior to purified or commercial diets. Typically, the unidentified factors found in certain feed ingredients have not been isolated and identified. These factors could be
providing vitamins, trace minerals, or a better amino acid balance, orcounteracting antagonists in the regular diet. In poultry, many of thegrowth responses obtained from UGF involved relationships between
known nutrients, such as natural chelates in corn distiller’s dried solubles that improved zinc utilization in a purified diet containing soybean(Scott et al., 1982).
Fish solubles, dried whey, brewer’s dried yeast, corn distiller’s driedsolubles, and other fermentation residues are the major special ingredients often added to poultry diets as potential sources of unrecognized
nutritional factors. In swine studies, Cunha (1977) reported that highquality alfalfa meal and pasture, animal protein concentrates, liver, soil,
dried distiller’s solubles, fish solubles, grass juice concentrate, dried
whey, and other feeds have been shown to contain a factor or factors
useful either for the growing pig or for the sow during gestation and lactation.
The use of short, lush, green leafy pastures will minimize vitamin
deficiencies in swine (Cunha, 1977). Likewise, most nutritionists recognize the possible benefits of using some UGF supplementation to ensure
optimal performance of diets for broilers and breeding hens (Scott et al.,1982). This practice, in addition to pasture use, may have a twofold advantage in providing possible unidentified growth factor responses and,at the same time, supplying additional amounts of some of the known
vitamins, as UGF supplements usually are good sources of many vitamins. The additional vitamins in the diet may prevent serious losses
when there is a loss of potency or omission of an important vitaminfrom the vitamin premix (Scott et al., 1982).

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Aoe, H., and Masuda, I. (1967). Bull. Jpn. Soc. Sci. Fish, 33, 674.
Basu, T.K., and Dickerson, J.W.T. (1996). In Vitamins in Human Health andDisease, p. 249. CAB International, Wallingford, United Kingdom.
Bishop, A., Gallop, P.M., and Karnovsky, M.L. (1998). Nutr. Rev. 56, 287.
Bravo, L. (1998). Nutr. Rev. 56, 317.
Burtle, G.J., and Lovell, R.T. (1989). Can. J. Fish. Aquat. Sci. 46, 218.
Cao, G., Russell, R.M., Lischner, N., and Prior, R.L. (1998). J. Nutr. 128, 2383.
Cheldelin, V.H., and Baich, A. (1972) In The Vitamins (W.H. Sebrell Jr. and R.S.
Harris, eds.), Vol. 5, 2nd Ed., p. 398. Academic Press, New York.
Clements, R.S., and Darnell, B. (1980). Am. J. Clin. Nutr. 33, 1954.
Cody, M.M. (1991). In Handbook of Vitamins (L.J. Machlin, ed.), p. 565. Marcel Dekker, New York.
Cromwell, G.L., Stahly, T.S., Coffey, R.D., Monegue, H.J., and Randolph, J.H.
(1993). J. Anim. Sci. 71, 1831.
Cunha, T.J. (1971). In The Vitamins (W.H. Sebrell Jr. and R.S. Harris, eds.), Vol.
3, 2nd Ed., p. 394. Academic Press, New York.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Denbow, D.M., Ravindran, V., Kornegay, E.T., Yi, Z., and Hulet, R.M. (1995).
Poult. Sci. 74, 1831.
Forbes, R.M., Erdman, J.W., Parker, H.M., Kondo, H., and Ketelsen, S.M.
(1983). J. Nutr. 113, 205.
Frankel, E.N., Kanner, J., German, J.B., Parks, E., and Kinsella, J.E. (1993).
Lancet 341, 454.
Gerloff, B.J., Herdt, T.H., Wells, W.W., Liesman, J.S., and Emery, R.S. (1986). J.
Anim. Sci. 62, 1682.
Halver, J.E. (1982). Comp. Biochem. Physiol. 73, 43.
Han, Y.M., Yang, F., Zhou, A.G., Miller, E.R., Ku, P.K., Hogberg, M.G., andLei, X.G. (1997). J. Anim. Sci. 75, 1017.
Harris, E.D. (1992). Nutr. Rev. 50, 263.
Hegsted, D.M., Hayes, K.C., Gallagher, A., and Hanford, H. (1973). J. Nutr.
103, 302.
Herrmann, K. (1976). J. Food Techol. 11, 433.
Hertog, M.G.L., Feskens, E.J.M., Hollman, P.C.H., Katan, M.B., andKromhout, D. (1993a). Lancet. 342, 1007.
Hertog, M.G.L., Hollman, P.C.H., Katan, M.B., and Kromhout, D. (1993b).
Nutr. Cancer 20, 21.
Holub, B.J., Bregeron, B., and Woodward, T. (1982). J. Nutr. 113(6), xxi(Abstr.).
Killgore, J., Smidt, C., Duich, L., Romero-Chapman, N., Tinker, D., Reiser, K.,
Melko, M., Hyde, D., and Rucker, R. (1989). Science 245, 850.
Kroes, J. (1978). In Handbook Series in Nutrition and Food, Section E: Nutritional Disorders (M. Rechcigl Jr., ed.), Vol. 2, p. 143. CRC Press, Boca Raton, Florida.
Kroes, J.F., Hegsted, D.M., and Hayes, K.C. (1973). J. Nutr. 103, 1448.
Kukis, A., and Mookerjea, S. (1978). Nutr. Rev. 36, 233.
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CHAPTER SEVENTEEN

Lei, X.G., Ku, P.K., Miller, E.R., and Yokoyama, M.T. (1993). J. Anim. Sci. 71,
3359.
Lindley, D.C., and Cunha, T.J. (1946). J. Nutr. 32, 47.
Maynard, L.A., Loosli, J.K., Hintz, H.F., and Warner, R.G. (1979). Animal Nutrition, 7th Ed, p. 283. McGraw-Hill, New York.
McLaren, B.A., Keller, E., Donnell, D.J., and Elvehjem, C.A. (1947). Arch.
Biochem. Biophys. 15, 169.
Michell, R.H. (1975). Biochem. Biophys. Acta 415, 81.
Moertel, C.G., Fleming, T.R., Rubin, J., Kvols, L.K., Sarna, G., Koch, R., Currie, V.E., Young, C.W., Jones, S.E., and Davingnon, J.P. (1982). N. Engl. J.
Med. 306, 201.
Morris, E.R. (1983). Fed. Proc. Fed. Am. Soc. Exp. Biol. 42, 1716.
Morse, D., Head, H.H., and Wilcox, C.J. (1992). J. Dairy Sci. 75, 1979.
NRC. (1988). Nutrient Requirements of Domestic Animals: Nutrition Requirements of Swine, 9th Ed. National Academy of Sciences-National ResearchCouncil, Washington, D.C.
NRC. (1993). Nutrient Requirements of Domestic Animals: Nutrition Requirements of Fish. National Academy of Sciences-National Research Council,
Washington, D.C.
NRC. (1994). Nutrient Requirements of Domestic Animals: Nutrition Requirements of Poultry, 9th Ed. National Academy of Sciences-National ResearchCouncil, Washington, D.C.
Putney, J.W., Aub, D.L., Taylor, C.W., and Merritt, J.E. (1986). Fed. Proc. Fed.
Am. Soc. Exp. Biol. 45, 263.
Rusznyak, S., and Szent-Györgyi, A. (1936). Nature (London) 138, 27(Abstr.).
Salisbury, S.A., Forrest, H.S., Cruse, W.B.T., and Kennard, O. (1979). Nature
280, 843.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Simons, P.C.M., Versteegh, H.A.J., Jongbloed, A.W., Kemme, P.A., Slump, P.,
Bos, K.D., Wolters, M.G.E., Beudeker, R.F., and Verschoor, G.J. (1990). Br.
J. Nutr. 64, 525.
Stahl, C.H., Han, Y.M., Roneker, K.R., House, W.A., and Lei, X.G. (1999). J.
Anim. Sci. 77, 2135.
Varma, S.D., and Kinoshita, J.H. (1976). Biochem. Pharm. 25, 2505.
Vlietinck, A.J., Berghe, D.A.V., and Haemers, A. (1988). In Plant Flavonoids in
Biology and Medicine II (V. Cody, E. Middeton Jr., J.B. Harborne, and A.
Beretz, eds.), p. 283. Alan R. Liss, New York.
Williams, S.N., and McDowell, L.R. (1985). In Nutrition of Grazing Ruminantsin Warm Climates (L.R. McDowell, ed.), p. 317. Academic Press, NewYork.
Williamson, P.R., Moog, R.S., Dooley, D.M., and Kagan, H.M. (1986). J. Biol.
Chem. 261, 16302.
Yagi, K., Kotaki, A., and Yamamoto, Y. (1965). J. Vitaminol. 11, 14.
Yang, F., de Villiers, J.S., McClain, C.J., and Varilek, G.W. (1998). J. Nutr. 128,
2334.
Yi, Z., Kornegay, E.T., Ravindran, V., and Denbow, D.M. (1996). Poult. Sci. 75,
240.
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chapter eighteen

E S S E N T I A L FAT T Y A C I D S

INTRODUCTION
Although essential fatty acids (EFA) are not vitamins by definition,
a deficiency disease or condition with dietary insufficiency does result,and in some ways, a similarity to vitamin deficiencies can be seen. The
finding that components of fat, other than the fat-soluble vitamins, aredietary essentials is of nutritional and medical importance. Excellent reviews in the literature of EFA have been prepared by Holman (1978a,b)and Hansen (1994).
Knowledge that carbohydrates can be readily converted into fat andthat essential lipid constituents such as phospholipids and cholesterol
can be made in the body led to the view that dietary lipids were not required. In 1926, Evans and Burr changed this viewpoint by reportingthat the total deprivation of fat in the diet of rats induced a syndrome of
deficiency that could be corrected by certain components of fat. The
EFAs originally included linoleic, linolenic, and arachidonic acids. However, arachidonic was later found to be synthesized from linoleic acid.
Most species have a dietary requirement for linoleic acid, while others(e.g., fish) require linolenic acid. Studies are reevaluating the beneficial
effects of linolenic acid in species that previously were considered toneed only linoleic acid as a dietary essential.

HISTORY
Historical aspects of EFAs have been reviewed (Aaes-Jørgensen,
1982; Mead, 1982). The earliest report that components of fat otherthan fat-soluble vitamins are dietary essentials for rats was made by
Evans and Burr (1926). Burr and Burr (1929, 1930) first demonstrated
675

18

CHAPTER EIGHTEEN

an essential dietary requirement by the rat for a specific unsaturated
fatty acid configuration that could not be synthesized by the animal. The
name “essential fatty acids” was coined to describe these unsaturated
fatty acids of linoleic and linolenic. Hume et al. (1940) reported thatarachidonic acid was also an EFA.
Most of the history of EFA is associated with Dr. R.T. Holman, of
the Hormel Institute, University of Minnesota, who has contributed
more to the knowledge of essential fatty acids than any other individual.
He is responsible for the delineation of metabolic conversions of polyunsaturated fatty acids, interactions among families of fatty acids, and determining quantitative requirements for linoleic and linolenic acids inanimals and humans.

CHEMICAL STRUCTURE AND PROPERTIES
Chemical structures of linoleic, linolenic, and arachidonic acids aswell as other fatty acids associated with EFA, are shown in Fig. 18.1.
There are three common families of unsaturated 18-carbon fatty acids
and one family of unsaturated 16-carbon fatty acids. The exact structureof an unsaturated fatty acid is given by three numbers: (1) the numberof carbon atoms in the chain, (2) the number of double bonds, and (3)the omega (ω) number, which indicates the number of carbon atoms
from the terminal methyl group to the carbon atom of the first doublebond. The omega system, which was originated by Holman, designatedthose unsaturated fatty acids belonging to each series. Another system(not used in this book) substitutes n for ω(e.g., n-6 versus ω-6). The ω9 and ω-7 series can be derived from endogenously synthesized oleic
acid (18:1ω-9) and palmitoleic acid (16:1ω-7), respectively. The ω-6 series is derived from linoleic acid (18:2ω-6) and the ω-3 series fromlinolenic acid (18:3ω-3). These latter two fatty acids are considered essential as they are products of plants and cannot be synthesized by animals. Thus, it appears that linoleic acid (18:2ω-6) is essential for most
species because of the inability of animals to synthesize a double bond
between carbons 6 and 7 counting from the terminal methyl group.
The polyunsaturated fatty acids are liquids at room temperature.
Double bonds of natural fatty acids would normally be found in natureas the cis-form. Ruminant animals ingest unsaturated fatty acids with
their plant foods. Subsequently, bacteria in the rumen use these unsaturated fatty acids as acceptors for excess hydrogen produced during bacterial anaerobic fermentation, thus resulting in saturation of the fatty
acids. However, in some cases, the end product is not a saturated bond
676

E S S E N T I A L FAT T Y A C I D S

Fig. 18.1 Structures of essential fatty acids and other unsaturated fatty acids.

but an unsaturated one with a different configuration—trans instead ofcis—or in a different position, or both. As a consequence, the body andmilk fat of ruminants contain trans-fatty acids, about 2 to 9% in thecase of butterfat.
A similar process, which uses hydrogen gas and a nickel catalyst, isemployed industrially to turn liquid edible oils into solid fats. During thehydrogenation process, some of the naturally occurring cis-double
bonds are isomerized to the trans-conformation, resulting in a decreased
bond angle and an acyl chain resembling a saturated fatty acid (Lichtenstein, 1993). Hydrogenation also results in the saturation of a portionof the existing double bonds in the fatty acyl chain, thereby decreasing
the polyunsaturated fatty acid (PUFA) and increasing the saturated and677

CHAPTER EIGHTEEN

monounsaturated fatty acid content of the fat. The trans-fatty acid content of commercial edible fats may vary from 0% for diet margarines
high in linoleic acid to more than 50% of fatty acids for certain shortenings and frying fats (Katan and Mensink, 1992).
One school of thought regarding trans- versus cis-fatty acids is that
dietary substitution of vegetable fats for animal fats reduces the risk ofcardiovascular heart disease (CHD). However, all forms of vegetable fat
are not alike, and research indicates that consumption of hydrogenatedrather than unhydrogenated vegetable oils may negatively influenceplasma lipids and risk of CHD. Low-density lipoprotein (LDL) cholesterol was raised by both trans-fatty acids and saturated fatty acids compared to a diet high in oleic acid (Katan and Mensink, 1992). Also,trans-fatty acids depressed high-density lipoprotein (HDL) cholesterol,
the so-called good cholesterol,” whereas saturates did not have this effect.
New forms of linoleic acid, conjugated linoleate acid (CLA) have received considerable attention as chemopreventive agents since they havebeen shown to inhibit rat mammary tumorigenesis, mouse forestomach
neoplasia, and mouse skin carcinogenesis (Belury, 1995; Parodi, 1999).
These compounds are linoleic acid derivatives with cis-9, trans-11-;trans-9, cis-11-; trans-9, trans-11-; trans-10, trans-12-; and trans-10, cis12-octadecadienoic acids accounting for the major isomers. The CLAs
are found predominantly in foods from ruminants, first identified as an
anticarcinogen following isolation from grilled ground beef extracts (Haet al., 1987).
Linoleic acid is a colorless oil that melts at −12oC. It is soluble inether, absolute alcohol, and other fat solvents and oils. It has an iodine
value of 181 and a molecular weight of 280.44. Arachidonic acid is an
oil that melts at −49.5oC, has an iodine value of 333.5, and has a molecular weight of 304.46 (Scott et al., 1982).

ANALYTICAL PROCEDURES
Essential and nonessential fatty acids are now readily determined by
gas-liquid chromatographic procedures (Holman et al., 1989) that aremuch more precise than alkaline isomerization analyses. Methyl estersof the fatty acids are formed before injection into the column; they arethen distributed between a moving gas phase (nitrogen, helium, or argon) and a stationary liquid phase (Hofstetter et al., 1965). Bioassays
are available in which growth rates of newly weaned rats are dependent
on the linoleic acid concentrations of feeds being tested.
678

E S S E N T I A L FAT T Y A C I D SMETABOLISM AND FUNCTIONSFats and fatty acid metabolism in relation to digestion, absorption,and excretion are discussed elsewhere (Dupont, 1990). Fatty acid-binding proteins (FABPs) have been identified as a family of cytosolic proteins found in heart, liver, and epithelial cells lining the small intestine(Anonymous, 1985). FABPs are believed to be integrally involved in thecellular uptake as well as intracellular transport and/or compartmentalization of fatty acids. It has been postulated that the intestinal FABPs
may actually participate in cellular fatty acid transport across the intestinal mucosa, as well as in selected intracellular events.
In humans, placental transfer of fat to the fetus occurs late in pregnancy. The late second trimester or early third trimester fetus has only
1.7% of its body weight as fat, compared with 15% in the full-term infant (Sosenko, 1995).
After fat absorption in monogastric animals, fatty acid compositionof body fat is directly related to fatty acid composition of the diet. In ruminants, however, polyunsaturated fatty acids are hydrogenated to alarge extent by ruminal microorganisms, resulting in more saturated
body fat of the animal. In all species, certain fatty acids form structuralcomponents and serve indispensable biochemical functions.
The relationship among the three common families of unsaturated
18-carbon fatty acids is shown in Fig. 18.2. Members of a particularfamily may be metabolically converted to more proximally unsaturated(toward the carboxyl group) or chain-elongated fatty acids, but no conversion from one ω family to another occurs in mammals. For example,
linoleic acid (18:2ω-6) is converted to arachidonic acid (20:4ω-6) in animals, and linolenic acid (18:3ω-3) may be converted to eicosapentaenoic (20:5ω-3) and docosahaexenoic acid (22:6ω-3). Members of the
ω-6 and ω-3 families are considered essential fatty acids for mammals,
because they cannot be synthesized de novo.
Linoleic acid and linolenic acid are the precursors of the entire ω-6and ω-3 families of polyunsaturated fatty acids, respectively. All members of the ω-6 and ω-3 families are active as essential fatty acids, andmany have been shown to be more active than their original precursor.
Studies employing graded dose levels of arachidonic acid fed to rats haverevealed that the deposition of arachidonic acid in liver is greater whenarachidonic acid itself is fed in the diet than when linoleic acid is fed.
This indicates that the conversion of 18:2ω-6 acid to arachidonic acidfor deposit in tissue lipids is a less efficient process than the deposition
679

CHAPTER EIGHTEEN

Fig. 18.2 Metabolic transformations of oleic, linoleic, and linolenic fatty acids.

of dietary 20:4ω-6 acid directly into tissue lipids, and that the potencyof 20:4ω-6 is greater than that of 18:2ω-6 (Holman, 1978a).
The effect of dietary deficiency of linoleic acid on the fatty acid composition of testicular tissue lipids for swine fed different levels of linoleic
acid is illustrated in Fig. 18.3 (Sewell and McDowell, 1966). The same
phenomenon has been observed in a large variety of both tissues andspecies, the differences being mainly in magnitude (Holman, 1978a,1986). Changes of greatest magnitude have occurred in heart and liver
lipids. At zero intake of linoleic acid, the major differences in fatty acidcomposition are in the polyunsaturated acids themselves. In linoleic aciddeficiency, 18:2ω-6, 20:4ω-6, and 22:5ω-6 are much lower than foundin normal animals. Palmitoleic acid, 16:1ω-7, and oleic acid, 18:1ω-9,
are higher than normal, but the most striking increase is in 20:3ω-9,which is formed endogenously from oleic acid. This acid, which is anormal component of tissue lipids in trace amounts, increases very dramatically in linoleic acid deficiency. It is found in the phospholipids inthe 2 position, the same position in which arachidonic acid, 22:5ω-6,and other polyunsaturated acids are normally found.
Similar studies have been made with graded dose levels of linolenicacid as the sole fatty acid supplement to a fat-free diet (Mohrhauer andHolman, 1963). Supplementation of the fat-free diet with 18:3ω-3680

Triene:Tetraene Ratio

E S S E N T I A L FAT T Y A C I D S

Dietary Linoleate (% Calories)

Fig. 18.3 Relationship of triene:tetraene ratio of testes tissue from young swine
fed varying levels of dietary linoleic acid. (Modified from Sewell and McDowell, 1966.)

causes dramatic increases in 20:5ω-3, 22:5ω-3, and 22:6ω-3 in comparison with the amounts found in the lipids of fat-deficient animals. Ayreand Hulbert (1996) noted that polyunsaturated fatty acids in muscle
phospholipids from rats fed high ω-6 or ω-3 fatty acid diets reflected thecomposition of their respective diets.
Studies on dose of a single fatty acid versus response of several fatty
acids in tissues have shown that each family of fatty acids suppresses metabolism of other families of fatty acids (Holman, 1964; Hwang et al.,1988).
In the absence of the main ω-6 (linoleic) and ω-3 (linolenic) families
in the diet, animals are capable of synthesizing some polyunsaturated
acids from endogenous precursors. Both oleic (18:1ω-9) and palmitoleic(16:1ω-7) acids themselves, and their respective families, are enhancedin the tissue lipids. None of these polyunsaturated acids is fully efficacious in meeting the requirement for polyunsaturation, for although
they are present in enhanced quantity in linoleic acid deficiency, the animals often die.
The enzymatic systems that perform chain elongation, desaturation,and insertion of fatty acids into various lipid molecules apparently handle all groups of fatty acids, for there is competition between substrates
at every step in each of these processes. The ω-3 family effectively suppresses metabolism of the ω-6 family. Likewise, the ω-6 family is able to681

CHAPTER EIGHTEEN

suppress metabolism of the ω-3 family, but less effectively. The ω-6 family, however, suppresses the formation of polyunsaturated acids fromoleic acid, as is manifested in linoleic acid deficiency. Ability of the precursor acids to compete for these enzyme systems is in the order linolenic> linoleic > oleic.
Depending on animal species, different EFAs are not equal in relationship to requirements or in ability to prevent all signs of EFA deficiency. For most species, linolenic acid does not fully relieve dermal signsof linoleic acid deficiency, even at high levels (Holman, 1978a). However, arachidonic acid (20:4ω-6) is twice as active as its precursor,
linoleic acid, in reducing dermal signs attributed to the deficiency.
The eicosanoids (eikosi = 20 in Greek), which are important in theregulation of widely diverse physiological processes, are derived from ω6 (e.g., 20:4) and ω-3 (e.g., 20:5) fatty acids. They have different biological activity depending on the precursor molecule. Prostaglandins,
thromboxanes, prostacyclins, leukotrienes, and hydroxy fatty acids areamong the eicosanoids that can be formed by enzymatic conversion of
di-homo-γ-linolenic, arachidonic, and eicosapentaenoic acids. Table
18.1 shows that eicosanoids have effects related to blood pressure, capillary permeability, inflammatory reactions, and blood platelet functions.
One of the most important specific metabolic functions of EFAs is
as precursors for a diverse group of “local hormones” called
prostaglandins. These biologically potent compounds seem to play aregulatory role in many cellular processes. Prostaglandins have beenshown to be involved in blood clotting, renal free water excretion, renal
blood flow, reproduction, bronchoconstriction, gastrointestinal motility
and water loss, endocrine function, and neurotransmitter release (Scottet al., 1982). Prostaglandins are intimately involved in regulation of immune function and thus resistance to infection (Boissonneault and Johnston, 1983).
Prostaglandins are formed by elongation and desaturation oflinoleic acid to dihomo-γ-linoleic acid (20:3ω-6) (DHGL) and to arachidonic acid (20:4ω-6), and from long-chain fatty acids of the linolenicfamily (20:5ω-3), eicosapentaenoic acid. These fatty acids are found inmembrane phospholipids. Practically all cells are capable either of producing or of being influenced by prostaglandins. A large number ofknown biologically active prostaglandins have been identified.
Prostaglandins formed from DHGL without further desaturation toarachidonate comprise the 1 series of prostaglandins. Prostaglandins
formed from arachidonate comprise the 2 series. The 3 series of682

E S S E N T I A L FAT T Y A C I D S■ Table 18.1

Cell Type

Biological Effects of Eicosanoids Derived from Two Essential Fatty Acid
Families
Arachidonic Acid
(20:4, ω-6)

Eicosapentaenoic Acid
(20:5, ω-3)

Several cells

PGE2
Immunosuppression
Vasodilating
Permeability
Hyperalgesia

PGE3
Immunosuppression
Inactive?

Platelets

TXA2
Aggregating
Vasoconstricting

TXA3
Antiaggregating
Inactive?

Endothelial

PGI2
Antiaggregating
Vasodilating

PGI3
Antiaggregating
Vasodilating

Leukocytes

LTB4 (inflammatory agent)
Chemotaxis
Aggregating
Receptor bind
Permeability

LTB5 (immunosuppressive agent)
Chemotaxis
Aggregating
Receptor bind
Permeability

Source: Modified from Drevon (1992).

prostaglandins is formed from eicosapentaenoic acid (20:5ω-3).
The discovery of prostaglandin-like molecules in human urine andplasma (Morrow et al., 1994) pointed to a nonenzymatic peroxidationof arachidonate, resulting in many different oxygenated products withprostaglandin-like structures. They are called isoprostanes because
many of them resemble the prostaglandins, with some differences instereochemistry, for example, 8-epi-prostaglandin F2-a versus prostaglandin F2-a (Hansen, 1994).
Generally, prostaglandins and leukotrienes constitute a group of extracellular mediator molecules that are part of an organism’s defense
system (Calder, 1998). Prostaglandins and leukotrienes are formed during the inflammatory process, and if the inflammation is caused by invading bacteria, the formation of prostaglandins and leukotrienes will
stimulate macrophages and other leukocytes to begin the process of destroying the bacteria.
Lipid sources could alter the development of autoimmune disease
and the life span of short-lived animals. Many investigators have observed that polyunsaturated lipids that hinder cardiovascular disease(when compared to saturated fats) can be proinflammatory (Fernandes
and Venkatraman, 1993).
Macrophages are involved in immune and inflammatory functions
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and possess the enzymes necessary for prostaglandin and leukotriene
biosynthesis. Consuming high levels of ω-3 fatty acids may provide considerable health benefits in relation to inflammatory diseases, such as
atopic dermatitis and rheumatoid arthritis (Drevon, 1992). The mechanism behind the potentially beneficial effect of ω-3 fatty acids on someinflammatory diseases may be related to altered eicosanoid formation.
Reduction of eicosanoid biosynthesis by inflammatory cells is of clinicalinterest because of the immunosuppressive potential of elevated levels of
prostaglandin E (PGE) (Kinsella et al., 1990) and possibly leukotriene B4(LTB4) (Goodwin, 1985). For instance, production of leukotriene derived from 20:5ω-3 (e.g., LTB5) is an immunosuppressive result for oneof the more potent inflammatory agents (LTB4), which is derived from
20:4ω-6 (Drevon, 1992).
Many age-associated diseases, including malignancy and autoimmune disease with a viral etiology, appear to be exacerbated by high-fat
diets with a large proportion of vegetable oils high in ω-6 fatty acids.
These oils could increase autoimmune disease by increasing free radical
formation and decreasing levels of antioxidant enzyme mRNA, thus further decreasing immune function, in particular by inhibiting the development of anti-inflammatory cytokines such as interleukin (IL-2) andtransforming growth factor (TGFb). In contrast, ω-3 lipids could protect
against autoimmunity by enhancing TGFb mRNA levels and preventing
an increase in oncogene expression (Fernandes, 1995).
Polyunsaturated fatty acids have a structural function as an integral
part of phospholipids, the building unit of biomembranes. This is inferred from the specific composition of the fatty acids in these phospholipids (the β position normally being esterified with the highly unsaturated members of the EFA families) and from the fact that in EFA
deficiency, these fatty acids are replaced by eicosatrienoic acid (20:3ω9), biosynthesized from oleic acid (18:1ω-9), with the known concomitant deleterious effects on biomembrane function and integrity. The
phospholipids of cell membranes will influence membrane viscosity andpermeability and thereby possibly the enzyme activity of membrane proteins. Different types of eicosanoids are formed from different essential
fatty acids. The general rule is that eicosanoids derived from eicosapentaenoic acid are less potent than the corresponding compounds derived
from arachidonic acid (Drevon, 1992).
It has been suggested that EFA deficiency and replacement of thelinoleic acid family in membrane structures may cause a disruption in
spatial arrangements in mitochondria that results in less efficient oxidative phosphorylation and a derangement of basal metabolism. Such a
684

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process may be the partial uncoupling of oxidative phosphorylation in
mitochondria. In poultry, presence of linoleic acid may not be absolutely
necessary in the body since a deficiency will not always result in death.
Fatty acids that replace the linoleic acid family in tissue lipids seem tocause a reduction in the metabolic efficiency and functioning of the animal, but life often can still be maintained (Scott et al., 1982).
A disturbed water balance is a characteristic defect of EFA deficiency and can include increased water loss through the skin, increased
urinary arginine-vasopressin loss, increased water intake, and reduced
urine output (Holman, 1968; Hansen and Jensen, 1986; Hansen, 1994).
Increased water loss through skin results from a defect in the permeability barriers of skin, which is an indication that EFAs are involved inmembrane structure. Histological studies have shown many changes inskin structure as a result of the deficiency.
Additional functions of EFAs include provision of adequate fluidity
to sustain cellular function and for lipid transport (Holman, 1978a).
Phospholipids and cholesteryl esters containing an abnormally high proportion of saturated fatty acids would tend to be more rigid or less fluid
than would similar compounds with high proportions of polyunsaturated acids. Ethanol can penetrate the lipid bilayer of the cell membrane
and can cause changes in the structure and organization of the fatty acidcore, thus changing the membrane fluidity (Hoek and Rubin, 1990).
Ethanol may affect the uptake of very long chain fatty acids from thediet and/or their incorporation into lipids (Swanson et al., 1995).
One of the functions of polyunsaturated acids is to provide lipids
that are fluid at body temperature. Alloxan diabetes, hyperthyroidism,
dietary cholesterol, saturated-fat diet, or mineral oil all involve thetransport of a nonessential lipid in large quantities. These conditions
have been found to accelerate EFA deficiency significantly. Studies suggest that one function of polyunsaturated acids is to provide necessary
structural components for circulating lipoproteins (Holman, 1978a).
In a review, Vergroesen (1977) summarized the beneficial effects ofadequate linoleic acid: (1) decreased blood cholesterol and triglyceride
levels, (2) decreased thrombotic tendency of platelets, (3) preventive andcurative effects in sodium-induced hypertension, (4) improvement of thephysiological function of the heart, and (5) normalization of the biochemical abnormalities in obesity and maturity-onset diabetes. Mechanisms of these responses are not clearly established; however, many ofthe biological effects are derived from the eicosanoids synthesized fromarachidonic acid (20:4ω-6), including prostaglandins, thromboxanes,
prostacyclins, and leukotrienes.
685

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Linoleic and linolenic acids stimulate the growth of mammary epithelium in normal rats. Dietary linoleate seems to be necessary to thedevelopment of mammary ducts and alveoli in immature mice and totheir maintenance in adult animals (Knazek and Liu, 1979). Using rats,
Ollivier-Bousquet et al. (1993) suggested that only ω-6 fatty acids (versus ω-3) are required for the optimal functioning of lactating mammary
epithelial cells.
CLA (see Chemical Structure and Properties) has been identified as
an anticarcinogen. The CLA has several unique structural and functional properties, resulting in chemical and physiological effects that are
different from those of all-cis- nonconjugated polyunsaturated fatty
acids. In turn, these unique qualities appear to modulate cellular
processes involved in carcinogenesis (Belury, 1995).
Synthesis of CLA requires the presence of free linoleic acid (the substrate), a free radical-generating species, and proteins rich in sulfur
residues (Dormandy and Wickens, 1987). These conditions occur in vivo
(through oxidative pathways and enzymatic isomerization) and in vitro(treatment of foods with heat) (Chin et al., 1992). The major dietary
sources of CLA are foods derived from ruminants, such as beef andcheese. More than 85% of the CLA in animal tissues is the cis-9, trans11 isomer form. They have been found in triglycerides, lipoproteins, andcell membrane phospholipids in several tissues of rodents, rabbits, andhumans. Intestinal bacterial flora of rats is capable of converting freelinoleic acid to cis-9, trans-11, and trans-9, cis-11 CLA isomers (Chin etal., 1994).
Increased attention has been given to the question of possible healthbenefits of ω-3 (linolenic family) of polyunsaturated fatty acids (Lands,
1986; Drevon, 1992; Hansen, 1994). Whatever the mechanisms involved, epidemiological studies of Greenland Eskimos (who subsist entirely on a marine diet high in ω-3 fatty acids) clearly indicated that theirdiet exerts potentially antithrombotic effects on platelet function, with alow death rate from coronary heart disease (Willis, 1984). In Japan, alower death rate from coronary heart disease was also related to higher
fish consumption. The Japanese studies showed a dose-response effect offish intake, incorporation of ω-3 fatty acids in plasma lipids, and reduced cardiovascular disease. Cerebrovascular deaths were also reduced(Leaf, 1992). Similar results were obtained in human or animal studies
in which fish oils rich in 20:5ω-3 were administered. The most striking
result of human studies is marked prolongation of bleeding time. This
clearly indicates that marine diets may reduce platelet plug formation indamaged blood vessels and may inhibit vessel-wall-induced clotting of686

E S S E N T I A L FAT T Y A C I D S

plasma, as observed in rats (Willis, 1984).
Using isolated working heart models from rats, Pepe and McLennan(1996) reported that dietary fish oil prevented the initiation and reduced
the severity of arrhythmias in the isolated hearts in response to a variety
of stimuli. These results establish that irrespective of any effects on
blood pressure or platelet function in vivo, dietary fish oil directly affects
myocardial properties, which may contribute to observed clinical reductions in cardiac mortality associated with fish consumption.
From studies on postinfarct transcutaneous angioplasty, there aresome conflicting data on the effect of very long chain ω-3 fatty acids onthe frequency of re-stenosis, while a prospective study on mortality after coronary artery infarction reported a significant decrease (29%) in
all-cause mortality after 2 years for patients using at least two fatty fishmeals weekly (Burr et al., 1989). Sudden cardiac death accounts forabout 50% of total coronary disease mortality in westernized industrial
countries. The lack of early symptoms for this disorder makes prevention the preferred strategy. In a rat model of cardiac ischemia, dietary ω6 (sunflower seed oil) and ω-3 (fish oil) polyunsaturated fatty acids wereshown to protect against arrhythmia compared with saturated fat;greatest protection was observed with fish oil (Topping, 1993). Studies
have shown that one aspirin every other day reduced CHD events by
47% in 5 years in 23,000 men. Aspirin interferes with platelet aggregation by inhibiting the enzyme cyclooxygenase in platelets. ω-3 Fatty acidsupplements have been shown to inhibit the same enzyme and to inhibit
platelet aggregation (Leaf, 1992).
The brain, as well as most of the nervous system, is a well-protectedorgan, especially with regard to polyunsaturated fatty acids. The nervous system has the greatest concentration of lipids after adipose tissue.
Interestingly, these lipids are practically all structural and are not related
to energy production; they participate directly in the functioning of cerebral membranes. Brain lipids are formed of polyunsaturated fatty acids
derived from dietary essential linoleic and linolenic acids. On average,
one fatty acid out of three in the brain is polyunsaturated, and thesefatty acids participate in the structure of phospholipids.
The ω-6 fatty acids are required for normal fetal growth, but ω-3
fatty acids are beneficial for surfactant synthesis (Viscardi, 1995). Surfactant is a phospholipid-rich substance that lines the air-alveolar interface in the lung and prevents alveolar collapse, a major cause of mortality and morbidity in premature infants.
The long-chain ω-3 fatty acids are found in high proportions in reproductive and nervous tissues. The elongated docosahexaenoic acid687

CHAPTER EIGHTEEN(22:6ω-3) is the most abundant fatty acid in the ethanolamine phospholipids of cerebral gray matter and the retina (Carlson et al., 1986; Connor et al., 1992). The need for ω-3 fatty acids in developing visual acuity was presented as evidence for a functional requirement for ω-3 fatty
acids in primates (Neuringer et al., 1984). Differences in physical activity and ability to learn have been related to low content of 22:6ω-3 inbrains of rats produced by feeding a diet low in linolenic acid (Lamptey
and Walker, 1976).
A diet deficient in linolenic acid alters nerve-ending fluidity and enzymatic activities, reduces the amplitude of electrophysiological parameters such as the electroretinogram, alters the resistance of the nervous
system to poisons, and reduces the performance of learning tasks(Bourre et al., 1989, 1993; Enslen et al., 1991; Zerouga et al., 1991). It
has been shown that changes in lipid membrane composition can modulate the binding of neurotransmitters or hormones to membrane receptors at the periphery (Murphy, 1990). Modifications of the neurotransmission pathways might induce the behavioral disturbances that are
seen in animals (Delion et al., 1994).
Diets rich in PUFAs have been associated with promotion of tumor
growth in rodents under certain conditions. However, studies in rats andmice (Cave, 1991) have shown that not all PUFAs should be incriminated. Increasing the fat contribution from ω-6 PUFAs (in vegetable oils)
from 0 to 4% dietary energy routinely enhances tumorigenesis in rodents, while diets with equivalent levels of long-chain highly unsaturated
ω-3 fatty acids found in fish oil often diminish formation of certain tumors.
Epidemiologic studies suggest that individuals who consume dietsrich in fat are at higher risk for colon cancer, whereas consumption offish products rich in ω-3 fatty acids, such as eicosapentaenoic acid (EPA)
and docosahexaenoic acid (DHA), is associated with low incidence ofcolorectal cancer (Willett et al., 1990). Fish oil seems to protect against
cancers that are most likely linked to diet—those of the colon, pancreas,
breast, and ovary.
Leaf (1992) reviewed the effect of ω-3 fatty acids in reducing cardiovascular disease; additional physiologic and pharmacologic effects offish oils were referenced as follows: (1) decreases blood pressure in normal and moderately hypertensive subjects, (2) decreases blood viscosity,(3) decreases microvascular albumin leakage in insulin-dependent diabetics, (4) decreases plasma triglycerides, (5) decreases vascular response
to norepinephrine, (6) decreases ventricular fibrillation from ischemia,

688

E S S E N T I A L FAT T Y A C I D S(7) decreases cardiac toxicity of cardiac glycosides, (8) decreases platelet
adhesion, (9) decreases leucocyte-endothelium interactions, (10) increases vascular compliance, and (11) increases platelet survival.

REQUIREMENTS
The dietary essentiality of both linoleic acid (18:2ω-6) and linolenicacid (18:3ω-3) is dependent on species and, to a certain extent, on thedefinition of an essential nutrient. Table 18.2 provides EFA requirementsfor various animals and humans.
The predominant EFA for most mammals and birds is linoleic acid;the requirement for linolenic acid is much less and is unknown for many
species. Determination of linoleic acid requirements have been based on
observations of gross dermal lesions as well as variations in tissue
polyunsaturated fatty acids. Shifts in fatty acid composition of metabolically active tissues (liver, heart, brain, etc.) that occur during onset of
deficiency are very similar in all species. Similarities between EFA deficiencies induced in various species are striking when the biochemical parameters of the deficiency are considered. Quantitative requirements forseveral species are also strikingly similar when measured with biochemical parameters.
Triene:tetraene is the ratio of abnormally elevated endogenous
metabolite of oleic acid, 20:3ω-9, to the metabolic product, 20:4ω-6, derived from linoleic acid (see Deficiency), and it has been used to estimate
the minimal linoleic acid requirement (Holman, 1960). The ratio drops
from a high value in deficiency to a low and rather constant value in theregion between 1 and 2% of calories. The implication from these biochemical parameters is that dietary requirement for linoleic acid lies between 1 and 2% of calories.
Holman (1960) suggested a ratio of 0.4 as the point at which theminimal linoleic acid requirement of the rat, as well as other species, hasbeen met. Fig. 18.3 illustrates the plotting of the triene-tetraene ratio
versus six dietary linoleic acid levels in 3-week-old swine testes (Sewelland McDowell, 1966). The ratio decreased markedly as dietary level oflinoleic acid increased from zero to 1% of dietary calories, with only aslight decrease occurring beyond this level of linoleic acid intake. Requirement for linoleic acid for pigs of this age is therefore less than 2%of dietary calories. The triene-tetraene ratio at the 1% level of linoleic
acid was 0.38, which is comparable to the figure of 0.4 suggested byHolman (1960).

689

■ Table 18.2

Essential Fatty Acid Requirements for Various Animals and Humans

AnimalPurpose or Class

Requirementa

Reference

Dairy cattle
Chicken

10% fat
1% linoleic
1% linoleic
1–1.4% linoleic

NRC (1989a)
NRC (1994)
NRC (1994)
Scott et al. (1982)

Japanese quail
Sheep

Calf milk replacer
Leghorn, 0–18 weeks
Leghorn, breeding
Leghorn,laying-breedingb
Broilers
0–8 weeks
8–20 weeks
Growing-breeding
Growing

Swine

Growing

Horse
Catc

All classes
Growingc

NRC (1994)
NRC (1994)
NRC (1994)
NRC (1994)
Bruckner et al.
(1984)
Sewell and
McDowell (1966)
NRC (1989b)
NRC (1986)

Dog
Mink

All classes
Adult
Pregnancy-lactationMales

1% linoleic
1% linoleic
0.8% linoleic
1% linoleic< 0.32 linoleic
as energy
1–2% linoleic
as energy
0.5% linoleic
0.5% 18:2 ω-6 and
0.02% 20:4 ω-6
1% linoleic
0.5% linoleic
1.5% linoleic
1.3% linoleic
as energy
0.5% linoleic
as energy
0.88–1.04% linoleic
as energy
0.68% linoleic
as energy

TurkeyRat

Females
Guinea pig

Growing

Mouse

All classes

Nonhumanprimates

All classes

Fish

Chum salmon
Rainbow trout
Yellowtail

Human

All classes

1–2% linoleic
as energy
1% linoleic and
1% linolenic
1% linoleic and
0.8% linolenic
2% 20:5ω-3 and22:ω-6
1–2% as energy

NRC (1985a)
NRC (1982a)
NRC (1982a)
NRC (1995)
NRC (1995)
NRC (1995)
NRC (1995)
NRC (1978)
NRC (1993)
NRC (1993)
NRC (1993)
RDA (1989)

aOften requirements are unknown; for some species, treatments or essential fatty

acid levels that have been successful are noted. Requirements are expressed either as percentage of diet or as percentage of calories (energy). Values can be on either as-fed (approximately 90% dry matter) or dry basis (see Appendix, Tables A1a.b).
bFor laying and breeding hens, 1.4% linoleic acid is required; after maximum egg
size is reached, 1% is adequate.
cThe cat has very limited abilities to desaturate linoleic and linolenic acid to longerchain fatty acids.

690

E S S E N T I A L FAT T Y A C I D S

Contrary to most species, evidence has shown that the cat family(e.g., cats and lions) is unable to desaturate linoleic and linolenic acids(NRC, 1986). As a result, these species may require a source of preformed longer-chain fatty acids, which means that these animals may
exhibit a specific requirement for polyunsaturated lipids of animal origin. Nevertheless, linoleic acid prevents several signs of EFA deficiency,
including scaly skin, increased transepidermal water loss, and enlarged
fatty livers. Thus, linoleic acid has a specific role as an EFA independentof arachidonic acid synthesis.
Most fish, unlike terrestrial animals, have a definite requirement forlinolenic acid (ω-3), while linoleic acid (ω-6) is often of less value. The
majority of fish and shellfish species studied required 18:3ω-3, some required a combination of 18:3ω-3 and 18:2ω-6, and only a few showeda preference for 18:2ω-6 (NRC, 1993). For some fish species (e.g., rainbow trout), the metabolite of linolenic acid, 22:6ω-3, was more effectivein stimulatory growth and other production parameters than linolenic
acid. At least four species of fish required ω-3 fatty acids, and 22:6ω-3
best satisfied the requirement. One suggestion to explain the differencein EFA requirements for fish was that ω-3 structure permitted a greater
degree of unsaturation, which was necessary in membrane phospholipids to maintain flexibility and permeability characteristics at low temperatures (NRC, 1993).
A number of factors influence development of EFA deficiency andthus requirements for EFA:
1. Age and carryover effects—Animals that had been fed normal dietsfor a longer time should have larger reserves of polyunsaturated fatty acids
and could withstand deficiency for a longer time than weanlings. Inducing
dermal signs of EFA deficiency is difficult in adult animals. Linoleic acid requirement of young chicks can be affected markedly by carryover of linoleic
acid from the egg to newly hatched chicks. If chicks hatch from eggs low inlinoleic acid and are fed purified diets very low in linoleic acid, the dietaryrequirement may be in excess of 1.4%, compared to the typical requirementof 1.0% (Scott et al., 1982).
2. Dietary fat and hormone imbalance—Animals that practice coprophagy have an additional source of lipids not available to animals that
do not. However, diets rich in saturated fatty acids or monounsaturated
fatty acids are also known to moderately enhance the development of EFA
deficiency. Peifer and Holman (1955) studied the effect of adding 1% cholesterol to the diet of EFA-deficient rats. An EFA deficiency syndrome,691

CHAPTER EIGHTEEN

judged by growth and dermal signs, occurred within periods of 2 weeks to
1 month. Comparable EFA deficiency signs were observed only after 3
months in rats on fat-free diets without cholesterol. Substances or conditions that induce hypercholesterolemia likewise accelerate EFA deficiency.
3. Growth rate—Any animal that is called upon to grow more rapidly,
and therefore to build more tissue, would have a higher requirement for EFA
and would consequently exhibit deficiency signs earlier.
4. Humidity and water balance—Low atmospheric humidity hastens
onset of dermal signs of EFA deficiency, probably through enhanced loss ofwater by evaporation, causing additional irritation of skin. Aaes-Jørgensen
and Dam (1954) reported an experiment in which female rats were raisedon diets with various amounts of fat for 16 weeks. They found that the water intake was higher and urine production lower in rats on diets with hydrogenated peanut oil or hydrogenated whale oil and in the absence of dietary fat than in rats on diets with lard, peanut oil, or coconut oil.
5. Sex of animal—Male animals are known to be more sensitive thanfemales to EFA deficiency. The requirement for the female rat was found tobe between 10 and 20 mg/day, while the male rat’s requirement exceeded 50
mg/day (Greenberg et al., 1950; NRC, 1995). Pudelkewicz et al. (1968) estimated the linoleic acid requirement of growing female and male rats to be
0.5 and 1.3% of dietary metabolizable energy, respectively.
6. Pen arrangement—Leat (1962) concluded, from feeding pigs, thatwhether the animals are penned individually or in groups is important in thedevelopment of EFA-deficiency signs. He found that the skin condition wasnoticeably better in pigs penned in pairs than in those penned individually.
He believed that keeping animals in close proximity with each other may
prevent dermatitis from becoming apparent merely through physical contact.
7. Temperature and environment—For fish, EFA requirements maychange with temperature and culture conditions. When rainbow trout fromthe same source were tested simultaneously in seawater and fresh water, theEFA deficiency was manifested more quickly in seawater (NRC, 1993).
Likewise, the fatty acid composition of fish lipids, especially membrane
lipids such as phospholipids, are significantly affected by acclimation temperature.

For species that require a predominance of linolenic versus linoleic
acid, information is lacking on linolenic acid requirements. For pregnant
rats and pups up to weaning, the minimum linoleic and linolenic acid intake should be 2.4 and 0.4% of total dietary energy, respectively (Bourre
et al., 1990). For the adult rat, linolenic acid requirement is reduced to692

E S S E N T I A L FAT T Y A C I D S

0.26% of dietary energy (Bourre et al., 1993).
For humans, many recommendations are that a minimum intake ofessential fatty acids should be 3% of total energy intake, whereas manynational nutrition committees have recommended that not more than
10% of total energy should be derived from PUFA, monoenes, and saturated fatty acids (Drevon, 1992).
On the basis of human milk composition, Neuringer et al. (1988)
recommend 0.7 to 1.3% of energy as linoleic acid, with the optimal ω6:ω-3 ratio 4 to 10:1. If the diet contains an ω-6:ω-3 fatty ratio greaterthan 50:1, ω-3 fatty acid deficiency is expected. However, a high ω-3level (ω-6:ω-3 1.4:1) depressed the cell-mediated immune response,
PGE2 production, and plasma vitamin E in aged dogs (Wander et al.,1997).

NATURAL SOURCES
The EFAs are widely distributed among food fats. For example,
vegetable oils of corn, soybean, cottonseed, peanut, and certain others
are excellent linoleic acid sources. Safflower oil contains 75% linoleic
acid, whereas corn oil, soybean oil, and cottonseed oil all contain approximately 50% linoleic acid. Linseed, canola, and soybean oils contain approximately 57, 8, and 7% linolenic acid, respectively (Drevon,
1992). Linolenic acid is particularly high in forage lipids. From thelipids of pasture grasses, 61% is reported as linolenic acid (Garton,1960).
Linoleic and arachidonic acid contents of feed ingredients are shownin Table 18.3. Linoleic acid and its dehydrogenation product 18:3ω-6
are found in highest abundance in plants, but more unsaturated andlonger-chain members of this family are found principally in animals.
Notable exceptions to these generalities are the occurrence of arachidonic acid and other higher members of the group in primitive plants
such as ferns and algae (Schlenk and Gellerman, 1965). Concentrationsof the long-chain linolenic acid (ω-3) family for marine products arelisted in Table 18.4.
Arachidonic acid (20:4ω-6) is the most abundant PUFA in animal
membranes; thus, when animal products are consumed as food, thearachidonic acid content varies with the amount of membrane. Crawford et al. (1989) determined the arachidonic acid composition in various species in the food chain as follows (percent of total fatty acids):
wild pig (8.5), antelope (7.4), dolphin muscle (16.8), herring muscle(0.6), squid (5.8), mollusk (2.3), algae (12.4), and phytoplankton (0.7).
693

CHAPTER EIGHTEEN■ Table 18.3

Typical Linoleic Acid and Arachidonic Acid in Various Foods and Feedstuffs (as-fed basis)

Food or Feedstuff
Alfalfa meal, dehydrated
BarleyBrewer’s grains, dehydrated
Coconut oil
Corn gluten meal
Corn oilCorn, yellow
Cottonseed meal, solvent extracted
Crab meal
Fish meal, anchovy
Fish meal, menhaden
Fish oil, menhaden
Fish solubles, condensed
Lard
Linseed oil
Meat meal
Milk, cow’s, dehydrated
Oats
Peanut meal
Poultry by-products meal
Poultry fat (offal)
Rice bran oil
Safflower oil
Sorghum
Soybean meal, solvent extracted
Soybean seed
Tallow
Wheat
Wheat bran

Linoleic Acid (%)

Arachidonic Acid (%)

0.40
0.83
2.94
1.10
3.83
55.40
1.82
0.80
0.33
0.20
0.15
2.70
0.20
18.30
13.90
0.34
0.01
1.49
1.25
1.72
22.30
36.50
72.70
1.08
0.35
7.97
4.30
0.58
2.25

—
—
—
—
—
—
—
—
—
—
—
—
—
0.3–1.0——————
0.5–1.0
—
—
—
—
—
0–0.2
—
—

Source: Data adapted from NRC (1982b).

CLA (see Chemical Structure and Properties; Metabolism andFunction) is a naturally occurring substance in food. Dietary sources of
CLA include milk fat, meat products, and vegetable oils. In foods, CLA
is highest (mg/g of fat) in ruminant meats and is found in smaller
amounts in poultry and eggs (Chin et al., 1992). Dairy products suchas natural cheeses, processed cheeses, and milks and yogurts that haveundergone a variety of heat-processing treatments all contain considerable amounts of CLA (Lin et al., 1995). Total CLA is generally increased in foods that are heat processed (dairy pasteurization, pan frying of meats, etc.) (Shantha et al., 1992). Overall, vegetable fats arepoorer sources of CLA. Concentrations of CLA in bovine milk can beincreased by dietary regimen and management (Jiang et al., 1996;

694

E S S E N T I A L FAT T Y A C I D S■ Table 18.4

Eicosapentaenoic (20:5ω-3) and Docosahexaenoic (22:6ω-3)
Acids in Fish or Marine Products

Carp
Catfish, channel
Cod, Atlantic
Herring
Mackerel
Salmon (wild)
Trout (wild)
Crab, blue
ShrimpOyster, east
Cod liver oil
Salmon oil
Menhaden oil

Total Fat
(g/100 g)

20:5

22:6

5.6
4.3
0.7
9.0
13.9
5.4
7.7
1.0
1.5
2.0
100.0
100.0
100.0

0.2
0.1
0.1
0.7
0.9
0.3
0.1
0.2
0.3
0.3
9.0
8.8
12.7

0.1
0.2
0.2
0.9
1.6
0.9
0.5
0.1
0.2
0.2
9.5
11.1
7.9

Sources: Data adapted from Drevon (1992) and Sanders (1994).

Chouinard et al., 1999).

DEFICIENCY
Effects of Deficiency
Induction of EFA deficiency in animals requires rigid exclusion offat from the diet, and even with supposed low-fat diets for humans, deficiency state in adults is difficult to attain (Holman, 1978b). Clinicalsigns of EFA deficiency induced by a fat-free diet require almost oneeighth of a rat’s normal lifetime to develop, and rarely have humans
been subjected to a low-fat diet under observation for a proportionate
span of time. Natural diets, even poor ones, usually contain adequate
amounts of EFA; therefore, the deficiency is far rarer than deficiencies ofprotein, vitamins, or minerals. Nevertheless, EFA deficiency does occur
when animals or humans receive insufficient dietary fat.
Linoleic acid deficiency signs and other criteria range from earlyclassic signs such as reduced growth rate, parakeratosis, increased water
permeability of skin, increased susceptibility to bacteria, and male andfemale sterility to more recently recognized signs such as decreased
prostaglandin biosynthesis, reduced myocardial contractility, abnormal
thrombocyte aggregation, and swelling of rat liver mitochondria (Ver695

CHAPTER EIGHTEEN

groesen, 1977). For all land species studied, the major feature of the deficiency is impairment of the exterior covering of the animal. Mammals
exhibit dermatitis, chickens exhibit faulty feathering, and moths are unable to form normal scales on their wings. All the manifestations indicate faulty membrane formation, a feature of deficiency that is common
to all tissues and species (Holman, 1978a).
For land animals, EFA deficiency is a term primarily referring tolinoleic acid deficiency. However, findings suggest that the linolenic family (ω-3) of fatty acids, particularly 20:5ω-6 and 22:6ω-3, are likewise
important. Fish oils, rich in ω-3 fatty acids have been linked to reduction of thrombosis and heart disease in humans as well as proper brain
development in children and rats (Drevon, 1992; Jumpsen et al., 1997).

Ruminants
The EFA deficiency of ruminants has been less extensively researched than that of nonruminants, with the deficiency in adult ruminants not readily demonstrated (Palmquist et al., 1977). The microbial
population appears to provide enough EFA to meet the requirements;
however, studies with lambs suggest that the required level of EFA may
be elevated in the presence of host microflora (Bruckner et al., 1984).
Gullickson et al. (1942) reported that calves fed a low-fat diet did not
develop EFA deficiency signs, but growth was suppressed. Cunningham
and Loosli (1954b) reported that calves receiving a fat-free synthetic
milk developed leg weakness and muscular twitches within 1 to 5 weeks
and died unless a source of fat was supplied. Lambert et al. (1954) alsostudied the effect of a “lipid-free,” semisynthetic milk fed to dairy
calves. They reported the following clinical signs: growth retardation after 3 weeks on trial; scaly dandruff; long dry hair; dull hair coat; excessive loss of hair on the back, shoulders, and tail; and diarrhea.
Weanling lambs fed a fat-free diet for 7 months showed no evidenceof skin lesions or other clinical signs typical of fat deficiency (Cunningham and Loosli, 1954a). In a second experiment, 2-day-old lambs andkids were given fat-free synthetic milk. The lambs and kids receiving thefat-free diets became weak and died within 1 to 7 weeks, while controls
were raised successfully on the same milk with 2% added lard.
Delivery of fatty acids at various levels for metabolism can influence
events important in dairy cow reproduction. A soybean oil emulsion(50% linoleic acid) was infused intravenously to Holstein heifers (Lucyet al., 1990). This resulted in increased plasma concentrations of
prostaglandin F 2-α(PGF2α) metabolite and increased ovarian follicles,
and the size of the largest follicle was greater. In a second study, feeding696

E S S E N T I A L FAT T Y A C I D S

rumen-protected fat to lactating dairy cows increased the numbers of 3to 5-mm follicles and follicles greater than 15 mm in diameter, and increased the size of the preovulatory follicle of a synchronized estrus cycle during the early postpartum period (Lucy et al., 1991). Garcia-Bojalil (1993) fed rumen-protected fat (0.5 kg/day), which improved
conception rates of lactating Holstein cows from 52 to 86%.
Inclusion of fish oil in the diet appears to result in an alteration inregression dynamics of the corpus luteum as evidenced by a greater proportion of cows having elevated concentration of plasma progesterone
after injection of PGF2α (Burke et al., 1996). Perhaps the increase in conception rate (39.5 versus 30.6%) could be attributable to increased survival of the embryo at the time of pregnancy recognition (e.g., when
PGF2α secretion is suppressed). It would appear that fatty acids are important to both stimulated follicles (elevated PGF2α) to bring about
pregnancy but later decrease PGF2, which would result in greater progesterone production and maintenance of pregnancy.

Swine
Witz and Beeson (1951) used a diet that contained only 0.06% lipid
and produced the following signs: slower growth rate; underdeveloped
digestive systems; small gallbladders; enlarged thyroid glands; delayed
sexual maturity; scaly dandruff-like dermatitis on the tail, back, andshoulders; loss of hair, with the remaining hair being dull and dry; abrown, gummy exudate on the belly and sides; necrotic areas on the skinaround the neck and shoulders; and an unthrifty appearance. Leat
(1962) fed pigs from 4.5 to 91 kg live weight a diet consisting of 0.07%
of the calories as linoleic acid. This diet resulted in pronounced scalinessof the skin, first noted after about 13 weeks on the diet. Scaliness seemed
to be confined to the dorsal surface and was most severe about theshoulders. The hair was dry and appeared to stand out from the skin at
all angles. When linoleic acid made up 0.5% of the dietary calories,
there was little or no flakiness of skin.
Sewell and McDowell (1966) fed 3-week-old male pigs purified diets containing six levels of linoleic acid. For the 10-week experiment, nodifferences were noted in weight gains, but dermal lesions (Fig. 18.4)
were observed after 6 to 7 weeks of the experiment. Scaly, dandruff-likedesquamation of the skin over the dorsal surface was the first noticeable
sign, and later, a brownish gummy exudate appeared around the ears
and axillary spaces, and under the flanks. Skin eruptions were also present about the ears, axillary spaces, and flanks in the severest cases. Lesions were observed only among pigs receiving 0.5% linoleic acid or
697

A

B
Fig. 18.4 (A) Dermal lesions of a linoleic acid-deficient pig (foreground) thatwas fed a diet with no linoleic acid (the black spots are flies that were attracted to the brown, gummy exudate). The other pig received 1.0% of calories as linoleic acid. (B) The same pigs as in A; the one on the right has EFA
deficiency, with dermal lesions particularly severe at the axillary spaces. (Courtesy of L.R. McDowell, R.F. Sewell, and the University of Georgia.)

698

E S S E N T I A L FAT T Y A C I D S

less; pigs receiving the 1% level and above were free of skin lesions.
Studies have been conducted in swine on the effect of PUFA in modulating infectious disease processes. Turek et al. (1994) found that dietary (ω-3 and ω-6) PUFAs affected alveolar macrophage tumor necrosis factor (TNF) production and leucine aminopeptidase (LAP) levels,and the production of T-cell growth factors by alveolar lymphocytes. In
another study, these researchers found that dietary PUFA can affect disease pathogenesis (e.g., Mycoplasma hyopneumonia, the most common
respiratory pathogen in swine) and that the ω3:ω6 PUFA ratio may
modulate the host response (Turek et al., 1996).

Poultry
Growing chicks fed a fat-free ration did not survive the fourth week
(Reiser, 1950). The most readily observed clinical sign of linoleic aciddeficiency in young chicks is slow growth rate. Machlin and Gordon(1961) found that adding safflower oil or linoleic acid, but not linolenicacid, to purified diets free of unsaturated fatty acids resulted in an immediate (within 7 days) growth response in chickens. Some microbial
synthesis of ω-6 fatty acids occurs in the gut of quail. On a diet deficientin linoleic acid, germ-free animals had a lower growth rate and more severe clinical signs than normal ones.
Linoleic acid deficiency in chicks has also been reported to result inan enlarged fatty liver, degeneration of testes, and subcutaneous edema,and, in some cases, general edema in the body occurs. Deficiency oflinoleic acid in the male can impair spermatogenesis and affect fertility(NRC, 1994). Linoleic acid-deficient chicks are more susceptible to respiratory infections (Scott et al., 1982). High mortality resulted from an
atypical respiratory infection for chicks fed linoleic acid-deficient dietsfrom hatching to 10 to 12 weeks of age.
Fritsche et al. (1991) reported that feeding laying hens a diet rich inω-3 PUFA (7% fish oil) significantly enhanced their primary antibody
response and altered lymphocyte proliferation. Feeding broiler chickens
diets rich in ω-3 PUFA reduced antibody-dependent cell cytotoxicity andaltered eicosanoid release by chicken immune cells (Fritsche and Cassity,1992).
Linoleic acid deficiency in laying hens results in depressed egg production, small egg size, a slight reduction in fertility, and a marked increase in early mortality of the embryo during incubation. Eggs fromhens severely deficient in linoleic acid will not hatch. For severely deficient hens it is necessary to feed pullets diets that are very low in linoleic
acid from hatching on. Linoleic acid is stored in the body for long peri699

CHAPTER EIGHTEEN

ods by animals reared on a diet containing adequate linoleic acid.

Other Animal Species
DOGS AND CATS
Dogs deficient in EFA have low growth rates, a characteristic dermatitis, swelling and redness of the paws, and increased susceptibility toinfection (NRC, 1985a). Beagle puppies fed a low-fat diet exhibited skinlesions within 2 to 3 months.
Clinical signs attributed to EFA deficiency in cats include listlessness; dry, unattractive hair coats; and severe dandruff (NRC, 1986).
Growth is poor, and susceptibility to infections is increased. Histological examination of the liver reveals fatty infiltration and parenchymal
disorganization. Both males and females lack libido, testes are underdeveloped, and estrus cycles are absent.

FISH
In common with other vertebrates, fish cannot synthesize either
18:2ω-6 or 18:3ω-3 de novo. One or both of these fatty acids must be
supplied preformed in the diet, depending on the EFA requirements. Fishvary considerably in their ability to convert 18-carbon unsaturated fatty
acids to longer-chain more highly unsaturated fatty acids of the same series. In general, freshwater fish require either dietary linoleic acid(18:2ω-6) or linolenic acid (18:3ω-3) or both, whereas marine fish require dietary EPA (20:5ω-3) and/or DHA (22:6ω-3).
The principal gross signs of EFA deficiency reported for various
fishes are dermal signs (fin rot), a shock syndrome, myocarditis, reduced
growth rate, reduced feed efficiency, and increased mortality (Castell etal., 1972; NRC, 1993). Essential fatty acid deficiency has also beenshown to reduce the reproductive performance of common carp, rainbow trout, and Red Sea bream (NRC, 1993).
Deficiency signs specific to trout are poor growth, elevated tissuelevels of ω-9 fatty acids (particularly 20:3ω-9), necrosis of the caudal fin,fatty pale liver, dermal depigmentation, increased muscle water content,
syncope accentuated by stress, increased mitochondrial swelling, increased respiration rate of liver homogenates, heart myopathy, and lowered hemoglobin level. Poor growth, low feed efficiency, high mortality,
and swollen pale livers were reported in chum salmon fed an EFA-deficient diet (NRC, 1993).
In certain warm-water fish—common carp, for example—it wasdemonstrated that dietary levels of 22:6ω-3 significantly affected egg
700

E S S E N T I A L FAT T Y A C I D S

hatchability (NRC, 1993). Shrimp would not produce eggs unless thediet contained 20:5ω-3 and 22:6ω-3. Watanabe et al. (1983) reported
that ω-3 PUFAs, such as EPA and DHA, are required for normal growthand development of ayu and Red Sea bream larvae. High mortalities andabnormalities, such as underdeveloped swim bladder and scoliosis, havebeen observed in Red Sea bream larvae.

FOXES
Foxes on low-EFA diets showed clinical signs of hyperkeratosis anddandruff (NRC, 1982a).

LABORATORY ANIMALS
Signs of EFA deficiency in the rat are reduction in growth (whichplateaus after about 12 to 18 weeks); scaly skin; rough, thin hair coat;necrosis of the tail; electrocardiographic abnormalities; fatty liver; impaired reproduction; and death. Many other less noticeable but equally
severe changes have been reported, including kidney lesions and a decrease in urine volume, lipid-containing macrophages in the lung, increased metabolic rate, decreased capillary resistance, and aberrant ventricular conduction (NRC, 1995).
During fetal growth and postnatal development, large amounts of
ω-3 and ω-6 are deposited in nervous system tissues, particularly in thecentral nervous system in rats. In rats, inadequate 22:6ω-3 supply tothese tissues during fetal development and postnatal life is associatedwith abnormal retinal function, visual acuity, and behavior (Enslen etal., 1991).
Newborns from rats fed linoleic acid-deficient diets exhibit the most
severe signs of deficiency and usually die within 3 days to 3 weeks after
birth, depending on the duration of the EFA feeding to the dam. In
young rats fed a fat-free diet, two of the earliest signs of EFA deficiency
are increased transepidermal water loss and increased urinary argininevasopressin excretion (Hansen and Jensen, 1986).
Mice with EFA deficiency have hair loss, dermatitis with scaling andcrusting of skin, and occasional diarrhea. Deficiency in older micecaused infertility without visible skin changes (NRC, 1995). In the deficient guinea pig, there is weight loss, with clinical signs including dermatitis; skin ulcers; fur loss; underdevelopment of spleen, testes, andgallbladder; and enlargement of kidneys, liver, adrenals, and heart.

NONHUMAN PRIMATES
In monkeys, EFA deficiency resulted in dryness and scaliness of the701

CHAPTER EIGHTEEN

skin, with loss of hair, although many months of the deficient diet wererequired for appearance of clinical signs (NRC, 1978).

RABBITS
Signs in rabbits of EFA deficiency are reduced growth, hair loss, degenerative changes in seminiferous tubules, and impaired sperm development (Ahluwalia et al., 1967).

HumansEssential fatty acid deficiency in humans has primarily been studied
in infants. Eczema has been reported in a number of studies of infants
maintained on a low-fat diet (Holman, 1978a). Hansen et al. (1958) presented data showing that young healthy infants, within a relatively shorttime, may develop symptoms when given diets extremely low in fat. After several weeks on trial, alterations in the skin were observed in the
majority of infants. The first sign detected was dryness, then thickening,
and later desquamation, with oozing in the intertriginous folds (Fig.
18.5). Addition of linoleic acid, as 2% of the caloric intake, restored theskin to normal, while the addition of saturated fatty acid had no effect.
Patients who are maintained by fat-free total parenteral nutrition(TPN) are candidates for EFA deficiency. Premature infants receiving
TPN are particularly susceptible to rapid development of EFA deficiency
signs (Cooke et al., 1984).
The need for ω-3 fatty acids in human diets has been established (seeMetabolism and Functions). Potential benefits of ω-3 fatty acids havebeen found to reduce thrombosis of platelet aggregation and heart disease, as well as hypertension. There are numerous reports of potentiallybeneficial effects of fish oil (ω-3 fatty acids) on triglycerides, blood rheology, blood pressure, and inflammation. Studies have also revealed an
anticancer effect (e.g., colon cancer) of supplemental ω-3 found in fishoils (Cave, 1991).
Essential fatty acid deficiency has been mostly related to linoleic
acid deficiency, but in most situations linolenic acid deficiency coexists(Uauy and de Andraca, 1995). One study found that ω-3 fatty acids—
especially docosahexaenoic acid (22:6ω-3), present in human milk—arenecessary for retinal and brain development in primates and humans(Uauy and Hoffman, 1991). Rod photoreceptor function and the maturation of visual acuity of human low birth weight infants are dependent
on the supply of these essential nutrients (Birch et al., 1992).

Assessment of Status
702

A

B
Fig. 18.5 Skin lesions associated with EFA deficiency in (A) an adult and (B)an infant before and after EFA administration. The adult received fat-free total
parenteral nutrition. (Courtesy of R.T. Holman, The Hormel Institute, University of Minnesota, and M.C. Riella, J.W. Broviac, M. Wells, and B.H. Scribner(1975), Ann. Intern. Med. 83:786–789.)
703

CHAPTER EIGHTEENIn addition to various deficiency clinical signs and responses to EFA
supplementation, the most accurate indicators of EFA status are biochemical changes associated with EFA deficiency. Anatomical signs of
EFA deficiency vary from species to species, but biochemical aberrations
associated with deficiency in rats are found to be the same in all otherspecies studied (Holman, 1978a). The biochemical signs of linoleic aciddeficiency have been known for many years. The chief alterations in various tissues are decreased levels of linoleic (18:2ω-6), arachidonic(20:4ω-6), and docosapentaenoic (22:5ω-6) acids, and increased levelsof eicosatrienoic (20:3ω-9) and docosatrienoic (22:3ω-9) acids. With the
EFA-deficient diet the tissues attempt to produce unsaturated acids fromthe oleic acid (18:1ω-9), which results in the accumulation of thetrienoic acid, eicosatrienoic acid (20:3ω-9). The triene:tetraene ratio is
the ratio of the abnormally elevated endogenous metabolite of oleic
acid, 20:3ω-9, to the metabolic product, 20:4ω-6, from the dietary essential linoleic acid (see Requirements). Therefore, it is the ratio of the“abnormal” to “normal” major polyunsaturated acids in tissue lipids.
The curve describing the triene-tetraene ratio versus dietary linoleate
had a sharp break at or near 1% of calories, suggesting that 1% of calories of linoleic acid may be a critical amount, the dietary requirement(Holman, 1960). Values of the ratio below 0.4 have been considered
normal for animals. Fig. 18.6 is an example of a study in which six levels of linoleic acid were fed to young male pigs (Sewell and McDowell,1966). As dietary linoleic acid was increased, arachidonic acid (20:4ω6) was elevated and 20:3ω-9 was suppressed. Castell et al. (1972) suggested that the ratio of 20:3ω-9:20:5ω-3 in polar lipids from the liver ofrainbow trout might be a useful index of linolenic acid status.

SUPPLEMENTATION
Linoleic acid deficiency is not likely to develop when diets containappreciable amounts of corn. Yellow corn is the major source of linoleic
acid in most feed formulas for swine and poultry. Diets composed ofcorn and soybean meal with no further supplementation are likely to beadequate in linoleic acid for chick growth but marginal for maximum
egg size. Since linoleic acid has a marked effect on egg size, it is necessary to ensure that a sufficient amount of linoleic acid is included in thediet of laying hens to enable them to lay eggs of maximum size as earlyas possible during the egg production year (Scott et al., 1982). A typicaldietary linoleic acid recommendation for poultry is 1% of total calories.

704

Total Fatty Acids (%)

E S S E N T I A L FAT T Y A C I D S

Dietary Linoleic Acid (% Calories)

Fig. 18.6 Influence of varying levels of dietary linoleic acid on linoleic (18:2 ω6), arachidonic (20:4 ω-6), eicosatrienoic (20:3 ω-9), and oleic (18:1 ω-9) acidcontent of testes tissue from young swine. (Modified from Sewell and McDowell, 1966.)

However, for laying and breeding hens the requirement would be 1.4%of calories until egg size is reached (Scott et al., 1982).
Animals fed diets containing sorghum, barley, or wheat instead ofcorn as the major grain may receive suboptimal quantities of linoleic
acid. Even more important, when roots and tubers (such as potatoes andcassava) or processed carbohydrates (such as cane sugar) make up a major part of the energy source, and when solvent-extracted protein supplements are used exclusively, there is potential danger of skin lesions related to linoleic acid deficiency (McDowell, 1977). For young
ruminants, the main supplementation concern is that milk replacers contain adequate concentrations of EFA.
Supplementing cattle with fish oils or protected fat may have a beneficial reproduction response. Typically, unsaturated fatty acids are biohydrogenated by ruminal microorganisms, therefore preventing their delivery as-is to the lower gut for absorption. However, eicosapentaenoic(20:5ω-3) and docosahexaenoic (22:6ω-3) fatty acids found in fish oil

705

CHAPTER EIGHTEEN

appear to escape biohydrogenation (Ashes et al., 1992; Palmquist andKinsey, 1994). Therefore, feeding fish meal may result in uptake of thesefatty acids for metabolism by reproductive tissues of the lactating cow(Smith and Marnett, 1991; Garcia-Bojalil, 1993) and result in higher reproductive rates.
Under typical conditions, EFA deficiency would not be expected inhumans. The average fat content of U.S. diets is 40% of calories. Likewise, there has been a marked shift in preference from animal fat to vegetable oils, which contain high quantities of linoleic acid. Even if all dietary fat were from one source, it is unlikely that EFA content of the dietwould be inadequate. Because polyunsaturated acids of the ω-6 family
are nearly ubiquitous in plants and animals, and in most natural foodsources they constitute more than the minimum nutrient requirement,even random selection of foods is not likely to induce EFA deficiency(Holman, 1978a).
Research suggests that human health could be improved with cisversus trans-fatty acids and conjugated linoleic acid. Trans-fatty acids(hydrogenated fats) raise the undesirable LDL cholesterol and depress
the favorable HDL cholesterol (Katan and Mensink, 1992). Conjugatedlinoleic acid from ruminant animal products have been shown to havean anticarcinogen effect (Belury, 1995).
The main concern for supplementation of EFA for humans would be
for infants not receiving mother’s milk, but rather a milk product low in
EFA. Particular attention is needed for premature infants to ensure thatparenteral nutrition includes formulas containing adequate EFA. Visualfunction of full-term infants fed human milk is enhanced for up to 3
years, supporting the concept of long-term benefits of human milk feeding on mental development (Birch et al., 1993). Docosahexaenoic acid
(22:6ω-3) is particularly required for brain development. Low birthweight infants randomized to human milk or formula tube feeding
demonstrated the benefits of human milk for IQ at age 8 years (Lucas etal., 1992). Plasma 22:6ω-3 concentrations of full-term infants fed formula are lower than that of breast-fed infants. This suggests that present formulas provide insufficient linolenic acid, or chain elongation-desaturation enzymes are not sufficiently active during early life to support
tissue accretion of 22:6ω-3. Also, studies of infants who were born atterm and died from sudden infant death syndrome revealed that brain
composition is affected by human milk feeding. The data indicated thathigher 22:6ω-3 content was found in the brain cortex of breast-fed infants relative to infants receiving formulas based on cow’s milk (Farquharson et al., 1992).
706

E S S E N T I A L FAT T Y A C I D S

Long-chain polyunsaturated fatty acids are being included in infant
formulas. It is clear that the balance between 20:4ω-6 and 22:6ω-3 inthe diet is a powerful determinant of the level of these fatty acids in thedeveloping brain. Addition of 20:4ω-6 or 22:6ω-3 alone may be inappropriate; balanced addition of both ω-6 and ω-3 long-chain PUFAs(20:4ω-6 and 22:6ω-3) seems to be required (Jumpsen et al., 1997).
Chylomicronemia (increased fat particles in blood) in patients withsevere familiar hypertriglyceridemia has been shown to respond to longterm treatment with ω-3 fatty acids (Pschierer et al., 1995). Long-term
supplementation of a diet with ω-3 fatty acids results in a persistent decrease of serum triglyceride concentrations, a pronounced reduction in
the number of chylomicrons present in the fasting state, and an improvement in plasma viscosity.
For human diets, increasing attention is being paid to the dietaryrole of long-chain ω-3 PUFAs (e.g., 20:5ω-3 and 22:6ω-3). Researchsuggests that marine oil fatty acids (linolenic or ω-3 family) play a rolein ameliorating thrombosis, heart disease, hypertension, inflammation(e.g., arthritis), and some types of cancer, and in improving the immune
response. As a result, increased dietary intake of ω-3 fatty acids has
been recommended as a means of reducing risk for coronary heart disease (Kromhout et al., 1985; Nair et al., 1999; Tinker et al., 1999).
Table eggs enriched with ω-3 fatty acids provide an alternative to fish
as a source of these proposed healthful fatty acids (Marshall et al.,1994). Inclusion of various fish oils, seeds, and seed oils in layer rations
readily results in the incorporation of up to 220 mg of ω-3 fatty acids
per egg yolk (Hargis and Van Elswyk, 1993). This level of ω-3 fatty
acids is equivalent to that which would be consumed in a 100-g (3.5oz.) serving of lean fish; therefore, consumption of one egg enrichedwith ω-3 fatty acids could replace a serving of fish (Marshall et al.,1994).
In swine, feeding diets containing canola or fish oils has resulted inincreased tissue levels of the ω-3 family of fatty acids (Leskanich et al.,
1997; Soler-Velásquez et al., 1998). Menhaden fish oil at 7% in a sow’slate-gestation and lactation diet greatly elevated the content of ω-3 fatty
acids in the nursing pig immune cells (Fritsche et al., 1993). Likewise,
the desire to manipulate the fatty acid composition of ruminant tissues
is motivated by human health concerns about saturated fatty acids in animal food products and by the emerging roles of unsaturated fatty acids
as regulators of cell function. In cattle (Fotouhi and Jenkins, 1992) andsheep (Jenkins, 1995), the reaction of unsaturated fatty acids with primary amines produced fatty acyl amides that resisted biohydrogenation707

CHAPTER EIGHTEEN

and caused less disruption of ruminal fermentation. In dairy cattle, conversion of soybean oil to butylsoyamide protected unsaturated fatty
acids from ruminal biohydrogenation, causing linoleic acid to increasein the plasma and milk of dairy cows (Jenkins et al., 1996).

TOXICITY
There are nutritional disadvantages from excessive intakes of EFAs.
Before the role of antioxidants and the dietary requirement for vitamin
E were understood, a large amount of literature accumulated concerning the alleged toxicity of polyunsaturated fats (Holman, 1978a). These
readily oxidized acids increase the requirement (see Chapter 4) for vitamin E, which functions as an antioxidant in the body. Several experiments have shown that levels of the vitamin that were normally sufficient to prevent vitamin E deficiency signs such as muscular dystrophyand encephalomalacia proved inadequate as the intakes of EFAs were increased. It is difficult to experimentally separate the effects of high levels of PUFAs from a relative deficiency of tocopherol, but it is obvious
that, at least under some circumstances, high levels of PUFAs may have
undesirable effects.
Not only are excess PUFAs detrimental to vitamin E, but individual
fatty acids are detrimental to other fatty acids. Excess fatty acids of one
family will suppress the metabolism of other families of fatty acids(Hwang et al., 1988). Suggested favorable ω-6:ω-3 fatty acid ratios forhumans are 4 to 10:1, respectfully. Increased ω-3 to result in a 1.4 ω6:l.0ω-3 ratio has been shown to depress the cell-mediated immune response and PGE2α production in dogs (Whelan, 1996; Wander et al.,1997).

■ REFERENCES
Aaes-Jørgensen, E. (1982) Prog. Lipid Res. 20, 123.
Aaes-Jørgensen, E., and Dam, H. (1954) Brit. J. Nutr. 8, 290.
Ahluwalia, B., Pincus, G., and Holman, T. (1967) J. Nutr. 92, 205.
Anonymous (1985). Nutr. Rev. 43(11), 350.
Ashes, J.R., Siebert, B.D., Gulati, S.K., Cuthbertson, A.Z., and Scott, T.W.
(1992) Lipids 27, 629.
Ayre, K.J., and Hulbert, A.J. (1996) J. Nutr. 126, 653.
Belury, M.A. (1995) Nutr. Rev. 53, 83.
Birch, E.E., Birch, D.G., Hoffman, D.R., and Uauy, R.D. (1992) Ophthal. Vis.
Sci. 33, 3242.
Birch, E.E., Birch, D.G., Hoffman, D.R., Hale, L., Everett, M., and Uauy, R.D.
708

E S S E N T I A L FAT T Y A C I D S(1993) J. Pediatr. Ophthalmol. Strabismus 30, 33.
Boissonneault, G.A., and Johnston, P.V. (1983) J. Nutr. 113, 1187.
Bourre, J.M., Francois, M., Youyou, A., Dumont, O., Piciotti, M., Pascal, G.,
and Durand, G. (1989) J. Nutr. 119, 1880.
Bourre, J.M., Piciotti, M., Dumont, O., Pascal, G., and Durand, G. (1990)
Lipids 25, 465.
Bourre, J.M., Dumont, O., Pascal, G., and Durand, G. (1993) J. Nutr. 123,
1313.
Bruckner, G., Gurnewalk, K.K., Tucker, R.E., and Mitchell Jr., G.E. (1984) J.
Anim. Sci. 58, 971.
Burke, J.M., Staples, C.R., Risco, C.A., and Thatcher, W.W. (1996) In 7th Annual Florida Ruminant Nutrition Symposium, p. 21. University of Florida,
Gainesville.
Burr, G.O., and Burr, M.M. (1929) J. Biol. Chem. 82, 345.
Burr, G.O., and Burr, M.M. (1930) J. Biol. Chem. 86, 587.
Burr, M.L., Fehily, A.M., and Gilbert, J.F. (1989) Lancet 2, 757.
Calder, P.C. (1998). Nutr. Rev. 56, S 70.
Carlson, S.E., Carver, J.D., and House, S.G. (1986) J. Nutr. 116, 718.
Castell, J.D., Sinnhuber, R.O., Wales, J.H., and Lee, D.J. (1972) J. Nutr. 102, 77.
Cave, W.T. Jr. (1991) FASEB J. 5, 2160.
Chin, S.F., Liu, W., Storkson, J.M., Ha, Y.L., and Pariza, M.W. (1992) J. Food
Comp. Anal. 5, 185.
Chin, S.F., Storkson, J.M., Liu, W., Albright, K.J., and Pariza, M.W. (1994) J.
Nutr. 124, 694.
Chouinard, P.Y., Corneau, L., Barbano, D.M., Metzer, L.E., and Bauman, D.E.
(1999). J. Nutr. 129, 1579.
Connor, W.E., Neuringer, M., and Reisbick, S. (1992) Nutr. Rev. 50, 21.
Cooke, R.J., Zee, P., and Yeh, Y.Y. (1984) J. Pediatr. Gastroenterol. Nutr. 3, 446.
Crawford, M.A., Doyle, W., Drury, P., Ghebremeskel, K., Harbige, L., Leyton,
J., and Williams, G. (1989) In Dietary ω3 and ω6 Fatty Acids. Biological
Effects and Nutritional Essentiality (C. Galli and A.P. Simopoulos, eds.), p.
5. Plenum Press, New York.
Cunningham, H.M., and Loosli, J.K. (1954a) J. Anim. Sci. 13, 265.
Cunningham, H.M., and Loosli, J.K. (1954b) J. Dairy Sci. 37, 453.
Delion, S., Chalon, S., Hérault, J., Guilloteau, D., Besnard, J.C., and Durand, G.
(1994) J. Nutr. 124, 2466.
Dormandy, T.L., and Wickens, D.G. (1987) Chem. Phys. Lipids 45, 353.
Drevon, C.A. (1992) Nutr. Rev. 50, 38.
Dupont, J. (1990). In Present Knowledge in Nutrition (M.L. Brown, ed.), 6thed., p. 56. International Life Science Institute, Washington, D.C.
Enslen, M., Milon, H., and Malnoe, A. (1991) Lipids, 26, 203.
Evans, H.M., and Burr, G.O. (1926) Proc. Soc. Exp. Biol. Med. 24, 740.
Farquharson, J., Cockburn, F., and Ainslie, P.W. (1992) Lancet 340, 810.
Fernandes, G. (1995) Nutr. Rev. 53, 572.
Fernandes, G., and Venkatraman, J.T. (1993) Nutr. Res. 13, 519.
Fotouhi, N., and Jenkins, T.C. (1992) J. Anim. Sci. 70, 3607.
Fritsche, K.L., and Cassity, N.A. (1992) Poult. Sci. 71, 1646.
Fritsche, K.L., Cassity, N.A., and Huang, S.C. (1991) Poult. Sci. 70, 611.
709

CHAPTER EIGHTEEN

Fritsche, K.L., Alexander, D.W., Cassity, N.A., and Huang, S. (1993). Lipids 28,
677.
Garcia-Bojalil, C.M. (1993) Reproductive, Productive and Immunological Responses of Holstein Dairy Cows Fed Diets Varying in Concentration and
Ruminal Degradability of Protein and Supplemented with Ruminally Inert
Fat. Ph.D. Dissertation, University of Florida, Gainesville.
Garton, G.A. (1960) Nature 187, 511.
Goodwin, J.S. (1985) Prostaglandins and Immunity. Martinus Nijhoff, Boston,
Massachusets.
Greenberg, S.M., Colbert, C.E., Savage, E.E., and Deuel, J. (1950) J. Nutr. 41,
473.
Gullickson, T.W., Fountaine, F.C., and Fitch, J.B. (1942) J. Dairy Sci. 25, 117.
Ha, Y.L., Grimm, N.K., and Pariza, M.W. (1987) Carcinogenesis 8, 1881.
Hansen, A.E., Haggard, M.E., Boelsche, A.N., Adam, D.J., and Wiese, H.F.
(1958) J. Nutr. 66, 565.
Hansen, H.S. (1994) Nutr. Rev. 52, 162.
Hansen, H.S., and Jensen, B. (1986) J. Nutr. 116, 198.
Hargis, P.S., and Van Elswyk, M.E. (1993) Poult. Sci. 70, 874.
Hoek, J., and Rubin, E. (1990) Alcohol 25, 143.
Hofstetter, H., Sen, N., and Holman, R.T. (1965) J. Am. Oil Chem. Soc. 42,
537.
Holman, R.T. (1960) J. Nutr. 70, 405.
Holman, R.T. (1964) Fed. Proc. Am. Soc. Exp. Biol. 23, 1062.
Holman, R.T. (1968) Prog. Chem. Fats Other Lipids 9, 279.
Holman, R.T. (1978a) In Handbook Series in Nutrition and Food, Section E:
Nutrition Disorders, Volume 3 (M. Rechcigl Jr., ed.), p. 491. CRC Press,
West Palm Beach, Florida.
Holman, R.T. (1978b) In Handbook Series in Nutrition and Food, Section E:
Nutrition Disorders, Volume 3 (M. Rechcigl Jr., ed.), p. 335. CRC Press,
West Palm Beach, Florida.
Holman, R.T. (1986). J. Am. Coll. Nutr. 56, 303.
Holman, R.T., Johnson, S.B., and Kokmen, E. (1989). Proc. Natl. Acad. Sci. U
S A 86, 4720.
Hume, E.M., Nunn, L.A., Smedley-MacLean, I., and Smith, H.H. (1940).
Biochem. J. 34, 879.
Hwang, D.H., Boudreau, M., and Chanmugam, P. (1988). J. Nutr. 118, 427.
Jenkins, T.C. (1995). J. Anim. Sci. 73, 818.
Jenkins, T.C., Bateman, H.G., and Block, S.M. (1996). J. Dairy Sci. 75, 585.
Jiang, J., Bjoerck, L., Fonden, R., and Emanuelson, M. (1996). J. Dairy Sci. 79,
438.
Jumpsen, J., Lien, E.L., Goh, Y.K., and Clandinin, M.T. (1997). J. Nutr. 127,
724.
Katan, M.B., and Mensink, R.P. (1992). Nutr. Rev. 50, 46.
Kinsella, J.E., Lokesh, B., Broughton, K.S., and Whelan, J. (1990). Nutrition 5,
24.
Knazek, R.A., and Liu, S.C. (1979). Proc. Soc. Exp. Biol. Med. 162, 346.
Kromhout, D., Bosschieter, E.B., and Coulander, C. deL. (1985). N. Engl. J.
Med. 312, 1205.
710

E S S E N T I A L FAT T Y A C I D S

Lambert, M.R., Jacobson, N.L., Allen, R.S., and Zaletel, J.H. (1954). J. Nutr.
52, 259.
Lamptey, M.S., and Walker, B.L. (1976). J. Nutr. 106, 86.
Lands, W.E.M. (1986). Nutr. Rev. 44, 189.
Leaf, A. (1992). Nutr. Rev. 50, 150.
Leat, W.M. (1962). Brit. J. Nutr. 16, 559.
Leskanich, C.O., Matthews, K.R., Warkup, C.C., Noble, R.C., and Huzzledine,M. (1997). J. Anim. Sci. 75, 673.
Lichtenstein, A. (1993). Nutr. Rev. 51, 340.
Lin, H., Boylston, T.D., Chang, M.J., Luedecke, L.O., and Shultz, T. (1995). J.
Dairy Sci. 78, 2358.
Lucas, A., Morley, R., Cole, T.J., Lister, G., and Leeson-Payne, C. (1992). Lancet
339, 261.
Lucy, M.C., Gross, T.S., and Thatcher, W.W. (1990). In Livestock Reproductionin Latin America, p. 119. Atomic Energy Agency, Vienna.
Lucy, M.C., Staples, C.R., Michel, F.M., and Thatcher, W.W. (1991). J. Dairy
Sci. 74, 483.
Machlin, L.J., and Gordon, R.S. (1961). J. Nutr. 75, 157.
Marshall, A.C., Kubena, K.S., Hinton, K.R., Hargis, P.S., and Van Elswyk, M.E.
(1994). Poult. Sci. 73, 1334.
McDowell, L.R. (1977). Geographical Distribution of Nutritional Diseases inAnimals. Department of Animal Science, University of Florida, Gainesville.
Mead, J.F. (1982). Prog. Lipid Res. 20, 1.
Mohrhauer, H., and Holman, R.T. (1963). J. Lipid Res. 4, 151.
Morrow, J.D., Minton, T.A., Badr, K.F., and Roberts, L.J. (1994). Biochim. Biophys. Acta 1210, 244.
Murphy, M.G. (1990). J. Nutr. Biochem. 1, 68.
Nair, S.S.D., Leitch, J., Falconer, J., and Garg, M.L. (1999). J. Nutr. 129, 1518.
Neuringer, M., Connor, W.E., Van Petten, C., and Barstad, L. (1984). J. Clin. Invest. 73, 272.
Neuringer, M., Anderson, G.J., and Connor, W.E. (1988). Annu. Rev. Nutr. 8,
517.
NRC. Nutrient Requirements of Domestic Animals. National Academy of Sciences-National Research Council, Washington, D.C.
(1978). Nutrient Requirements of Nonhuman Primates.
(1982a). Nutrient Requirements of Mink and Foxes.
(1985a). Nutrient Requirements of Dogs, 2nd Ed.
(1986). Nutrient Requirements of Cats, 3rd Ed.
(1989a). Nutrient Requirements of Dairy Cattle, 6th Ed.
(1989b). Nutrient Requirements of Horses, 5th Ed.
(1993). Nutrient Requirements of Fish.
(1994). Nutrient Requirements of Poultry, 9th Ed.
(1995). Nutrient Requirements of Laboratory Animals, 2nd Ed.
NRC. (1982b). United States-Canadian Tables of Feed Composition, 3rd Ed.
National Academy of Sciences-National Research Council, Washington,
D.C.
Ollivier-Bousquet, M. Guesnet, P., Seddiki, T., and Durand, G. (1993). J. Nutr.
123, 2090.
711

CHAPTER EIGHTEEN

Palmquist, D.L., and Kinsey, D.J. (1994). J. Dairy Sci.(Suppl. 1) 77, 350(Abstr.).
Palmquist, D.L., Mattos, W., and Stone, R.L. (1977). Lipids 12, 235.
Parodi, P.W. (1999). J. Dairy Sci. 82, 1339.
Peifer, J.J., and Holman, R.T. (1955). Arch. Biochem. Biophys. 57, 520.
Pepe, S., and McLennan, P.L. (1996). J. Nutr. 126, 34.
Pschierer, V., Richter, W.O., and Schwandt, P. (1995). J. Nutr. 125, 1490.
Pudelkewicz, C., Seuffert, J., and Holman, R.T. (1968). J. Nutr. 94, 138.
Reiser, R. (1950). J. Nutr. 42, 319.
Sanders, K.M. (1994). Feed Management 45(4), 53.
Schlenk, H., and Gellerman, J.L. (1965). J. Am. Oil Chem. Soc. 42, 504.
Scott, N.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Sewell, R.F., and McDowell, L.R. (1966). J. Nutr. 89, 64.
Shantha, N.C., Decker, E.A., and Ustunol, Z. (1992). J. Am. Oil Chem. Soc. 69,
425.
Smith, W.L., and Marnett, L.J. (1991). Biochim. Biophys. Acta, 1083, 1.
Soler-Velásquez, M.P., Brendemuhl, J.H., McDowell, L.R., Sheppard, K.A.,
Johnson, D.D., and Williams, S.N. (1998). J. Anim. Sci. 76, 110.
Sosenko, I.R.S. (1995). J. Nutr. 125, 1652S.
Swanson, R.L., Baumgardner, C.A., and Geer, B.W. (1995). J. Nutr. 125, 553.
Tinker, L.F., Parks, E.J., Behr, S.R., Schneeman, B.O., and Davis, P.A. (1999). J.
Nutr. 129, 1126.
Topping, D. (1993). Nutr. Rev. 51, 271.
Turek, J.J., Schoenlein, I.A., Clark, L.K., and Van Alstin, W.C. (1994). J.
Leukoc. Biol. 56, 599.
Turek, J., Schoenlein, A., Watkins, A., Van Alstine, G., Clark, L., and Knox, K.
(1996). J. Nutr. 126, 1541.
Uauy, R., and de Andraca, I. (1995). J. Nutr. 125, 2278S.
Uauy, R., and Hoffman, D.R. (1991). Semin. Perinatol. 15, 449.
Vergroesen, A.J. (1977). Nutr. Rev. 35, 1.
Viscardi, R.M. (1995). J. Nutr. 125, 1645S.
Wander, R.C., Hall, J.A., Gradin, J.L., Du, S.H., and Jewell, D.E. (1997). J.
Nutr. 127, 1198.
Watanabe, T., Kitajima, C., and Fujita, S. (1983). Aquaculture 34, 115.
Whelan, J. (1996). J. Nutr. 126, 1086.
Willett, W.C., Stampfer, M.J., Colditz, G.A., Rosner, B.A., and Speizer, F.E.
(1990). N. Engl. J. Med. 323, 1664.
Willis, A.L. (1984). In Nutrition Reviews, Present Knowledge in Nutrition (R.E.
Olson, ed.), p. 90. The Nutrition Foundation, Inc., Washington, D.C.
Witz, W.M., and Beeson, W.M. (1951). J. Anim. Sci. 10, 112.
Zerouga, M., Beauge, F., Niel, E., Durand, G., and Bourre, J.M. (1991).
Biochim. Biophys. Acta 1086, 295.

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chapter nineteen

V I TA M I N
S U P P L E M E N TAT I O N

INTRODUCTION
In the early part of this century, pasture, other forages, distiller’s solubles or grains, brewer’s grains, fermentation products, and meat, milk,
and fish by-products were depended on as sources of the vitamins foranimal feeding. As animal feeding became more sophisticated, as fastergrowing and higher-producing animals were developed, and as the trend
toward more intensified operations occurred, it became necessary to add
an increasing number of vitamins to properly fortify animal diets.
Vitamins represent only a minute fraction of animal feeds, typicallyamounting to only 0.05% by weight and 1 to 2% of feed cost for swine
and poultry operations depending on the diet used and the level of supplementation required (McNaughton, 1990). Yet a balanced vitaminfortification program for meeting requirements of nonruminant animals
under a wide range of feeds and different production systems will morethan offset the cost of adding vitamins.
Because of proper diet selection and vitamin supplementation, human deficiency diseases attributed to vitamin A (xerophthalmia andnight blindness), vitamin D (rickets), vitamin C (scurvy), thiamin(beriberi), niacin (pellagra), and vitamin B12 (pernicious anemia) havebeen eliminated in varying degrees in developing countries but still posea problem for susceptible groups. Vitamin A deficiency particularly is
still a problem in many world areas. In human nutrition, vitamin supplementation should be provided to susceptible population groups, withthe precaution of avoiding quantities in extreme excess, in particular vitamin A.
Supplementation guidelines specific for each vitamin have been presented in the respective chapters (Chapters 2 through 18). The present
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chapter will discuss general supplementation considerations for both animals and humans.

FACTORS RESULTING IN INADEQUATE DIETARYINTAKES OF VITAMINS
Vitamin dietary intake and utilization is influenced by many factors,
including particular feed ingredients, bioavailability, harvesting, processing, storage, feed intake, antagonists, least-cost feed formulations,and other factors (NRC, 1973; McDowell, 2000).

Agronomic Effects and Harvesting Conditions
Vitamin levels will vary in feed ingredients because of crop location,
fertilization, plant genetics, plant disease, and weather. Intensive cropping practices and use of new crop varieties may result in reduced levelsof certain vitamins in many feedstuffs. In forage crops, factors that favor production of lush, green plants also favor production of many vitamins, particularly β-carotene, vitamin E, and vitamin K. The vitamin
C content of tomatoes depends primarily on intensity of sunlight striking the fruits of the tomato during the immediate preharvest period(Scott, 1973).
Harvesting conditions often play a major role in the vitamin content
of many feedstuffs. Vitamin content of corn is drastically reduced whenharvest months are not conducive to full ripening. If corn has been subjected to alternate periods of freezing and thawing while it contains ahigh amount of moisture, fermentation occurs in corn kernels, and there
is a loss of vitamin content, particularly of vitamin E and cryptoxanthin.
In one study, vitamin E activity in blighted corn was 59% lower than insound corn, and activity of the vitamin in lightweight corn averaged
21% lower than in sound corn (Hoffmann-La Roche, 1991). Young etal. (1975) reported that the rate of oxidation of natural tocopherol was
higher in high-moisture corn than in low-moisture corn due to increased
peroxidation of the lipid. Certain legumes, particularly alfalfa and soybeans, contain the enzyme lipoxidase that, unless quickly inactivated,
readily destroys much of the carotenes.

Processing and Storage Effects
Many vitamins are delicate substances that can suffer loss of activity due to unfavorable circumstances encountered during processing or
storage of premixes and feeds. Stress factors for vitamins include humidity, pressure (pelleting), friction (abrasion), heat, light, oxidation-re714

V I TA M I N S U P P L E M E N TAT I O N

duction, rancidity, trace minerals, pH, and interactions with other vitamins, carriers, enzymes, and feed additives (NRC, 1973).
Humidity is the primary factor that can decrease the stability of vitamins in premixes and feedstuffs. Water softens the matrix, for example, of vitamin A, thus the vitamin becomes more permeable to oxygen.
Trace elements, acids, and bases are activated only by water. Humidity
augments the negative effects exerted by choline chloride, trace elements, and other chemical reactions that are not found in dry feed.
Thus, the water level is responsible for a higher reactivity of vitaminswith other feed components. Elevated moisture content or incorrect
storage of premixes and feedstuffs is the root of almost all stability problems. Christian (1983) determined the stability of vitamin A in a premix.
After 3 months of storage, the vitamin A retention was 88% under low
temperature and low humidity, 86% under high temperature and low
humidity, and 2% under high temperature and high humidity. They concluded that humidity was significantly more stressful than temperature.
Vitamins that undergo friction or are mixed and stored with minerals are subject to loss of potency. Friction is an important factor because
it erodes the coating that protects several vitamins and reduces vitamin
crystals to a smaller particle size. Friction is very high in pelleting. Someabrasion is inevitable in the mixing process, but fortunately, most minerals contain little moisture, with the exception of salt, which is, ofcourse, somewhat hygroscopic if exposed to environmental moisture.
Therefore, packaging, careful transport, and storage become important,and few companies would willingly use very high levels of salt in a supplement containing fat-soluble vitamins.
Hazards to vitamins from minerals are abrasion and direct destruction by certain trace elements, particularly copper, zinc, and iron; manganese and selenium are the least reactive. Free metal ion is the most reactive (metal filings), followed by sulfate, carbonate, oxide, and the least
reactive form is chelated. Chelated mineral forms become incapable ofinitiating formation of free radicals. In fat-soluble vitamins, esters are
significantly more stable than alcohols. The hydroxy group of alcohols
is extremely sensitive to oxidation. The five double bonds in retinyl acetate still make the compound sensitive to oxidation. Vitamin A is significantly more stable in vitamin premixes than in vitamin-trace mineralpremixes because trace minerals catalyze oxidation of the five double
bonds (Coelho, 1996). Dove and Ewan (1986) determined the stabilityof α-tocopherol in feeds without and with trace minerals. At the end of
3 months of storage at 25 to 30oC, α-tocopherol retention was 50 and
30%, respectively. The addition of 245 ppm copper as copper sulfate715

CHAPTER NINETEEN

produced 0% retention after 15 days. In addition to trace minerals,choline chloride is highly destructive to vitamins (see Chapter 14) andshould not be included in a vitamin premix.
Some vitamins are destroyed by light. Riboflavin is stable to mostfactors involved in processing; however, it is readily destroyed by either
visible or ultraviolet light. Vitamin B6, vitamin C, and folacin can also
be destroyed by light. It is necessary, therefore, to protect premixes offeeds containing these vitamins from light and radiation (Stamberg andPeterson, 1946).
Sun-field curing of cut hay is essential to provide vitamin D activity
but results in loss of other vitamin potency. Mangelson et al. (1949)
showed that mechanical dehydration at 177oC within 1 hour after cutting produced an alfalfa meal that contained 2.5 times more carotene as
did sun-cured alfalfa. There was no loss of riboflavin, pantothenic acid,
niacin, or folacin during dehydration. Field-cured alfalfa was lower inriboflavin, and when alfalfa was exposed to rain, there was a large lossof pantothenic acid and niacin (Scott, 1973).
Dehydration of alfalfa at 135oC resulted in an average of 18% lossof α-tocopherol. When dehydrated alfalfa meal was stored for 12 weeks
at 32oC, the α-tocopherol loss averaged 65% (Livingston et al., 1968).
Corn is often dried rapidly under high temperatures, resulting in lossesof vitamin E activity and other heat-sensitive vitamins. When corn was
artificially dried for 40 minutes at 88oC, losses of α-tocopherol averaged
19%, and when corn was dried for 54 minutes at 107oC, losses averaged
41% (Adams, 1973).
While pelleting generally improves the value of energy and protein
carriers in a feed, this is not true for some vitamins. During pelleting offeeds, four elements destructive for a number of vitamins are applied in
combined action: friction, heat, pressure, and humidity. Increasing thepelleting temperature or conditioning time generally enhances redox reactions and destroys vitamins. As with friction, not all vitamins or formsof vitamins respond to these factors equally. Significant amounts of thevitamin E in the alcohol form and ascorbic acid, but very little of the vitamin E in the acetate form or choline, are destroyed as pelleting temperatures and conditioning times increase. Gadient (1986) reported thatvitamins A, D3, K3, C, and thiamin are most likely to show stability
problems in pelleted feeds. Pelleting of feed may have a beneficial effecton availability of vitamins such as niacin and biotin, which are often
present in bound forms (Scott, 1973). Feed manufacturers have increased pelleting temperatures for all animal feeds in order to control

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Salmonella organisms and increase digestibility, and are using steam pelleting, prepelleting conditioners, and feed expanders, which lead to increased vitamin degradation (Coelho, 1994).
In extrusion, the dominant stress factors are pressure, heat, moisture, and redox reactions. Extrusion is the most aggressive processagainst vitamins due to the high temperature (107 to 135oC), pressure(400 to 1,000 psi), and moisture (30%) involved in the process (Coelho,1996).
Processing with the goal of producing better-quality fish meals andfish solubles under conditions in which putrefaction is prevented has resulted in lower levels of vitamin K and vitamin B12 in these feedstuffs
than were present when the products were allowed to undergo a considerable degree of putrefaction. Early studies on vitamin K and vitamin
B12 proved that fish meal and rice bran exposed to the action of microorganisms showed increased content of these vitamins. On the other
hand, processing of many raw fishes with heat is required to inactivate
a potent thiaminase that destroys thiamin. Also, attempts to preservefish with nitrates led to the production of carcinogenic nitrosamines.
Each stage of human food preparation and storage results in vitamin loss. Fruits and vegetables that are harvested long before use undergo heavy vitamin losses by enzymatic decomposition. Vitamin C isparticularly liable to this type of destruction. In apples stored under domestic conditions, the vitamin C content may fall to about one-third of
the original value after only 2 or 3 months. Blanching vegetables before
canning or freezing also results in vitamin loss. Estimates suggest thatlosses due to blanching fluctuate between 13 and 60% for vitamin C, 2
and 30% for thiamin, and 5 and 40% for riboflavin (Marks, 1975). Vitamin losses during heat sterilization are generally small because oxygenis excluded during this process. Thiamin is the vitamin most susceptible
because of its labile nature in heat, and considerable thiamin losses havebeen observed in meat. Irradiation results in damage to vitamins, ofwhich the most sensitive vitamins are thiamin, riboflavin, vitamin A,
and vitamin E; niacin is relatively stable. Storage in cans, freezing, anddehydration are all relatively good methods of preservation concerning
vitamin retention.

Reduced Feed Intake
When feed intake is reduced, vitamin allowances should be adjusted
to ensure adequate vitamin intake for optimum performance. Restricting feed intake practices and/or improved feed conversion will decrease717

CHAPTER NINETEEN

dietary intake of all nutrients, including vitamins. Restricted feeding ofbroiler breeders, turkey breeder hens, and gestating sows and gilts may
result in marginal vitamin intake if diets are not adequately fortified(Hoffmann-La Roche, 1989; McDowell, 2000). Reduced feed intake
may also result from stress and disease.
Use of high-energy feeds such as fats to provide diets with greater
nutrient density for higher animal performance requires a higher vitaminconcentration in feeds. Nonruminant species provided diets ad libitum
consume quantities sufficient to meet energy requirements. Thus, vitamin fortification must be increased for high-energy diets because animals will consume less total feed. Feed consumption was compared in
broilers receiving metabolizable energy ranging from 2,800 to 3,550kcal/kg of feed (Friesecke, 1975). Feed and vitamin consumption were
each 19.1% lower in broilers consuming the diet with greater energy
density compared to those consuming the lowest-energy diet.
Ambient temperature also has an important influence on diet consumption, as animals consume greater quantities during cold temperatures and reduced amounts as a result of heat stress. Vitamins, as well asother nutrients, must therefore be adjusted to reflect changing dietaryconsumption.

Vitamin Variability and Insufficient Analysis
Tables of feed and food composition demonstrate the lack of complete vitamin information, with vitamin levels varying widely within agiven feedstuff. Kurnick et al. (1972) found that, of the feeds surveyed,
information on the niacin and riboflavin content of feedstuffs was more
complete than for any other vitamins, whereas values for vitamin B12and vitamin K were most deficient. Thus, 2 to 30% of the ingredients
lacked niacin, riboflavin, or pantothenic acid values, while 89 to 97%
did not have values for carotene, vitamin B12, or vitamin K. Information
about the other vitamins was not listed for 36 to 64% of the ingredients.
In more than 25 years since this report, the situation has not greatly improved; vitamin analyses of feeds are still woefully inadequate.
Variability of vitamin content within ingredients is generally large
and difficult to quantify and anticipate. It is well recognized that vitamin levels shown in tables of vitamin composition of feedstuffs represent
average values and that actual vitamin content of each feedstuff varies
over a fairly wide range. Vitamin content of feed ingredients differs drastically from sample to sample; therefore, at least two standard deviations (average, 10%) need to be deducted from the initial content. Sev-

718

V I TA M I N S U P P L E M E N TAT I O Neral months elapse between harvest, processing, and finally consumptionof feed ingredients. Since most natural vitamins are not chemically protected in feed ingredients, there is considerable loss between harvesting
and consumption (average, 25% loss) (BASF, 1991).
Methods of processing and storage, as previously mentioned, account for variability as well as different analysis techniques. Often, it isquestionable whether the accuracy of vitamin levels from feedstuffs, calculated using tabular values, can be ensured (Kurnick et al., 1972).
Proof of this is the statement by the 1982 NRC publication on U.S.Canadian Tables of Feed Composition that “. . . organic constituents(e.g., crude protein, cell wall constituents, ether extract, and aminoacids) can vary as much as ±y15%, the inorganic constituents as much
as ±y30%, and the energy values as much as ±y10%.” Therefore, average values in feed composition tables may vary considerably from thenutrient value in a specific group of feeds.

Vitamin Bioavailability
Even accurate feedstuff analyses of vitamin concentrations do notprovide bioavailability data needed for certain vitamins. Bound forms ofvitamins in natural ingredients often are unavailable to animals.
Bioavailability of choline, niacin, and vitamin B6 is adequate in somefeeds but limited or variable in others. For example, bioavailability ofcholine is 100% in corn but varies from 60 to 75% in soybean meal;that of niacin is 100% in soybean meal but zero in wheat and sorghum
and varies from 0 to 30% in corn; that of vitamin B6 is 65% in soybean
meal and varies from 45 to 56% in corn (Hoffmann-La Roche, 1991).
The niacin in cereal grains and their by-products is in a bound form,
which is virtually unavailable for the pig and chick (Cunha, 1982). For
alfalfa meal, corn, cottonseed meal, and soybean meal, bioavailability ofbiotin is estimated at 100% (Cunha, 1984a). However, biotin availability is variable for other feedstuffs, for example, 20 to 50% in barley,
62% in corn gluten meal, 30% in fish meal, 20 to 60% in sorghum,
32% in oats, and 0 to 62% in wheat. Likewise, ascorbic acid in cookedcabbage is present in the bound form, ascorbinogen, a form that is absorbed very poorly by humans (Marks, 1975).
Some data obtained for the pig showed that responses to vitamins
may differ depending on whether vitamins are being added to a purified
or natural diet (Cunha, 1977). Requirements of the pig for niacin, riboflavin, and pantothenic acid were considerably higher on a natural
diet than requirements established earlier from experiments using puri-719

CHAPTER NINETEEN

fied diets (McMillen et al., 1949). This shows that results obtained withpurified diets must also be verified with natural diets and that bioavailability of vitamins may be greater in purified diets.

Computerized Least-Cost Feed Formulations
Vitamins are not usually entered as specifications in computerized
feed formulations. Therefore, vitamin-rich feedstuffs—such as alfalfa,
distiller’s solubles or grains; brewer’s grains; fermentation products; andmeat, milk, and fish by-products—are often excluded or reduced whenleast-cost feed formulations are computed. The resulting least-cost dietconsisting of a grain and soybean meal is usually lower in vitamins thana more complex one containing more costly vitamin-rich feeds (Roche,1979).

FACTORS AFFECTING VITAMIN REQUIREMENTS
AND UTILIZATION
Physiological Makeup and Production Function
Vitamin needs of animals and humans depend greatly on their physiological makeup, age, health, and nutritional status and function (e.g.,
producing meat, milk, eggs, hair or wool, or developing a fetus) (Roche,1979). For example, dairy cows producing greater volumes of milk havehigher vitamin requirements than dry cows or cows producing low
quantities. Breeder hens have higher vitamin requirements for optimum
hatchability, since vitamin requirements for egg production are generally
less than for egg hatchability. Higher levels of vitamins A, D3, and E areneeded in diets for breeder hens than in feeds for rapidly growing broilers. Selection for faster growth rate may allow animals to reach muchhigher weights at much younger ages, with less feed consumed. DudleyCash (1994) concluded that since genetic potential has improved at therate of 0.8% feed conversion yearly, and most of the NRC vitamin requirement data are 20 to 40 years old, vitamin requirements determinedseveral decades ago may not apply to today’s poultry. Selection for fasterweight gains in swine and increased number of litters per year also demands elevated vitamin requirements (Cunha, 1980a, 1984b).
Different breeds and strains of animals have been shown to vary intheir vitamin requirements. Vitamin needs of new strains developed for
improved production are higher. Leg problems seen in fast-growing
strains of broilers can be corrected in part by higher levels of biotin, folacin, niacin, and choline (Roche, 1979).
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Confinement Rearing Without Access to Pasture
Moving swine and poultry operations into complete confinement
without access to pasture has had a profound effect on vitamin nutrition(as well as mineral nutrition). Pasture could be depended on to provide
significant quantities of most vitamins, since young, lush, green grasses
or legumes are excellent vitamin sources. More available forms of vitamins A and E are present in pastures and green forages, which containample quantities of β-carotene and α-tocopherol versus lower bioavailable forms in grains. Confinement rearing to include poultry in cages
and swine on slatted floors allows limited animal access to feces (coprophagy), which is rich in many vitamins. Confinement rearing requires producers to pay more attention to higher vitamin requirements
needed when this management system is used (Cunha, 1984b).

Stress, Disease, or Adverse Environmental Conditions
Intensified production increases stress and subclinical disease level
conditions because of higher densities of animals in confined areas.
Stress and disease conditions in animals may increase the basic requirement for certain vitamins. A number of studies have indicated that nutrient levels that are adequate for growth, feed efficiency, gestation, andlactation may not be adequate for normal immunity and for maximizing
the animal’s resistance to disease (Cunha, 1985; Nockels, 1988). Diseases or parasites affecting the gastrointestinal tract will reduce intestinal absorption of vitamins, both from dietary sources and those synthesized by microorganisms. If they cause diarrhea or vomiting, this will
also decrease intestinal absorption and increase needs. Vitamin A deficiency is often seen in heavily parasitized animals that supposedly werereceiving an adequate amount of the vitamin. Mycotoxins are known tocause digestive disturbances such as vomiting and diarrhea, as well as internal bleeding, and to interfere with absorption of dietary vitamins A,
D, E, and K. In broiler chickens, moldy corn (mycotoxins) has been associated with deficiencies of vitamins D (rickets) and E (encephalomalacia) in spite of the fact that these vitamins were supplemented at levels
regarded as satisfactory.
In recent years, a malady dubbed “spiking syndrome” in broilers
caused a sharp rise in mortality at about 14 days of age. Some nutritionists feel that this problem may be associated with Fusarium mycotoxins, although the exact cause is not clearly defined. Increased levelsof thiamin ameliorate the rise in mortality, and it has been suggested thatwhen either corn quality is poor or mycotoxin levels and/or mold counts
721

CHAPTER NINETEEN

are high, thiamin should be increased by 1.11 to 1.65 mg/kg in thestarter feed (Coelho, 1995).
Mortality from fowl typhoid (Salmonella gallinarum) was reducedin chicks fed vitamin levels greater than normal (Hill, 1961). Vitamin E
supplementation at a high level decreased chick mortality due to Escherichia coli challenge from 40 to 5% (Tengerdy and Nockels, 1975).
Scott et al. (1982) concluded that coccidiosis produces a triple stress onvitamin K requirements as follows: (1) Coccidiosis reduces feed intake,thereby reducing vitamin K intake; (2) coccidiosis injures the intestinal
tract and reduces absorption of the vitamin; and (3) treatment with sulfaquinoxaline or other coccidiostats causes an increased requirement forvitamin K.

Vitamin Antagonists
Vitamin antagonists (antimetabolites) interfere with the activity ofvarious vitamins, and Oldfield (1987) summarized the action of antagonists. The antagonist could cleave the metabolite molecule and render
it inactive, as occurs with thiaminase and thiamin; it could complex withthe metabolite, with similar results, as happens between avidin and biotin; or, by reason of structural similarity, it could occupy reaction sites
and thereby deny them to the metabolite, as with dicumarol and vitamin
K. The presence of vitamin antagonists in animal and human dietsshould be considered in adjusting vitamin allowances, as most vitamins
have antagonists that reduce their utilization (see Chapters 2 through18). Some common antagonists are as follows:
1. Thiaminase, found in raw fish and some feedstuffs, is a thiamin antagonist. Pyrithiamin is another thiamin antagonist.
2. Dicumarol, found in certain plants, interferes with blood clotting by
blocking the action of vitamin K.
3. Avidin, found in raw egg white, and streptavidin, from Streptomyces
molds, are biotin antimetabolites.
4. Rancid fats inactivate biotin and destroy vitamins A, D, and E andpossibly others.
5. Oral contraceptives and drug therapy to control tuberculosis are antagonistic to vitamin B6.

Use of Antimicrobial Drugs
Some antimicrobial drugs will increase vitamin needs of animals byaltering intestinal microflora and inhibiting synthesis of certain vitamins. Certain sulfonamides may increase requirements of biotin, folacin,722

V I TA M I N S U P P L E M E N TAT I O N

vitamin K, and possibly others when intestinal synthesis is reduced. Thismay be of little significance except when drugs that are antagonistic toward a particular vitamin are added in excess, that is, sulfaquinoxaline
versus vitamin K, amprolium versus thiamin, and sulfonamide potentiators versus folacin (Perry, 1978).

Levels of Other Nutrients in the Diet
Level of fat in the diet may affect absorption of the fat-soluble vitamins A, D, E, and K, as well as the requirement for vitamin E and possibly other vitamins. Fat-soluble vitamins may fail to be absorbed if digestion of fat is impaired. The high cost of fat as an energy source has
resulted in minimal fat levels in least-cost feed formulations, which may
result in reduced absorption of fat-soluble vitamins (Hoffmann-La
Roche, 1991).
Many interrelationships of vitamins with other nutrients exist andtherefore affect requirements. For example, prominent interrelationships
exist for vitamin E with selenium, vitamin D with calcium and phosphorus, choline with methionine, and niacin with tryptophan.

Body Vitamin Reserves
Body storage of vitamins from previous intake will affect daily requirements of these nutrients. This is truer for the fat-soluble vitamins
A, D, and E and for vitamin B12 than for the other water-soluble vitamins and vitamin K. Vitamin A may be stored by an animal in its liverand fatty tissue in sufficient quantities to meet requirements for up to 6
months or even longer.

OPTIMUM VITAMIN ALLOWANCESThe National Research Council (NRC) requirements for a vitaminare usually close to minimum levels required to prevent deficiency signs
and for conditions of health and adequate performance, provided sufficient amounts of all other nutrients are supplied. Vitamin requirementsfor various animal species (NRC) and humans (RDA, 1989) are presented in each chapter and in the appendix, Table A1. Most nutritionists usually consider NRC requirements for vitamins to be close to minimum requirements sufficient to prevent clinical deficiency signs, andthey may be adjusted upward according to experience within industry insituations in which a higher level of vitamins is needed.
Allowances of a vitamin are those total levels from all sources fed tocompensate for factors influencing vitamin needs of animals. These in723

CHAPTER NINETEEN

fluencing factors include (1) those that may lead to inadequate levels of
the vitamin in the diet (see Factors Resulting in Inadequate Dietary Intakes of Vitamins) and (2) those that may affect the animal’s ability toutilize the vitamin under commercial production conditions (see Factors
Affecting Vitamin Requirements and Utilization). The higher the allowance, the greater the extent to which it may compensate for the influencing factors. Thus, under commercial production conditions, vitamin allowances higher than NRC requirements may be needed to allow
optimum performance (Roche, 1979).
The optimum supplementation level is the vitamin concentrationthat achieves the best growth rate, feed utilization, and health (including
immune competency) and provides adequate body reserves (Coelho,1996).
The concept of optimum vitamin nutrition under commercial production conditions is illustrated in Fig. 19.1 (Roche, 1979). The marginal zone represents vitamin levels that are lower than requirementsthat may predispose animals to deficiency. The requirements zone represents minimum vitamin quantities that are needed to prevent deficiency signs, but may lead to suboptimum performance even though animals appear normal. The allowances in the optimum zone permit
animals to achieve their full genetic potential for optimum performance.
In the excess zone, vitamin levels range from levels still safe, but uneconomical, to concentrations that may produce toxic effects. Usually only
vitamins A and D, under practical feeding conditions, pose the possibility of toxicity problems for livestock. Optimum allowances of any vitamin are depicted as a range in Fig. 19.1 because factors influencing vitamin needs are highly variable, and optimum allowances to allow
maximum response may vary from animal to animal of the same species,
type, and age within the same population and from day to day (Roche,1979).
It should be emphasized that subacute deficiencies can exist although the actual deficiency signs do not appear. Such borderline deficiencies are both the most costly and the most difficult to cope with andoften go unnoticed and unrectified, yet they may result in poor and expensive gains, impaired reproduction, or depressed production. Also,
under farm conditions, one will usually not find a single vitamin deficiency. Instead, deficiencies are usually a combination of factors, and often deficiency signs will not be clear cut. If the NRC minimum requirement for a vitamin is the level that barely prevents clinical deficiency
signs, then this level moves in relationship to the level required for optimum production responses. This means that if a greater quantity of a vi724

V I TA M I N S U P P L E M E N TAT I O N

Fig. 19.1 Optimum vitamin nutrition for animals under commercial production conditions. Allowances are for total vitamin levels from all sources fed tocompensate for factors affecting animals’ vitamin needs. (Adapted from
Roche, 1979. Printed with permission of Hoffmann-La Roche Inc., Nutley,
New Jersey.)

tamin is required for an optimum response (because of the influencing
factors), a greater quantity would also be required to prevent deficiency
signs (Fig. 19.2). Similarly, if a lesser quantity is required for an optimum response, less would also be required to prevent deficiencies (Perry,1978). Optimum animal performance required under modern commercial conditions cannot be obtained by fortifying diets to just meet minimum vitamin requirements. Establishment of adequate margins of safety
must provide for those factors that may increase certain dietary vitaminrequirements and for variability in active vitamin potencies and availability within individual feed ingredients.
The NRC requirements often do not take into account that certain
vitamins have special functions in relation to disease conditions withhigher than the recommended levels needed for response (Cunha, 1985).
In pigs artificially infected with Treponema hyodysenteriae, the agent
causing diarrhea, high supplementation with vitamin E (200 mg/day) in
combination with selenium (0.2 mg/day) markedly reduced the numberof pigs that became clinically ill (Tiege et al., 1978). Clinical signs andpathological changes were less severe compared with pigs with vitamin725

CHAPTER NINETEEN

Fig. 19.2 Optimum vitamin nutrition for animals under commercial production conditions and influencing factors. (Adapted from Perry, 1978. Printed
with permission of Hoffmann-La Roche Inc., Nutley, New Jersey.)E deficiency. Thus, high doses of vitamin E increase resistance against
disease. In practice, feeds contaminated with mycotoxins increase requirements for fat-soluble and other vitamins (e.g., biotin, folacin, andpossibly others); therefore, supplementation should be increased above
NRC minimum requirements. Apart from these fat-soluble vitamins, addition of folacin also improved performance in pigs fed moldy grain(Purser, 1981), and of biotin in pigs fed feeds containing certain molds(Cunha, 1984a). Besides other nutrients, vitamins play a major role in
the immune response, the body’s defense system against infectious disease. Vitamin supplementation above requirements was shown to be required for optimum immune responses (Ellis and Vorhies, 1976; Cunha,
1985; Weiss, 1998).
In relation to poultry, Kennedy et al. (1992) suggested that the NRC
vitamin requirements have essentially been the same for the last 40
years. The industry vitamin average allowances have increased significantly (30 to 500%) to keep pace with greater genetic potential, faster
growth rates, better feed efficiency, poorer quality ingredients, larger
726

V I TA M I N S U P P L E M E N TAT I O N

poultry houses, and generally higher disease levels, all of which caused
increased stress in one study (McNaughton, 1990). A reasonableamount of logic would suggest that vitamin requirements determinedseveral decades ago may not apply to today’s poultry feeds (DudleyCash, 1994).
Dritz et al. (1995) fed sows higher vitamin concentrations than
NRC requirements, with the result that 0.1 more pigs were born alive,
0.2 more pigs were weaned per litter, and weaned pigs were 1.1 kg heavier than controls. Stahly (1994) fed growing pigs under two management schemes to create a moderate or high level of antigen exposure.
Within each antigen group, pigs were fed one of five dietary concentrations of B vitamins (niacin, riboflavin, pantothenic acid, B12, or folacin).
The addition of B vitamin concentrations of 370 to 470% above the current NRC (1998) recommendation resulted in 21% greater body weightgains and 10% improvement in feed utilization for pigs in either antigen
group.
Interest has increased regarding the elimination of vitamin supplementation from poultry and swine diets for varying lengths of time.
Skinner et al. (1991), for example, found that broiler performance was
not impaired when vitamin supplementation was deleted from the dietfor the last 21 days of the feed period. In contrast with these results,
however, Gwyther et al. (1992) reported that NRC vitamin recommendations were much too low to maintain broiler performance. This latterstudy indicated that broiler vitamin requirements exceed those recommended by the NRC, and that the elimination of vitamin supplementation from broiler diets would severely impair performance. Teeter andDeyhim (1996) eliminated vitamins and/or minerals from broiler diets
for the last 21 days of life, during which time the birds were exposed toheat stress. There was significant reduction in live bird and carcass performance. Performance tended to be poorest when the diet contained
added trace minerals and no added vitamins, suggesting oxidation of thevitamins already present.

VITAMIN SUPPLEMENTATION MOST NEEDED BYLIVESTOCK
Vitamin requirements, as previously noted, are highly variable
within the various species and classes of animals. Supplementation allowances need to be set at levels that reflect different management systems and are high enough to take care of fluctuations in environmental
temperatures, energy content of feed, or other factors that might influ727

CHAPTER NINETEEN

ence feed consumption or the vitamin requirements in other ways
(McGinnis, 1986; McDowell, 1998).
A new concept of vitamin supplementation for both livestock andhumans relates to providing antioxidant vitamins (carotenoids, vitamin
E, and vitamin C) in excess of supposed dietary requirements (see Chapters 2, 4, and 15). These nutrients play important roles in animal healthby inactivating harmful free radicals produced through normal cellular
activity and from various stressors. Both in vitro and in vivo studies
showed that these antioxidant vitamins generally enhance different aspects of cellular and noncellular immunity. A compromised immune system will result in reduced animal production efficiency through increased susceptibility to diseases, thereby leading to increased animal
morbidity and mortality.
The following section will briefly discuss vitamins that are normallyprovided in ruminant, poultry, swine, and horse diets (McDowell,
2000).

Ruminants
Grazing ruminants generally need supplemental vitamin A only if
pastures are low in carotene and possibly vitamin E (influenced by selenium status). Vitamin D is provided by ultraviolet light activity on theskin, while all other vitamins are provided by ruminal or intestinal, microbial, or tissue synthesis.
Ruminants housed under more strict confinement conditions generally require vitamins A and E and may require vitamin D if they are deprived of sunlight. Additional supplemental vitamin E would be needed
to stabilize the meat color of finishing animals. Under specific conditions
relating to stress and high productivity, ruminants may benefit from supplemental B vitamins, particularly thiamin and niacin. Future research
may find a need for biotin and carnitine supplementation.
Adding a complete B-vitamin mixture to cattle entering the feedlot
during the first month can reduce stress and increase gains. In one study,
supplemental B vitamins given to feedlot calves tended to reduce morbidity of animals (Zinn et al., 1987). Apparently, under stress conditionsof feedlots, the microbial population in the rumen is not synthesizing
certain B vitamins at adequate levels.
Steer feedlot calves were feed-restricted and infected with bovine
herpes virus-1 (BHV-1), with one treatment group receiving injections of
B vitamins (Dubeski et al., 1996a). The IgG titers tended to be higher forthe calves receiving B-vitamin injections, indicating that the humoral immune response was enhanced by B-vitamin treatment. B-vitamin status728

V I TA M I N S U P P L E M E N TAT I O N

in stressed calves at the time of vaccination or disease challenge may affect the success of the immune response. The mild respiratory (BHV-1)
infection in steer calves markedly decreased plasma concentrations of vitamin B6, vitamin B12, pantothenic acid, and ascorbate but not folacin(Dubeski et al., 1996b). Depletion of B-vitamin and ascorbate statusduring shipping and marketing may contribute to the enhanced susceptibility of cattle to infectious disease during the first few weeks after arrival at feedlots.

Poultry
Poultry managed under intensive production systems are particularly susceptible to vitamin deficiencies (Scott et al., 1982). Reasons forthis susceptibility are as follows: (1) Poultry derive little or no benefit
from microbial synthesis of vitamins in the gastrointestinal tract, (2)poultry have high requirements for vitamins, and (3) the high-density
concentration of modern poultry operations places many stresses on thebirds that may increase their vitamin requirements. Typical grain-oilseed
meal (e.g., corn-soybean meal) poultry diets are generally supplemented
with vitamins A, D, D3, E, and K; riboflavin; niacin; pantothenic acid;
B12; and choline (Scott et al., 1982). Thiamin, vitamin B6, biotin, and folacin are usually present in adequate quantities in the major ingredients,
such as corn-soybean meal-based diets. Carnitine may be found to be ofvalue in future studies.
Riboflavin and vitamins A, D, and B12 are usually low in poultry diets. However, adding other vitamins to poultry diets is good insurance.
Vitamins D and B12 are almost completely absent from diets based oncorn and soybean meal. Vitamin K is generally added to poultry diets
more than to those for other species, because birds have less intestinalsynthesis due to a shorter intestinal tract and faster rate of food passage.
Birds in cages require more dietary K and B vitamins than those on floor
housing because of more limited opportunity for coprophagy.

Swine
Vitamin supplementation of swine diets is obviously necessary (Figs.
19.3 and 19.4) because vitamin needs have become more critical as complete confinement feeding has increased. Swine in confinement, without
access to vitamin-rich pasture, and housed on slatted floors, which limits vitamins available from feces consumption, have greater need forsupplemental vitamins. The vitamins most likely to be marginal or deficient in corn-soybean diets are vitamins A, D, E, B12, riboflavin, niacin,
pantothenic acid, and occasionally vitamin K and choline.
729

CHAPTER NINETEEN

Fig. 19.3 Small, weak pigs at birth due to a lack of B vitamins. (Courtesy of T.
J. Cunha and University of Florida.)

Almost all swine diets in the United Stated are now fortified with vitamins A, D, E, and B12; riboflavin; niacin; pantothenic acid; andcholine. An increasing number of feed manufacturers are adding vitamin
K, and many are adding biotin and B6 to diets. Diets are fortified withthese vitamins even though not all experiments indicate a need for eachof them. Most feed manufacturers add them as a precaution to take careof stress factors, subclinical disease level, and other conditions on the
average farm that may increase vitamin needs (Cunha, 1977). It appears
that carnitine supplementation of weanling pigs has potential (Newton
and Burtle, 1992).

HorsesThere is a lack of experimental information on the level of vitaminsrequired in well-balanced horse diets, as well as on which vitamins need
to be added (Cunha, 1980b). The vitamins most likely to be deficient inall classes of horses are vitamins A and E, with vitamin D also deficientin horses in confinement. Inadequate vitamin D may be provided to
racehorses that are exercised only briefly in the early morning, whensunlight provides less antirachitic protection. Requirements for vitamins
A, D, and E can be met with a high-quality (e.g., green) sun-cured hay.
Deficiencies of vitamin K and the B vitamins appear to be less likely in730

V I TA M I N S U P P L E M E N TAT I O NABFig. 19.4 The effect of B-vitamin supplementation on deficient pigs obtained
from farms in Michigan. (A) These pigs were about 80 days old and weighed
an average of 9.1 kg. (B) The same pigs after 35 days of supplementation.
(Courtesy of R.W. Luecke, Michigan State University.)

the mature horse than in other monogastric species, as many vitamins
occur in the cecum of the horse. It is not known, however, how much of
the vitamins synthesized in the cecum is absorbed in the large intestine.
Since it is difficult to depend on the intestinal synthesis, many horseowners use B-vitamin supplementation of diets for young horses and for
those being developed for racing or performance purposes (Cunha,1991).
Vitamin supplementation has become more critical for horses as thetrend toward total confinement has increased. Currently, few horses receive a high level of vitamin intake from a lush, green pasture or from a731

CHAPTER NINETEEN

high-quality leafy, green hay. Cunha (1980b) suggested that a vitaminpremix for horses contain vitamins A, D, E, K, B6, and B12; thiamin; riboflavin; niacin; pantothenic acid; folacin; and choline. Biotin supplementation is also recommended since Comben et al. (1984) showed abenefit of the vitamin for hoof integrity. More recently, it has been suggested that carnitine supplementation is beneficial for horses. Vitamin E
was found to have an effect on the susceptibility of horses to exerciseinduced muscle damage (BASF, 1997a). Dietary levels of vitamin E
greater than 80 IU/kg, and potentially 300 IU/kg, are required to maintain the blood and muscle vitamin E concentrations in horses undergoing exercise conditioning.

VITAMIN SUPPLEMENTATION FOR HUMANS
People in developed countries consume an average of 3,000 calories
and almost 40 g of protein daily, while those in developing countries
consume two-thirds as many calories and only one-fifth as much protein. Thus, calorie and protein deficiency are the most important problems in nutrition. In developing countries, vitamin deficiencies usuallycoexist with protein-calorie deficiency. In developed countries, on theother hand, inappropriate eating patterns can lead to poor nutrition andresult in vitamin deficiencies (Marks, 1975).
A number of factors influence the vitamin deficiency state in humans. Diseases caused by primary or dietary deficiency result solely
from inadequate quantities of the vitamin in the diet. Lack of vitamin intake can result from a number of factors (Marks, 1975), including crop
failure, losses during food preparation and storage, poverty, ignorance,loss of appetite, apathy, food taboos or fads, dental problems, andchronic disease. Even social changes can affect dietary intake. For example, old people, particularly those living alone, tend to abandon ordinary meal preparation and live on small amounts of so-called easy
foods (coffee, soup, and bread) that can lead to scurvy and other deficiency diseases.
However, some vitamin deficiencies occur even under conditions inwhich the diet would normally be considered adequate by most standards. Such deficiencies, usually referred to as secondary or conditioned
deficiencies, arise from metabolic stress or organic disease. The increased metabolic demands during pregnancy and lactation are primeexamples of metabolic stress states requiring extra vitamin therapy. Useof oral contraceptives and of various drugs, such as tranquilizers and antibiotics, can result in conditioned deficiencies. Particular attention to vi732

V I TA M I N S U P P L E M E N TAT I O N

tamin supplementation is needed for individuals with inborn errors of
metabolism, for whom requirements are much higher for certain vitamins, and for hospital patients receiving total parenteral nutrition.
Certain disorders or diseases and some medications may interferewith vitamin intake, digestion, absorption, metabolism, or excretion andthus change requirements. The stress placed on carbohydrate metabolism in the alcoholic is another example of a condition requiring morethan normal daily vitamin consumption. A number of B-vitamin deficiencies, including those of thiamin, niacin, B6, B12, and folacin, have resulted from alcoholism. For example, a major cause of folacin deficiency
in the United States is chronic alcoholism. Diets of alcoholics are likely
to contain inadequate quantities of a number of nutrients.
Another important cause of conditioned deficiency results from disease. In these cases a defect in the host results in decreased absorptionof the vitamin from the diet. Pernicious anemia is a conditioned vitamin B12 deficiency because a lack of gastric intrinsic factor severely limits absorption of the vitamin from the gastrointestinal tract. Anotherexample of this type is vitamin K deficiency that results from poor absorption and is caused by inadequate supplies of bile. Vitamin therapy
in such cases is usually administered parenterally (e.g., intravenous injection).
Typical well-balanced diets that include ample quantities of vegetables, fruits, and animal products should provide vitamins in adequate
quantities. For most individuals consuming such diets, there should be
no need for supplemental vitamins. Nevertheless, many circumstances—including individual variations, lack of proper diet, older age,and physiological and emotional stress—may warrant vitamin supplementation. Many individuals also consume multiple vitamins (andsometimes minerals) daily, which gives them a sense of well-being andin essence is a type of nutritional insurance against possible deficiencies.
In addition to vitamin deficiencies, it is also important to consider possible effects of excessive quantities of vitamins (hypervitaminosis) inhuman diets, particularly regarding vitamin A and, to a lesser degree,
vitamin D.
The antioxidant nutrients—vitamin E, vitamin C, and β-carotene—
have become the focus for their protective role in disease prevention,
particularly cardiovascular disease (atherosclerosis) and cancer. Thesenutrients play important roles in health by inactivating harmful free radicals produced through normal cellular activity and from various stressors. Both in vitro and in vivo studies have shown that these antioxidant
vitamins generally enhance different aspects of cellular and noncellular733

CHAPTER NINETEEN

immunity. Many individuals, particularly those in more developed countries, take pharmacological levels of the antioxidant vitamins as well as
the antioxidant mineral selenium. This trend is likely to continue because health benefits will continue to be amplified as more and morestudies validate the health benefits of these nutrients.
The vitamins least likely to be needed for supplementation in humandiets are choline, pantothenic acid, and biotin. Those that warrant supplementation are the following:
1. Vitamin A—Deficiency of vitamin A is widespread in the world andis most common in young children who receive inadequate green and yellow
vegetables and foods derived from animals.
2. Vitamin D—In developed countries there is vitamin D fortificationof human diets; however, most of the world depends on sunlight. Children
not receiving sufficient sunlight are at great risk of deficiency, and older populations need to consider supplementation. Decreased ability of the kidney
to convert 25-OHD to 1,25-(OH)2D is a problem in the elderly (Tsai et al.,1984). Thus, even with more sunlight and more dietary intake, it is possiblethat elderly individuals would not maintain equivalent activity of 1,25(OH)2D. Diseases and parasites that impair the ability of the liver and kidney to convert vitamin D to 1,25-(OH)2D may also be problems, especiallyin developing countries.
3. Vitamin E—This vitamin is most likely deficient in individuals consuming low-fat diets or diets high in unsaturated fats and would be morecritical in regions deficient in selenium, as vitamin E supplementation is beneficial for diverse disease conditions.
4. Vitamin K—Newborns are commonly given a single dose of vitamin
K to prevent abnormal bleeding. Many persons take aspirin for pain, tominimize coronary problems, and for other reasons. There are indications
that this may increase vitamin K needs.
5. Thiamin—Individuals subsisting mostly on refined grains (e.g., polished rice) are most likely in need of thiamin supplementation. In developed
countries, people have for a number of years received food fortified with thiamin (e.g., bread enrichment). In Far Eastern countries, rice enriched withthiamin has dramatically reduced the incidence of beriberi.
6. Riboflavin—Often, individuals in developing countries do not havethe economic resources needed to consume major dietary sources of riboflavin, such as meat, milk, and dairy products.
7. Niacin—Supplementation is important when diets are primarily
based on corn and sorghum and for individuals with metabolic defects (e.g.,conversion of tryptophan to niacin) or when grain has not been heated or734

V I TA M I N S U P P L E M E N TAT I O N

treated sufficiently to make niacin available. Pharmacological effects haveincluded reduction of serum cholesterol and treatment of schizophrenia.
8. Vitamin B6—Supplementation is most important for young, pregnant, or lactating women. Requirements are increased by oral contraceptives, certain drug therapy, radiation sickness, urinary calculi, and in errorsof metabolism. Vitamin B6 has been used to alleviate nausea during pregnancy and in megadose therapy for a wide variety of conditions, including
premenstrual syndrome and behavioral disorders.
9. Folacin—A high percentage of the human population is estimatedto be at a moderate to high risk for folacin deficiency (see Chapter 12) during growth, pregnancy, and old age. Supplementation is particularly needed
for individuals not consuming optimum quantities of fruits and vegetables.
To prevent certain birth defects, starting in 1998 certain foods (e.g., cereals
and bread) in the United States were required by law to contain supplemental folacin.
10. Vitamin B12—Organ meats and other animal products are the bestsources of vitamin B12. Vegetarians as well as individuals with the malabsorption defect of pernicious anemia need B12 supplementation.
11. Vitamin C—Supplementation is likely needed for individuals notconsuming optimum quantities of fruits and vegetables. Megavitamin dosesof vitamin C for most individuals are not harmful.
12. Essential fatty acids—Under typical conditions, linoleic acid(18:2ω6) deficiency would not be expected for humans. The need for supplementing the linolenic acid family (e.g., 22:6ω3) has been suggested forprevention of cardiovascular disorders, hypertension, inflammation, andsome types of cancer, as well as normal brain development and improvement of immune response.

PROVIDING VITAMIN SUPPLEMENTS
The physical and chemical forms of various vitamin products used
to fortify feeds are usually different from the forms found in feedstuffs.
Modification of these naturally occurring vitamin forms is required to
improve their stability, compatibility, dispersion, and handling characteristics for feed fortification.
Various chemical and physical vitamin forms available for supplementation are presented in Table 19.1 (Adams, 1978; Hoffmann-La
Roche, 1991). To obtain sufficient stability, compatibility with otherfeed components, and the properties required for application, vitaminproducers try to devise vitamin forms according to the following methods (Schneider and Hoppe, 1986):
735

■ Table 19.1

Product Forms of Vitamins and Application Uses

736

Product Form
VitaminA

ChemicalPhysical

Alcohol (retinol)
Acetate and/or palmitate

Crystalline or liquid
Beadlets
Spray- or drum-dried powders
Liquid concentrates

Propionate
D(D2 and D3)

E

Ergocalciferol (D2), orcholecalciferol (D3)d- or dl-α-tocopheryl acetate
Mixed tocopherols
d-α-tocopheryl succinate

K

Menadione (K3),
MSB (menadione
sodium bisulfite),
MSBC (menadione sodium
bisulfite complex), or
MPB (menadione dimethyl
pyrimidinol bisulfite)
Phytomenadione (K1)

Oil dilutions
Oil absorbates
Liquid concentrates
Beadlets (with vitamin A)
Spray- or drum-dried powders
Liquid concentrates (with vitamin A)

Application Uses
Liquid and dry oral pharmaceuticals
Dry feeds; dry oral pharmaceuticals
Dry feeds; dry oral pharmaceuticals;
water-dispersible vitamin products
Liquid feed supplements; oral andparenteral pharmaceuticals
Nonpelleted feeds
Nonpelleted feeds
Oral and parenteral pharmaceuticals;
liquid feed supplements

Oil dilutions
Oil absorbates

Dry feeds; dry oral pharmaceuticals
Dry feeds; dry oral pharmaceuticals
Liquid feed supplements; oral andparenteral pharmaceuticals
Nonpelleted feeds
Nonpelleted feeds

Absorbate powder or oil
Spray-dried powders
Oils
PowderFeeds, foods, and pharmaceuticals
Water-dispersible vitamin products
Foods and pharmaceuticals
Pharmaceuticals

Dry dilutions
Water-dispersible powders

Dry feeds or pharmaceuticals
Water-dispersible vitamin products

LiquidParenteral pharmaceuticals

■ Table 19.1

Continued
Product Form

VitaminChemicalPhysicalApplication Uses

Thiamin (B1)

Thiamin mononitrate
Thiamin hydrochloride

Crystalline; dry dilutions
Dry dilutions
Crystalline

Feeds, foods, and pharmaceuticals
Feeds
Parenteral and oral pharmaceuticals; feeds

Riboflavin (B2)

Riboflavin:chemically synthesized
crystalline product;
fermentation product
Riboflavin-5′-phosphate
pharmaceuticals

High-potency powder;
Spray-dried powders

Feeds, foods, and dry oral pharmaceuticals

Water-soluble powder

Parenteral and liquid oral

Niacin (B3)

Niacin (niacinamide;
nicotinic acid)

Crystalline
Dry dilutions

Feeds and pharmaceuticals
Feeds

Vitamin B6

Pyridoxinehydrochloride

Dry dilution
Crystalline

Feeds
Oral and parenteral pharmaceuticals

Pantothenic acid

Calcium d- or dl-pantothenate
Calcium dl-pantothenatecalcium chloride complex
d-Panthenol

Powders
PowderFeeds, foods, and pharmaceuticals
Feeds

LiquidParenteral and liquid oral pharmaceuticals;
cosmetics

737

Biotind-Biotin

Crystalline
Dry dilutions

Feeds, foods, and pharmaceuticals
Feeds

Folacin

Folacin

Crystalline
Dry dilutions

Feeds, foods, and pharmaceuticals
Feeds

■ Table 19.1

Continued

738

Product Form
VitaminChemicalVitamin B12 (cyanocobalamin):(a) Crystalline productfrom fermentation(b) Chemically synthesized crystalline
product

B12

C

Ascorbic acid
Sodium ascorbate

Choline

Choline chloride
Choline bitartrate

PhysicalApplication Uses

Dry dilutions

Feeds

Water-soluble dilutions oral andparenteral pharmaceuticals

Water-dispersible vitamin products;

Dry dilutions
Coated products
Crystalline
PowderFeeds
Feeds and foods
Parenteral pharmaceuticals; feeds
Antioxidant-preservative for foods

70% liquid
25–60% dry powders
Crystalline
Water-soluble powders

Feeds
Feeds
Pharmaceuticals
Water-dispersible vitamin products;
pharmaceuticals

Sources: Modified from Adams (1978) and Hoffmann-La Roche (1991).

V I TA M I N S U P P L E M E N TAT I O N■ Synthesis of stable derivatives■ Addition of stabilizing agents■ Coating■ Absorption of liquid vitamins on suitable carriers■ Transformation of fat-soluble or poorly soluble vitamins into water-soluble or dispersible forms■ Transformation of water-soluble vitamins or derivatives into poorly

water-soluble or fat-soluble forms■ Standardization of content■ Providing high bioavailability
In view of the nutritional importance of vitamins A, D, and E, many
commercial vitamin producers have succeeded in enhancing stability ofthese vitamins in two ways: (1) by mechanical means, enveloping minutedroplets of the vitamin or vitamins in a stable fat or gelatin, forming
small beads, thus preventing most of the vitamin from coming into contact with oxygen until it is digested in the animal intestinal tract, and (2)through use of effective antioxidants that markedly prolong the induction period that precedes active vitamin oxidation. The stabilized
beadlet containing an effective antioxidant will protect these vitamins
for storage periods up to 4 to 8 weeks without much loss of vitamin potency. New technology has further improved vitamin A and D3 stability
by a cross-linking process, such as the reaction between the gelatin andthe sugar, that makes the beadlet insoluble in water, giving it a more resistant coating that can sustain higher pressure, friction, temperature,and humidity (Coelho, 1996). Instability of vitamin D in peroxidizing
diets is sometimes overlooked. Studies have shown that very high dietarylevels of vitamin D are completely destroyed in diets containing high levels of peroxidizing polyunsaturated fatty acids, and chicks suffer severe
rickets by 3 weeks of age (Scott, 1973). Rickets was prevented by a normal level of vitamin D when the diet was supplemented with 57 mg of
ethoxyquin per kilogram of diet.
Stress factors affecting vitamin stability during manufacture andstorage of a custom premix may include heat, oxygen, moisture, oxidation, reduction, trace minerals, and pH. Stability of some vitamins is notaffected (Frye, 1978). Riboflavin, niacin, d-biotin, d-pantothenic acid,vitamin B12, and choline generally have excellent stability in custom premixes. Other vitamins, such as vitamin A acetate, vitamin D3, and vitamin E acetate, are also available in stabilized forms in custom premixes.
Vitamin K, unstabilized vitamin A, unstabilized vitamin D, thiamin, fo-739

CHAPTER NINETEEN

lacin, vitamin C, and vitamin B6 have poor stability in custom premixes
under various stress conditions (Coelho, 1996).
Most vitamins are supplemented in dry form; however, liquid vitamin supplements are useful for certain feeding operations. Their appeal
is broad, the advantages being lower feed cost per kilogram of gain, convenience in handling, ready adaptation in formulation, easy control, reduced time and labor, control of feed dust, and good palatability (Perry,1968). Liquid supplements are not without disadvantages—such as
physical stability and corrosion potential—most of which are minimal
with properly formulated supplements and equipment and adequate
feeder use knowledge (Bauernfeind, 1969). Ready availability of themany water-soluble or water-dispersible nutrients, such as energy
sources (molasses), nonprotein nitrogen (urea), trace minerals, and vitamins (A, D, and E), have permitted standard and custom formulation forruminants in feedlots and on pasture. Special economical liquid emulsions of vitamins A, D, and E, which remain acceptably uniform in physical distribution and with adequate stability patterns, are marketed forthis purpose.
When control of animal feed intake is not complete but there isgood control over water consumption, special vitamin A, D, and E dry
beadlet or liquid products are available to add to drinking water. They
disperse in cold water into a fine emulsion with desirable physical, taste,and chemical stability characteristics for short-time use. This vitaminform is useful in drinking water for poultry and other monogastric animals. Water-soluble vitamins and drugs, in properly formulated products, can also be added to water.
Because of the high variability and unknown bioavailability of vitamins in feeds, feed manufacturers have come to rely to a large extent oncommercially synthesized vitamins. In practice, feed manufacturers usually ignore to a certain degree the contribution of many vitamins in feedstuffs and provide complete vitamin supplementation. Bauernfeind(1969) summarized the advantages of synthetic vitamins in animal feeds:
1. Biological and physical characteristics are known; potency is uniform, stability is adequate with few exceptions, and supply is usually unlimited.
2. Weight added per ton to the animal diet is small; hence, adjustments
upward or downward or in different ratios can be made without upsetting
the remainder of the diet.
3. Ready-made combinations can be formulated in premix form for740

V I TA M I N S U P P L E M E N TAT I O N

quick addition to specific diets for a given species of animal for a defined
production objective with assurance of known diet values.
4. Cost is economical, hence vitamin restrictions of individual natural
feed ingredients can be removed from the computer program, thus increasing flexibility of programming for least-cost energy and protein needs andshortening computer operation time.
5. Assay costs of determining variability of natural vitamin content offeedstuffs can be decreased by supplementing with the chemically produced
nutrients.
6. Assay costs of determining the vitamin content of the final mixed dietcan be decreased since assay of one or two components of the premix will
give confidence of the mixing adequacy of all of the premix vitamins in the
final diet.
7. Use of chemically prepared vitamins eliminates unknowns existing attimes in natural ingredients or seasonal nutrient variation or physiologicalnutrient availability.
8. Chemically prepared vitamins have versatility in form and applications—adaptability to dry feeds, liquid supplements, drinking water solutions, drenches, and parenteral and capsule forms.

FORMULATING VITAMIN PREMIXESThe custom vitamin premix is generally considered to be a concentrated mixture of vitamins added in the manufacture of complete feeds
or feed supplements, and it is customized to meet vitamin specificationsof the feed manufacturer.
Use of custom vitamin premixes in feed manufacturing facilitates
uniform distribution of vitamins in complete feeds and ensures thatspecifications for vitamin fortification of feeds are met.
Vitamin premixes are mixtures of specific vitamins required in combination with some type of carrier material that is added to feeds—usually at a rate of 1 to 5 lb per ton of complete feed—at the time of mixing to ensure uniform distribution of these ingredients in mixed feeds.
Commonly used carriers include soybean meal, ground grain, corn
gluten meal, wheat middlings, and several other mill feeds.
Zhuge and Klopfenstein (1986) compared vitamin premix carriers
and found that destruction of vitamin A was greatest when groundsorghum was used as a diluent and least when the diluent was rice hulls
or ground corn. Other carriers, such as wheat middlings, corn gluten
meal, and soybean meal, are less suitable than rice hulls. Most vitamin741

CHAPTER NINETEEN

premixes are delivered in 50- to 55-lb plastic-lined paper bags. This
combination of materials limits recycling opportunities. A viable alternative is the use of 1-ton capacity bags or bulk bags.
Custom vitamin premixes should possess certain physical and chemical properties to ensure optimum quality and dispersion in finishedfeeds. The most important physicochemical properties for vitamin premixes are flowability, hydroscopicity, lumping, compression, andchargeability (BASF, 1997b).
Desirable properties suggested by Aiello (1978) and Hoffmann-La
Roche (1989, 1994) are as follows:
1. Proper particle size distribution. A good standard is 100% through a#20 U.S. standard sieve.
2. Free of contaminants.
3. Minimal moisture content. Excessive moisture (> 12%) can promote
destruction of certain vitamins as well as bacterial and mold growth andcauses handling problems due to caking and lumping.
4. Desirable mixing properties. Free-flowing, noncaking, nondusting,nonelectrostatic, nonhygroscopic, and nonsegregating.
5. Favorable pH. Most vitamins have their own specific pH level for optimum stability; however, a pH level of 5.5 is generally considered ideal for
multivitamin premixes.
6. Proper bulk density. This contributes to proper premix particle sizefor uniform dispersion, free-flowing properties, and ease of premix handlingand packaging.
7. Storage potential. Premixes should also be formulated to ensure optimum vitamin levels and stability under practical storage conditions. This
includes addition of antioxidants to improve stability of vitamins such as A,
D, and E.
8. Not mixing with antagonists. To prevent vitamin destruction, it isbest not to include choline and trace minerals in vitamin premixes.

■ REFERENCES
Adams, C.R. (1973). In Effect of Processing on the Nutritional Value of Feeds,
p. 142. National Academy of Sciences, Washington, D.C.
Adams, C.R. (1978). Proc. Roche Vitam. Nutr. Update Meet., p. 69. Arkansas
Nutrition Conference, Hot Springs.
Aiello, R. (1978). Proc. Roche Vitam. Nutr. Update Meet., p. 54. Arkansas Nutrition Conference, Hot Springs.
BASF. (1991). Vitamins—One of the Most Important Discoveries of the Century,742

V I TA M I N S U P P L E M E N TAT I O N

BASF Corporation, Parsippany, New Jersey.
BASF. (1997a). Antioxidant Status and the Susceptibility of Horses to ExerciseInduced Muscle Damage, KC9618, BASF Corporation, Mount Olive, New
Jersey.
BASF. (1997b). Physico-Chemical Properties of Vitamins, Trace Minerals andCarriers, KC9617, BASF Corporation, Mount Olive, New Jersey.
Bauernfeind, J.C. (1969). World Rev. Anim. Prod. 5, 20.
Christian, L.D. (1983). In Proc. AFIA Nutrition Council, p. 22. Ames, Iowa.
Coelho, M. (1994). Feedstuffs 66(25), 13.
Coelho, M.B. (1995). In Proc. Maryland Nutrition Conference for Feed Manufacturers, p. 46. College Park, Maryland.
Coelho, M.B. (1996). Impact of Vitamin Sources and Feed Processing on Vitamin Stability, p. 38. BASF Technical Symposium, Seattle, Washington.
Comben, N., Clark, R.J., and Sutherland, D.J.B. (1984). Vet. Rec. 115, 642.
Cunha, T.J. (1977). Swine Feeding and Nutrition. Academic Press, New York.
Cunha, T.J. (1980a). J. Anim. Sci. 51, 1429.
Cunha, T.J. (1980b). Horse Feeding and Nutrition. Academic Press, New York.
Cunha, T.J. (1982). Niacin in Animal Feeding and Nutrition. National Feed Ingredients Association (NFIA), Fairlawn, New Jersey.
Cunha, T.J. (1984a). Feed Manage. 35, 14
Cunha, T.J. (1984b). Squibb Int. Swine Update 3, 1.
Cunha, T.J. (1985). Feedstuffs 57, 37.
Cunha, T.J. (1991). Horse Feeding and Nutrition, 2nd Ed. Academic Press, NewYork.
Dove, C.R., and Ewan, R.C. (1986). Swine Research Report, AS-580-J. Iowa
State University, Ames.
Dritz, S.S., Tokach, M.D., Nelssen, J.L., Goodband, R.D., and Lynh, G. (1995).
In Proc. Kansas State University Swine Day, p. 7. Lawrence, Kansas.
Dubeski, P.L., d’Offay, J.M., Owens, F.N., and Gill, D.R. (1996a). J. Anim. Sci.
74, 1367.
Dubeski, P.L., Owens, F.N., Song, W.O., Coburn, S.P., and Mahuren, J.D.
(1996b). J. Anim. Sci. 74, 1358.
Dudley-Cash, W.A. (1994). Feedstuffs 66(6), 12.
Ellis, R.P., and Vorhies, M.W. (1976). J. Am. Vet. Med. Assoc. 168, 231.
Friesecke, H. (1975). In Pantothenic Acid, No. 1533. Hoffmann-La Roche,
Basel, Switzerland.
Frye, T.M. (1978). Proc. Roche Vitam. Nutr. Update Meet., p. 61. Arkansas Nutrition Conference, Hot Springs.
Gadient, M. (1986). Proc. Nutr. Conf. Feed Manuf., p. 73. College Park, Maryland.
Gwyther, M.J., Tillman, P.B., Frye, T.M., and Lentz, E.L. (1992). In 13th Annual
Meeting Southern Poultry Science. Atlanta, Georgia.
Hill, C.H. (1961). Poult. Sci. 40, 762.
Hoffmann-La Roche. (1989). Vitamin Nutrition for Poultry, RCD 7817/689.
Hoffmann-La Roche, Inc., Nutley, New Jersey.
Hoffmann-La Roche. (1991). Vitamin Nutrition for Swine, RCD 8260/191.
Hoffmann-La Roche, Inc., Nutley, New Jersey.
Hoffmann-La Roche. (1994). Vitamin Nutrition for Ruminants, RCD 8775/
743

CHAPTER NINETEEN

894. Hoffmann-La Roche, Inc., Nutley, New Jersey.
Kennedy, D.G., Rice, D.A., Bruce, D.W., Goodall, E.A., and McIlroy, S.G.
(1992). Br. Poult. Sci. 33, 1015.
Kurnick, A.A., Hanold, F.J., and Stangeland, V.A. (1972). Proc. Georgia Nutr.
Conf., p. 107. Atlanta Georgia.
Livingston, A.L., Nelson, J.W., and Kohler, G.O. (1968). J. Agric. Food Chem.
16, 492.
Mangelson, F.L., Draper, C.I., Greenwood, D.A., and Crandall, B.H. (1949).
Poult. Sci. 28, 603.
Marks, J. (1975). A Guide to the Vitamins. Their Role in Health and Disease, p.
73. Medical and Technical Publ., Lancaster, England.
McDowell, L.R. (2000). Asian–Aust. J. Anim. Sci. 113, 115.
McGinnis, C.H. (1986). Bioavailability of Nutrients to Feed Ingredients, p. 1.
National Feed Ingredient Association (NFIA), Des Moines, Iowa.
McMillen, W.N., Luecke, R.W., and Thorpe, F. Jr. (1949). J. Anim. Sci. 8, 518.
McNaughton, J. (1990). Feedstuffs 62(36), 13.
Newton, G.L., and Burtle, G.J. (1992). Current Concepts in Carnitine Research,p. 59. CRC Press, Boca Raton, Florida.
Nockels, C.F. (1988). In Proc. 1988 Georgia Nutr. Conf. for the Feed Industry.
Atlanta, Georgia.
NRC. (1973). Effect of Processing on the Nutritional Value of Feeds. National
Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1982). United States-Canadian Tables of Feed Composition, 3rd Ed. National Academy of Sciences-National Research Council, Washington, D.C.
NRC. (1998). Nutrient Requirements of Swine, 10th Ed., National Academy of
Sciences-National Research Council, Washington, D.C.
Oldfield, J.E. (1987). J. Nutr. 117, 1322.
Perry, S.C. (1978). Proc. Roche Vitam. Nutr. Update Meet., p. 29. Arkansas Nutrition Conference, Hot Springs.
Perry, T.W. (1968). Feedstuffs 40(25), 48.
Purser, K. (1981). Anim. Nutr. Health April, 38.
RDA. (1989). Recommended Dietary Allowances, 9th Ed. National Academy of
Sciences-National Research Council, Washington, D.C.
Roche. (1979). Optimum Vitamin Nutrition. Hoffmann-La Roche, Nutley, New
Jersey.
Schneider, J., and Hoppe, P.P. (1986). Bioavailability of Nutrients in Feed Ingredients, p. 1. National Feed Ingredients Association (NFIA), Des Moines,
Iowa.
Scott, M.L. (1973). In Effect of Processing on the Nutritional Value of Feeds, p.
119. National Academy of Sciences, Washington, D.C.
Scott, M.L., Nesheim, M.C., and Young, R.J. (1982). Nutrition of the Chicken,
p. 119. Scott, Ithaca, New York.
Skinner, J.T., Izat, A.L., and Waldroup, P.W. (1991). Poult. Sci. 70(Suppl. 1),
112.
Stahly, T.S. (1994). In Carolina Swine Nutrition Conference. Raleigh, North
Carolina.
Stamberg, O.E., and Peterson, C.F. (1946). Poult. Sci. 25, 394.

744

V I TA M I N S U P P L E M E N TAT I O N

Teeter, R.G., and Deyhim, F. (1996). In El Foro 96 de Alimentos Balanceados,
p. 71. Watt Publishing Co., Mt. Morris, Illinois.
Tengerdy, R.P., and Nockels, C.F. (1975). Poult. Sci. 54, 1292.
Tiege, J., Saxegaard, F., and Froslie, A. (1978). Acta Vet. Scand. 19, 133.
Tsai, K.S., Health, H., Kumar, R., and Riggs, B.L. (1984). Clin. Res. 32, 411A.
Weiss, W.P. (1998). J. Dairy Sci. 81, 2493.
Young, L.G., Lun, A., Pos, J., Forshaw, R.P., and Edmeades, D. (1975). J. Anim.
Sci. 40, 495.
Zhuge, G., and Klopfenstein, C.F. (1986). Poult. Sci. 65, 987.
Zinn, R.A., Owens, F.N., Stuart, R.L., Dunbar, J.R., and Norman, B.B. (1987).
J. Anim. Sci. 65, 267.

745

appendix

V I TA M I N R E Q U I R E M E N T S
OF LIVESTOCKAND HUMANS;FEED COMPOSITION;METRIC CONVERSIONS■ Table A1a

Vitamin Requirements for Ruminants and Horses (units per kg of diet,AnimalA

D

E

K

ThiaminBeef cattlea
Feedlot
Pregnant heifers and cows
Lactating cows and bulls
Growing

IU
2200
2800
3900
—

IU
275
275
275
—

IU
—
—
—
15–60

MSb
MS
MS
—

MS
MS
MS
MS

Dairy Cattlec
Growing
Lactating cows and bulls
Calf milk replacer

IU
2200
3200
3800

IU
300
1000
600

IU
25
15
40

—
MS
MS
—

—
MS
MS
6.5 ppm

SheepdReplacement ewes, 60 kg
Pregnant ewes, 70 kg
Lactating ewes, 70 kg
Replacement ram, 70 kg
Ewes, maintenance, 70 kg

IU
1567
3306
2380
1979
2742

IU/100 kg BW
555
555
555
555
555

15
15
15
15
15

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

Goate
All classes

IU
5000

IU
1400

IU
100

—
MS

—
MS

HorsefGrowing
MaintenanceWorking
Pregnant
Lactating

IU
2000
2000
2000
3000
3000

IU
800
300
300
600
600

IU
80
50
80
80
80

—
MS
MS
MS
MS
MS

mg
3
3
5
3
3

aNRC (1996). Nutrient Requirements of Beef Cattle. National Academy of Sci-

ences-National Research Council, Washington, D.C.
bMS = microbial synthesis.
cNRC (1989). Nutrient Requirements of Dairy Cattle. National Academy of Sci-

ences-National Research Council, Washington, D.C.

748

100% dry basis unless otherwise specified)Riboflavin Niacin

B6

Pantothenic
Acid

BiotinFolacin

B12

Choline

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

MS
MS
MS
MS

—
MS
MS
6.5 ppm—
MSMS
2.6 ppm—
MSMS
6.5 ppm—
MSMS
13 ppm—MS
MS
0.1 ppm—
MSMS
0.5 ppm—
MSMS
0.07 ppm

—
MS
MS
0.26%

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

MS
MS
MS
MS
MS

—
MS

—
MS

—
MS

—
MS

—
MS

—
MS

—
MS

—
MS

mg
2
2
2
2
2

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

—
MS
MS
MS
MS
MS

dNRC (1985a). Nutrient Requirements of Sheep. National Academy of Sciences-

National Research Council, Washington, D.C.
eMorand-Fehr, P. (1981). In Goat Production. (C. Gall, ed.) p.193, Academic Press,

New York.
fNRC (1989). Nutrient Requirements of Horses. National Academy of Sciences-Na-

tional Research Council, Washington, D.C.

749

■ Table A1b

Vitamin Requirements for Monogastric Animals (units per kg of diet,AnimalSwineaGrowing-finishing
3–5 kg
5–10 kg
10–20 kg
20–50 kg
50–120 kg
Breeding
Gilts and sows
Young and adult boars
Lactating gilts and sows
Poultryb
Leghorn chickens
Growing
0–6 weeks
6–12 weeks
12–18 weeks
18 weeks to first egg
Broilers
0–3 weeks
3–6 weeks
6–8 weeksTurkeys
0–4 weeks
4–8 weeks
8–12 weeks
12–16 weeks
16–20 weeks
20–24 weeks

Breeders holding
Laying hens
GeeseStarting 0–4 weeks
Growing 4+ weeks
Breeding
Duck
0–2 weeks
Growing 2–7 weeks
Breeding
Pheasant
0–4 weeks
4–8 weeks
9–17 weeks
Breeding

750

A

D

E

K

Thiamin

IU

IU

IU

mg

mg

2200
2200
1750
1300
1300

220
220
200
150
150

16
16
11
11
11

0.5
0.5
0.5
0.5
0.5

1.5
1.0
1.0
1.0
1.0

4000
4000
2000

200
200
200

44
44
44

0.5
0.5
0.5

1
1
1

1500
1500
1500
1500

200
200
200
300

10
5
5
5

0.5
0.5
0.5
0.5

1.0
1.0
0.8
0.8

1500
1500
1500

200
200
200

10
10
10

0.5
0.5
0.5

1.8
1.8
1.8

5000
5000
5000
5000
5000
5000

1100
1100
1100
1100
1100
1100

12
12
10
10
10
10

1.75
1.5
1.0
0.75
0.75
0.5

2
2
2
2
2
2

IU

IU

mg

mg

mg

5000
5000

1100
1100

10
25

0.5
1.0

2
2

1500
1500
4000

200
200
200

10
5
10

0.5
0.5
0.5

1.8
1.3
0.8

2500
2500
4000

400
400
900

10
10
10

0.5
0.5
0.5

—
—
—

5000
5000
5000
5000

1100
1100
1100
1100

12
10
10
25

1.75
1.5
0.75
1

2
2
2
2

100% dry basis unless otherwise specified)Riboflavin Niacin

B6

Pantothenic
Acid

BiotinFolacin

B12

Choline

mg

mg

mg

mg

mg

mg

μg

mg

4.0
3.5
3.0
2.5
2.0

20.0
15.0
12.5
10.0
7.0

2.0
1.5
1.5
1.0
1.0

12
10
9
8
7

0.08
0.05
0.05
0.05
0.05

0.30
0.30
0.30
0.30
0.30

20
17.5
15
10
5

600
500
400
300
300

3.75
3.75
3.75

10
10
10

1
1
1

12
12
12

0.2
0.2
0.2

1.30
1.30
1.30

15
15
15

1250
1250
1000

3.6
1.8
1.8
2.2

27
11
11
11

3
3
3
3

10
10
10
10

0.15
0.10
0.10
0.10

0.55
0.25
0.25
0.25

9
3
3
4

1300
900
500
500

3.6
3.6
3.0

35
30
25

3.5
3.5
3.0

10
10
10

0.15
0.15
0.12

0.55
0.55
0.50

10
10
7

1300
1000
750

4.0
3.6
3.0
3.0
2.5
2.5

60
60
50
50
40
40

4.5
4.5
3.5
3.5
3
3

10
9
9
9
9
9

0.25
0.2
0.125
0.125
0.100
0.100

1.0
1.0
0.8
0.8
0.7
0.7

3
3
3
3
3
3

1600
1400
1100
1100
950
800

mg

mg

mg

mg

mg

mg

µg

mg

2.5
4.0

40
40

3
4

9
16

0.100
0.20

0.7
1.0

3
3

800
1000

3.8
2.5
4

65
35
20

3
3
4.5

15
10
10

0.15
0.10
0.15

0.55
0.25
0.35

9
3
4

1500
1000
—

4
4
4

55
55
55

2.5
2.5
3.0

11
11
11

—
—
—

—
—
—

—
—
—

—
—
—

3.4
3.4
3.0
4.0

70
70
40
30

4.5
3.5
3
4

10
10
10
16

0.25
0.2
0.1
0.2

1
1
0.7
1

3
3
3
3

1430
1300
1000
1000

751

■ Table A1b

Continued

AnimalBobwhite quail
Starting
0–6 weeks
Growing 6+ weeks
Breeding
Japanese quail
Starting and growing
Breeding
Dogc
GrowthMaintenance
Catd
Growing
Minke
Weaning 13 weeks
Foxe
Growth

752

A

D

E

K

Thiamin

IU

IU

IU

mg

mg

5000
5000
5000

1100
1100
1100

12
10
25

1.75
0.75
1.0

2
2
2

1650
3300

750
900

12
25

1
1

2
2

IU/kg BWIU/kg BWIU/kg BW

202
75

22
8

1.2
0.5

—
—

54
20

IU

IU

IU

IU

mg

3333

500

30

100

5

IU

5930

µg/kg BW

mg

—

27

mg

—

IU

2440

1.3
µg

—

—

—

1

Riboflavin Niacin

B6

Pantothenic
Acid
BiotinFolacin

B12

Choline

mg

mg

mg

mg

mg

mg

μg

mg

3.8
3.0
4.0

30
30
20

3.5
3
4

12
9
15

0.25
0.10
0.20

1.0
0.75
1.0

3
3
3

1500
1500
1000

4
4

40
20

3
3

10
15

0.3
0.15

1
1

3
3

2000
1500

µg/kg BWµg/kg BWµg/kg BWµg/kg BWµg/kg BWµg/kg BWµg/kg BW

100
50

450
225

60
22

400
200

-------

8
4

1.0
0.5

50
25

mg

mg

mg

mg

µg

µg

µg

g

4

40

4

5

70

800

20

2.4

mg

mg

mg

mg

mg

mg

µg

1.6

20

1.6

8

0.12

0.5

32.6

----

mg

mg

µg

mg

3.7

9.6

1.8

7.4

----

----

µg

----

0.2

753

■ Table A.1b

Continued

AnimalFishf
CatfishCommon carp
Tilapia
Trout
SalmonLaboratory animalsh
Rat
Growth, gestation,or maintenance
Mouse
Guinea pig
Growing
Hamster, golden
Growing
Nonhuman primatei

D

E

K

IU

IU

IU

IU

mg

mg

500
—
—
2400
—

50
100
50
50
50

—
—
—
—
—

25–50
—
50
50
50

1.0
0.5
—
1
—

mg

ppm

1000–2000
4000
—
2500
2500

Rabbitg
Growth, female
and breeding male
Reproducing

Ascorbic
Acid Thiamin

A

IU

580

—

40

—

—

—

1160

—

40

2

—

—

mg

mg

mg

mg

0.7

0.025

18

1

—

4

0.72

0.025

22

1

—

5

IU

IU

IU

mg

mg

mg

21,960

1000

40

5

200

2

mg

—

IU

mg

mg

mg

2

—

27

4

—

20

mg

mg

—

100

—

IU

IU

IU

10,000 –15,000

2000

50

mg

Sources: All references are from the Nutrient Requirement Series, National Academy of Sciences-National Research Council, Washington, D.C.
aNRC (1998). (Diets are 90% dry matter.)bNRC (1994). (Diets are 90% dry matter.)
cNRC (1985). (Requirements on body weight basis.)dNRC (1986). (Diets are 100% dry matter.)754

Riboflavin Niacin

B6

Pantothenic
Acid
BiotinFolacin

B12

Choline

mg

mg

mg

mg

mg

mg

mg

mg

9
7
6
4
7

14
28
—
10
—

3
6
—
3
6

15
30
10
20
20

—
1
—
0.15
—

1.5
—
—
1.0
2

—
—
—
0.01
—

400
500
—
1000
800

mg

µg/g

—

180

39

—

—

—

—

1200

—

—

—

—

—

—

—

—

mg

mg

mg

mg

mg

mg

µg

mg

3-4

15

6

10

0.2

1

50

750

7

15

8

16

0.2

0.5

10

2000

mg

mg

mg

mg

mg

mg

3

10

3

20

0.2

4

—

1800

mg

mg

mg

mg

mg

mg

µg

mg

15

—

6

10

0.2

2

10

1800

mg

mg

mg

mg

mg

mg

mg

mg

5

50

2.5

15

0.1

0.2

—

—

mg

mg

eNRC (1982a). (Diets are 100% dry matter.)
fNRC (1993). (Purified diets, therefore values represent near 100% bioavailability.)gNRC (1977). (Diets are 90% dry matter.)hNRC (1995). (Diets are 90% dry matter.)
iNRC (1978). (Diets are 90% dry matter.)755

■ Table A.1c

Vitamin Requirements for Humans
A

D

E

K

C

(μgRE/day)a(IU/day)

Male

1000

400

(IU/day)(µg/day)(mg/day)

10

65–80

60

FemalePregnant
Lactating

800
800
1300

400
400
400

8
10
12

55–60
65
65

60
70
95

Infants
0-6 months
6-12 months

375
375

300
400

3
4

5
10

30
35

Children and adolescents 400–700

400

7

15–30

45

Source: RDA (1989) Recommended Dietary Allowances. National Academy of Sciences-National Research Council, Washington, D.C.
aRetinol equivalents. 1 RE = 1µg retinol or 6 µg β-carotene.

756

Thiamin Riboflavin

NiacinB6

Pantothenic
Acid(mg/day)(mg/day)(mg/day)(mg/day)

1.5

1.7

19

2.0

1.1
1.5
1.6

1.3
1.6
1.7

15
17
1-20

0.3
0.4

0.4
0.5

0.7-1.0

0.8-1.2

BiotinFolacinB12(mg/day)(µg/day)(µg/day)(µg/day)

4–7

30–100

200

2

1.6
2.2
2.1

4–7
4–7
4–7

30–10030–10030–100

180
400
280

2
2.2
2.6

5
6

0.3
0.6

2
3

10
15

25
35

0.3
0.5

13

1.0-1.4

3-5

20-30

50-1000.7-1.4757

■ Table A2 Composition of Important Feeds (Vitamins Expressed at 100% Dry Matter)
Fat-Soluble Vitamins
Feed Name, Description

Alfalfa hay, sun cured
Alfalfa leaves, sun cured
Bahia grass, fresh
Bakery waste, dehydrated
Barley, grain
Bean, navy, seeds
Beet, sugar pulp, dehydrated
Blood, meal
Bluegrass, Kentucky, fresh
Brewer’s grains, dehydrated
Buckwheat, common, grain
Buttermilk, dehydrated (cattle)
Carrot, root, fresh
Casein, dehydrated (cattle)
Cattle, liver, fresh
Cattle, spleen, fresh
Chicken broilers, whole, fresh
Chicken, hens, whole, fresh
Citrus, dried pulp
Clover, red hay, sun cured
Coconut, meats,
meal mechanically extracted
solvent extracted
Corn, dent yellow cobs, ground
Gluten meal
Grain
SilageCotton seeds, meal,
mechanically extracted,
41% protein
Crab meal
Fish, anchovy meal,
mechanically extracted
Fish, menhaden meal,
mechanically extracted
Fish, sardine meal,
mechanically extracted
Flax (linseed meal)
Livers, meal
Meat, with blood,
meal rendered (tankage)
Milk, dehydrated (cattle)

758

Carotene
Provitamin A

A

D

E

K

mg/kg

IU/g

IU/kg

mg/kg

mg/kg

58.0
88.0
183
5.0
2.0
—
—
—
248
—
—
—
678
—
—
—
—
—
—
20

—
—
—
7.0
—
—
—
—
—
—
—
2.4
—
—
439.1
3.0
30.0
—
—
—

1575
373
—
—
—
—
637
—
—
—
—
—
—
—
—
—
—
—
—
1914

113.0
——
45.0
25.0
1
—
—
—
29
—
7
60
—
25
56
—
310
—
—

21.6
——
—
0.2
—
—
—
—
—
—
—
—
—
—
—
—
—
—
—

—
—
1
18
3
43

—
—
—
—
—
—

—
—
—
—
—
439

—
—
—
34
25
—

—
—
——
0.2
—

—
—
—

—
—
—

—
—
—

35
—
5.0

—
—
—

—

—

—

13

—

—

—

—

—

—

—
—

—
—

—
—

9
—

—
—

—
—

—
12.3

—
353

—
—

—
—

■ Table A.2

Continued
Water-Soluble Vitamins

BiotinCholine

Folic
Acid

NiacinPantothenic
Acid
Riboflavin

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg μg/kg

0.20
0.28
—
0.07
0.17
0.12
—
0.09
—
0.68
—
0.31
0.07
0.05
3.51
0.16
—
0.46
—
0.11

—
1062.0
—
1005
1177
1499
902
854
—
1757
501
1891
—
229
5093
2036
—
6288
867
—

3.6
5.8
—
0.2
0.6
1.4
—
0.1
—
7.7
—
0.4
1.2
0.5
8.4
4.8
—
0.5
—
—

42
47
—
28
94
28
18
34
66
47
21
9
58
1
269
25
230
225
24
43

28.6
29.0
—
9.0
9.1
2.3
1.5
2.6
—
8.9
13.1
40.1
30.1
2.9
164.9
8.2
—
20.4
15.4
11.2

13.4
20.6
—
1.5
1.8
2.0
0.8
2.2
11.0
1.6
5.4
33.1
4.9
1.7
92.2
15.3
15.6
6.4
2.5
17.8

3.0
4.6
—
3.2
5.0
7.1
0.4
0.4
8.8
0.7
4.2
3.7
5.8
0.5
6.3
3.1
2.9
2.4
1.6
2.2

—
—
—
—
—
—
4.7
—
7.3
—
0.3
—
—
—
4.8
49
—
—
0.8
—
—
—
2.6
21
12.0
—
0.5
—
18.0 1523
1.3
247
—
—
4.6
278
—
—
—
—

1-00-078
1-00-146
2-00-464
4-00-466
4-00-549
5-00-623
4-00-669
5-00-380
2-00-7865-02-1414-00-994
5-01-160
4-01-145
5-01-162
5-01-1665-07-942
5-07-945
5-07-9504-01-237
1-01-415

—
—
—
0.20
0.08
—

1036
1189
—
391
567
—

1.5
0.3
—
0.3
0.3
—

26
28
8
55
28
47

6.8
6.9
4.2
11.2
6.6
—

3.4
3.7
1.1
1.8
1.4
—

0.8
0.7
1.0
0.2
3.8
—

—
4.8
—
8.8
5.3
—

—
—
—
—
—
—

5-01-572
5-01-573
1-28-2345-28-241
4-02-9353-02-912

1.19
0.07
0.21

2965
2179
4036

2.3
0.1
0.2

38
49
89

11.2
7.0
10.9

5.7
6.7
8.2

7.0
0.5
0.5

5.4
7.2
5.0

—
475
233

5-01-6175-01-663
5-01-985

0.20

3398

0.2

60

9.4

5.2

0.6

5.1

133

5-02-009

0.11

3518

—

81

11.8

5.8

0.3

—

256

5-02-015

0.36
0.02

1962
12,281

3.1
6.0

41
221

15.8
31.5

3.5
39.1

4.6
0.2

6.1
—

—
542

5-02-045
5-00-389

—
0.40

2391
—

1.7
—

40
9

2.8
23.8

2.4
20.6

0.4
3.9

—
4.9

147
—

5-00-3865-01-167

Thiamin

B6

B12International
Feed No.

759

■ Table A2Continued
Fat-Soluble Vitamins

Feed Name, Description

Milk, skim, dehydrated
Millet, grain
Molasses, sugarcane
Oat, grain
Pangola grass, freshPea, seeds
Peanut kernels, meal,
mechanically extracted
Potato tubers, dehydrated
Poultry, by-products,
meal, renderedfeathers, hydrolyzed
Rape, seeds,
mechanically extracted
Rice bran with germgrain, groundgroats, polished
Rye, grain
Ryegrass, hay, sun cured
Safflower, seeds, meal,
mechanically extracted
Sesame seeds, meal,
mechanically extracted
SorghumgrainsilageSoybean seeds
meal, solvent extracted
Sunflower, common seeds,
mechanically extracted
Swine livers, fresh
Timothy, fresh
hay, sun cured
Tomato pomace, dehydrated
Triticale grain
Turnip roots, fresh
Wheat
bran
grain
Whey, dehydrated (cattle)
Yeast, brewer’s, dehydrated
Yeast, torula, dehydrated

Carotene
Provitamin A

A

D

E

K

mg/kg

IU/g

IU/kg

mg/kg

mg/kg

—
—
—
—
62
1

—
—
—
—
—
—

446
—
—
—
—
—

10
—
7
15
—
3

—
—
—
—
—
—

—
—

—
—

—
—

3
—

—
—

—
—

—
—

—
—

2
—

—
—

—
—
—
—
—
120

—
—
—
—
—
—

—
—
—
—
—
—

20
66
11
4
17
211

—
—
—
—
—
—

—

—

—

1

—

—

—

—

—

—

1
15
1
—

—
—
—
—

26
662
—
—

10
—
37
3

0.2
—
—
—

—
—
179
28
—
—
—

—
361.7
—
—
—
—
—

—
—
—
2138
—
—
—

—
—
111
38
—
—
—

—
—
—
—
—
—
—

3
—
—
—
—

—
—
0.5
—
—

—
—
—
—
—

21
17
—
2
—

—
—
—
—
—

Source: United States-Canadian Tables of Feed Composition (NRC, 1982b); National Academyof Sciences-National Research Council, Washington, D.C.

760

Water-Soluble Vitamins
BiotinCholine

Folic
Acid

NiacinPantothenic
Acid
Riboflavin

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg

mg/kg μg/kg

0.35
—
0.92
0.31
—
0.22

1480
489
1012
1116
—
662

0.7
—
0.1
0.4
—
0.3

12
26
49
16
—
36

38.6
12.2
50.3
8.8
—
21

20.5
4.2
3.8
1.7
—
2.0

3.9
8.1
1.2
7.1
cc
5.2

4.5
—
5.7
2.8
—
1.7

54
—
—
—
—
—

5-01-175
4-03-120
4-04-6964-03-309
2-03-493
5-03-600

0.35
0.11

2052
2879

0.7
0.7

186
37

49.7
22.0

8.8
1.1

6.6
—

8.0
15.5

—
—

5-03-649
4-07-850

0.09
0.05

6451
962

0.5
0.2

50
23

11.8
9.7

11.2
2.1

0.2
0.1

4.7
3.2

322
90

5-03-798
5-03-795

—
0.47
0.09
—
0.06
—

7103
1357
1076
1018
479
—

—
2.4
0.4
0.2
0.7
—

168
330
39
17
21
—

9.8
25.2
9.1
3.9
9.1
—

3.3
2.8
1.2
0.6
1.9
—

1.9
24.7
3.2
0.7
4.2
—

—
—
5.0
0.4
2.9
—

—
—
—
—
—
—

5-03-870
4-03-928
4-03-938
4-03-942
4-04-047
1-04-077

1.54

1287

0.5

—

—

—

—

—

—

5-04-109

—

1655

—

20

6.4

3.6

3.0

13.4

—

5-04-109

0.42
—
0.37
0.36

737
—
2939
2915

0.2
—
3.9
0.7

43
—
24
31

12.5
—
17.3
18.2

1.4
—
3.1
3.2

4.7
—
10.6
6.2

5.0
—
—
6.7

—
—
—
—

4-04-383
3-04-323
5-04-610
5-04-604

—
2.49
—
0.07
—
—
—

4214
—
—
811
—
514
—

—
6.9
—
2.3
—
—
2.8

293
544
—
29
—
—
72

33.3
77.9
—
7.9
—
—
19.0

3.4
90.3
11.5
10.1
6.7
0.5
6.5

3.4
7.7
2.9
1.7
12.3
—
7.1

12.4
10.0
—
—
—
—
—

—
935
—
—
—
—
—

5-09-340
5-04-792
2-04-912
2-04-912
1-04-893
4-20-362
4-05-067

0.32
0.11
0.38
1.08
1.47

1797
1085
1921
4227
3223

1.6
0.5
0.9
10.3
26

268
64
11
482
525

33.5
11.4
49.6
118.4
100.6

4.6
1.6
29.4
38.1
47.6

7.9
4.8
4.3
99.2
6.6

9.6
5.6
3.6
39.8
38.9

—
1.0
20
1
4

4-05-190
4-05-211
4-01-182
7-05-527
7-05-534

Thiamin

B6

B12International
Feed No.

761

■ Table A3

Metric Weights and Measures with Customary Equivalents

Capacity or Volume
1 cubic centimeter1 cubic meter1 milliliter
1 liter

1 kiloliter
Weight1 gram
1 kilogram
1 metric ton (1,000 kg)
Volume per unit area
1 liter/hectare
Weight per unit area
1 kilogram/square centimeter
1 kilogram/hectare
Acre per unit weight
1 square centimeter/kilogram
Temperature conversion formulas
Centigrade (Celsius)
Fahrenheit

=
=
=
=
=
=
=
=
=

0.061 cubic inch
35.315 cubic feet
1.308 cubic yards
0.0338 fluid ounce (U.S.)
33.81 fluid ounces (U.S.)
2.1134 pints (U.S.)
1.057 quarts (U.S.)
0.2642 gallon (U.S.)
264.18 gallons (U.S.)

=
=
=
=
=
=

0.03527 ounce (advp.)
35.274 ounces (advp.)
2.205 pounds (advp.)
0.984 ton (long)
1.102 tons (short)2204.6 pounds (advp.)=

0.107 gallon (U.S.)/acre==

14.22 pounds (avdp.)/square inch
0.892 pound (avdp.)/acre=

0.0703 square inch/pound (avdp.)==

5/9 (Fahrenheit −32)
9/5 Centigrade (Celsius) +32

Source: Modified from J. Anim. Sci. 25 (1966), 270.
Note: When conversions are made, the results should be rounded to a meaningful
number of digits relative to the accuracy of original measurements. Values for weights
and volumes are based on pure water at 4ϒC under 760 mm of atmospheric pressure.

762

INDEX
Absorption
biotin, 449
calcium, 104-106, 107-108,113,115
carnitine, 642
choline, 568-569, 570
folacin, 485-486
magnesium, 105
myo-inositol, 660
niacin, 351-352
pantothenic acid, 423
phosphorus, 104-106,108-109,
113-114
riboflavin, 315, 342
thiamin, 269-270, 274
vitamin A, 23-24
vitamin B,, 389
vitamin B,,, 529-531, 551
vitamin C, 602-603
vitamin D, 97
vitamin E, 161-162, 215
vitamin K, 232-234
Accutane, 73
Acetylcholine, 566, 567, 569, 571-572,
5 87-58 8
Acremonium coenophiatum, 286
Acrodynia, 385-386
ACTH (adrenocorticotropic hormone),
316
Acyl carrier protein (ACP), 423-426
Adenosylcobalamin, 532-533
ADP-ribose, 357
Aflatoxin, 70, 255
Air pollution, 112
a-keto acids, decarboxylation of,
271-272
Alcohol consumption
biotin deficiency, 470
folacin deficiency, 493, 509-510, 733

niacin deficiency, 374, 378
riboflavin requirements and, 324
thiamin deficiency, 277, 299, 304
vitamin A deficiency, 74
vitamin B, deficiency, 396, 407
vitamin B,, deficiency, 551
vitamin C deficiency, 632
vitamin D deficiency, 132
Aldose reductase, 669
Aldosterone and riboflavin, 316
Alfalfa
carotene content, 44,45
riboflavin content, 324, 341
vitamin D content, 115-116
vitamin destruction in field-cured, 716
vitamin E content, 176
vitamin K content, 243
Algae, 539, 540
Allowances, optimum vitamin, 723-727
Alzheimer’s diseasecarnitine and, 645, 654
choline and, 566, 588, 591
thiamin and, 299
Amino acid metabolism
folacin and, 487-490
vitamin B,, 391-393
Aminotransferases, 391
Amprolium, 267, 288, 303
Analytical procedures
biotin, 448
carnitine, 642
choline, 567-568
essential fatty acids (EFA),678
folacin, 484
niacin, 350-351
pantothenic acid, 422-423
riboflavin, 314-3 15thiamin, 269

763

INDEX
Analytical procedures (cont.)
vitamin A, 21-22
vitamin B,, 388-389
vitamin B,,, 528-529
vitamin C, 601-602
vitamin D, 95-97
vitamin E, 160-161
vitamin K, 231-232
Anemiacobalt and, 541,542-543
folacin and, 479-480, 489, 493, 495,
497-499,503-507,509,511,
5 14-5 15
intrinsic factor, 529
vitamin B, and, 400, 402, 403404, 405
vitamin B,, and, 523-524, 540,544-545,547452,557-558
vitamin C and, 611
vitamin E and, 202, 205
Anencephaly, 479, 508
Antagonists, vitamin, 722. See also specific vitamins
Antibiotics
choline and, 576
folacin requirements and, 493
myo-inositol deficiency, 663
pantothenic acid requirements and, 427
vitamin K deficiency and, 246, 248,
252,255
vitamin requirements, effects on,
722-723
Anticonvulsant drugs
folacin and, 493
vitamin D and, 139
Antioxidants, 733-734
0-carotene, 53, 76-77
carnitine, 644-645
carotenoids, 24, 75-78
control of lipid oxidation in meats,
2 12-2 13
disease resistance and, 167-168
flavonoids, 668-669
lycopene, 19, 78
metalloenzymes, 606role of, 204
in vitamin A supplements, 70, 71, 74
vitamin C, 600, 605-607, 626, 631-632
vitamin E, 156, 162, 164-166,212-213
Arachidonic acid, 675-680, 682-683,
691,693-694,704-705
Arthritis
vitamin C and, 621
vitamin E and, 203764

Ascorbic acid. See Vitamin C
Aspergillus ~ ~ Q V U 278
S,
Aspergillus nigec 665
Aspirin, 687
Atherocalcin, 238
Atherosclerosis
choline, 587, 591
vitamin C, 611
vitamin E, 216
Autoimmune disease, 683, 684
Avidin, 445446, 448, 474, 722

Bacillus thiaminolyticus, 288
Bats, vitamin B,, deficiency, 546
0-carotene, 19-21. See also Vitamin A
as anticarcinogen, 75-78
antioxidant, 53, 76-77
bactericidal effects, 76
cancer and, 38
conversion to vitamin A, 23,24, 41-42
function independent of vitamin A,
75-78
reproduction, effects on, 76-77
source of, natural, 43-45,47
supplementation, 74
Beef cattle. See Cattle
Beriberi, 12, 19, 29, 265-267, 280,
296-299
Betaine, 566, 569, 572, 574, 575, 577,
583,589
Bile and vitamin A, 27
Bioavailability, vitamin, 719-720
Biological assay. See Analytical procedures
Biotin, 445-478
analytical procedures, 448
antagonists, 447, 470, 722
bioavailability, 719
chemical structure and properties,
446-447
deficiency, 455-471
assessment of status, 470-471
cats, 466
dogs, 466
fish, 466
foxes and mink, 466467,468
horses, 4 6 4 4 6 5humans, 469-470
laboratory animals, 467-468
poultry, 460-464
primates, nonhuman, 4 6 8 4 6 9
rabbits, 469
ruminants, 456-457
swine, 457-460

INDEX
functions, 450-452
history, 445-446
introduction to, 445
metabolism, 448-450
requirements, 452-453
table of species, 453
sources, natural, 454-455
stability, 446, 453, 454-455, 474
supplementation, 471-474, 728-730,
732,737
toxicity, 474-475
Biotinidase, 449, 469
Bitot's spots, 63-64
Blacktongue, 347, 349, 368, 369, 371
Blindness, 15. See also Vision
Blood clotting, 235-236. See also Vitamin K
fatty acids and, 686, 702, 707
vitamin E and, 166-167
vitamin K deficiency, effects of, 244-253
Bonevitamin A and, 36, 52, 58, 78-79
vitamin C and, 618
vitamin D and, 106-107, 117-121,
123-132,140
vitamin K and, 238-239
Bone Gla protein (BGP), 238
Bracken fern (Pteridium aquilinum) poisoning, 2~92-293,302
Butylated hydroxytoluene (BHT), 65
B vitamins. See also specific vitamins
copropha ;y and, 7
monogasdc animals and, 6-7
ruminar'ts and, 6
Cabagin, 671
Calbindin, 105, 107, 108, 109-110
Calcitonin, 104
Calcitrol, 93. See also Vitamin D. Calcium
absorption, 113, 115
absorption by ruminants, 107-108
binding proteins, 105, 107, 108,109-110milk fever (parturient paresis), 121-122
mobilization by myo-inositol, 661
phytate and, 664-666
plants, calcinogenic, 142-145
rickets, 117-119
soft tissue calcification, 141-145
supplementation, 134, 140transplacental movement, 102transport, 105-106
vitamin B, deficiency, 394

vitamin D, 100, 102-123
bone effects, 106-107
intestinal effects, 104-106
kidney effects, 107
requirements, effect on, 113
toxicity, 141-146
vitamin K, 238-239, 241, 245
Cancercarotenoid protection against, 75, 77-78
choline deficiency and, 587, 591
fatty acids and, 686, 688, 702, 706, 707
folacin and, 479,483,493, 508-509,
510
linoleic acid and, 678
vitamin A deficiency, 38
vitamin B,, (laetrile) and, 670
vitamin C and, 607, 608, 611, 626
vitamin D and, 110, 133, 140
vitamin E and, 203,204-205
Canthaxanthin, 75
Carbohydrate metabolism
biotin, 4 5 0 4 5 1
niacin, 356
pantothenic acid, 424-426
thiamin, 271-272, 273,275-276
vitamin B,, 391, 392
vitamin B,,, 535
Carboxylases and biotin, 451,469
Carboxylation reactions and vitamin K,
236-238
Cardiovascular system
carnitine, 647, 650, 653
choline, 573, 587
dilated cardiomyopathy (DCM),
650-651
fatty acids, 678, 686-687, 688-689,
696,707flavonoids, 669
folacin, 479, 509, 515
thiamin, 280-282,289,294,296-298
vitamin C, 626
vitamin D
deficiency, 133
toxicity, 143
vitamin E, 181-183, 193, 197, 203,
204
Carnitine, 641-658
analytical procedures, 642
chemical structure and properties,
641-642
deficiency, 647-653
assessment of status, 653
dogs, 650-651

765

INDEX
Carnitine, deficiency (coat.)
fish, 651
horses, 650
humans, 652-653
laboratory animals, 651
poultry, 649-650
ruminants, 648
swine, 649
functions, 644-645
history, 641
introduction to, 641
metabolism, 642-644
requirements, 645
sources, natural, 646-647
supplementation, 653-655, 728-729
toxicity, 655
vitamin C and, 608
Carotene and carotenoids, 15, 16-17,
18-21. See also p-carotene; Vitamin A
absorption of, 23-24
as anticarcinogens, 75, 77-78
digestion, 22, 46
sources of, natural, 44-47
Carpal tunnel syndrome, 41 1
Carrots as carotene source, 45
Casal, Gaspar, 348
Catalase, 606
Cataracts
vitamin C and, 607, 626
vitamin E and, 203
Catsbiotindeficiency, 466
requirements, 453
cholinedeficiency, 580, 585
requirements, 577
essential fatty acids (EFA)
deficiency, 700
requirements, 689-691
folacin
deficiency, 503
requirements, 492
niacindeficiency, 369
requirements, 355, 358
pantothenic aciddeficiency, 436
requirements, 427
retinoid therapy, 73
riboflavindeficiency, 333-334
requirements, 322

766

thiamin
deficiency, 281, 293-294
requirements, 277
vitamin A
deficiency, 60-61
requirements, 40
storage, 27-28
supplementation, 72-73
toxicity, 79-80
vitamin B,
deficiency, 404
requirements, 395
vitamin B,, deficiency, 536, 546-547
vitamin C deficiency, 621
vitamin D
deficiency, 128
requirements, 111, 114
vitamin E
deficiency, 200-201
requirements, 171
vitamin K
deficiency, 250
requirements, 241
Cattlep-carotene supplementation, 76-77
biotindeficiency, 456
requirements, 453
supplementation, 456-457
calcium absorption, 107-108
carnitine deficiency, 648
carotene absorption, 23
cholinedeficiency, 579-581
metabolism, 570, 572
requirements, 576, 577
supplementation, 5’90-59 1
cobaltdeficiency, 525, 537
toxicosis, 558-559
essential fatty acids (EFA)
deficiency, 696-697
requirements, 690
supplementation, 704-705, 707-708
fo1acin
deficiency, 496-497
requirements, 492
supplementation, 496, 513
niacindeficiency, 363-365
requirements, 358
supplementation, 359, 364-365,
377-3 78

INDEX
pantothenic acid requirements, 427
production function and vitaminrequirements, 720
riboflavin, 322, 326
thiamin
deficiency, 280, 283-289
requirements, 277
supplementation, 303
vitamin A
deficiency, 33-34, 36, 38, 50-53
requirements, 40
storage, 28, 67-68
supplementation, 72, 73
vitamin B,
deficiency, 399
requirements, 395
vitamin B,,
deficiency, 541, 542, 543
requirements, 536
status determination, 554
supplementation, 555
vitamin C
deficiency, 613-615
supplementation, 628
vitamin D
deficiency, 119-123
requirements, 114
supplementation, 134-136
toxicity, 141
vitamin E
deficiency, 181-183,184,185-186
requirements, 171
supplementation, 209-213
vitamin K
requirements, 241
supplementation, 256
Cerebrospinal fluid pressure, vitamin A
deficiency and, 50, 51, 54
Cestrium diumum, 143
Chastek paralysis, 268
Cheilosis, 337, 374
Chickens
biotindeficiency, 4 6 0 4 6 4requirements, 453
carnitine deficiency, 649-650
cholinedeficiency, 283-584, 580
requirements, 577
toxicity, 592
essential fatty acids (EFA)
deficiency, 699
requirements, 690

folacin
deficiency, 499-501
requirements, 492
myo-inositol deficiency, 662-663
niacindeficiency, 367-368, 369
requirements, 358
supplementation, 376, 379
pantothenic aciddeficiency, 434-435
requirements, 426-427, 428
supplementation, 439
production function and vitaminrequirements, 720
restricted feeding and vitamin intake, 718
riboflavindeficiency, 330-332
requirements, 321, 322
thiamin
deficiency, 281, 289-292
requirements, 277
supplementation, 721-722
vitamin A
deficiency, 33, 36
requirements, 40
supplementation, 72
vitamin B,
deficiency, 401-402
requirements, 395
vitamin B,, requirements, 536
vitamin C, 617-619, 629
vitamin D
deficiency, 124-127
requirements, 114
supplementation, 138
toxicity, 141
vitamin E
deficiency, 194-196
requirements, 171
toxicity, 216
vitamin K
deficiency, 227-228,247-249
requirements, 240, 241
supplementation, 247
Cholecalciferol (vitamin D3), 91, 93,
94-98, 115-117, 146. See also
Vitamin D
Choline, 4, 565-596
analytical procedures, 567-568
bioavailability, 719
carnitine and, 643
chemical structure and properties,
566-567767

INDEX
Choline (cont.)
deficiency, 578-588
assessment of status, 588
cats, 580, 585
clinical signs, table of, 580
dogs, 580, 585
fish, 580, 585
horses, 585
humans, 586-588
laboratory animals, 580, 586
poultry, 580, 583-584
primates, nonhuman, 580, 586
rabbits, 580, 586
ruminants, 579-581
swine, 580,581-583
folacin and, 490, 501
functions, 571-573
introduction to, 565
metabolism, 568-571
requirements, 574-576
table of species, 577
stability, 591
supplementation, 588-591, 729-730,
732,738
toxicity, 592
Choline chloride and vitamin destruction,
716
Chromatography. See Analytical procedures
Chromium, 671
Chylomicronemia, 707
Citric acid cycle
coenzyme A and, 424,425
thiamin and, 271-272
vitamin B, and, 391
vitamin BIZ and, 534
Classification of vitamins, 4-5
Clostridium sporogenes, 288
Coagulation, blood. See also Vitamin K
overview of, 235-236
vitamin K deficiency, effects of, 244-253
Cobalamin, 526-528, 551. See also
Vitamin BIZ
Cobalophilin, 529
Cobalt, 6absorption, iron effects on, 531
deficiency
cattle, 525
ruminants, 541-543
sheep, 525
ruminant requirements, 537-538, 540
sources of, 540
status assessment, 554

768

supplementation, 556-557
toxicosis, 558-559
vitamin BIZ and, 523, 525
Coccidiosis, 41
vitamin C and, 617, 619
vitamin K deficiency, 722
Codecarboxylase. See Pyridoxal phosphate (PLP)
Cod liver oil, 11,43
Coenzyme A, 420-421,423-426
Coenzyme Q (ubiquinone), 318, 669-670
Coenzymes, 3
biotin, 450
cobalamin, 527, 532
folacin, 482,483, 487-488
niacin, 350-351, 355-357
riboflavin, 311, 317
thiamin, 301, 371
vitamin B,, 386, 391
Coffee, niacin content, 362
Collagenase inhibition by retinoic acid, 33
Collagen synthesis
vitamin C, 604-605
vitamin D, 107Computerized feed formulations, 720
Confinement rearing, 721
Conjugated linoleic acid (CLA), 678, 686,
693-694,706
Copperantioxidant, 606
coenzyme A and, 421
pyrroloquinoline quinone (PQQ) and,666
vitamin A stability and, 46, 71
vitamin C and, 607-608
vitamin destruction by, 715
vitamin E and, 177
Coprophagy, 7choline and, 576
folacin and, 490
thiamin and, 269, 274, 296
vitamin B, and, 395
vitamin BIZ and, 536, 548, 555
vitamin K and, 239-240, 243, 251
Cornbiotin content, 454-455
carotene content, 45-46, 74
choline content, 577
essential fatty acids (EFA)content, 704
niacin content, 348, 362, 365-366,
374-375
pantothenic acid content, 429,439
vitamin B, content, 409-410

INDEX
vitamin content, effects on, 714
vitamin E content, 176, 178, 214
Creatine and choline, 572
Crocodiles, thiamin deficiency, 296, 297
Curled-toe paralysis, 330, 331
Cyanocobalamin, 526-528, 555. See also
Vitamin B,,
Cystine and vitamin E, 170, 195
Cytochromes, 318, 669
Dairy cows. See Cattle
Dam, Henrik, 227-228
Davis, M., 12, 16
Decarboxylases
thiamin and, 271-272
vitamin B, and, 391
Dehydrogenases, 317-320, 356-357
Dementiacholine, 566, 588
folacin, 510
niacin, 374
Dermatitis/dermatosis
biotin deficiency, 456, 457, 462, 466,
467,469
essential fatty acids (EFA) deficiency,
696,697,701
niacin deficiency, 348-349, 367-368,
372-373
pantothenic acid deficiency, 420,
431-432,434-435,437
riboflavin deficiency, 337
vitamin B, deficiency, 385-386, 404,
405
vitamin C and, 614
vitamin E deficiency, 180, 199-200
Dexpanthenol, 441
Diabetes mellitus
niacin and, 360, 379
vitamin C and, 608, 626
Diarrhea
biotin deficiency, 466
folacin deficiency, 505, 507
niacin deficiency, 349, 363, 366-367,
369,371-372
pantothenic acid deficiency, 431,436
thiamin deficiency, 280
vitamin A deficiency, 30, 35,51, 6465, 75
vitamin E absorption, 215
Dicumarol, 230-231,244,250, 255
Digestion
biotin, 449
choline, 568
folacin, 485

pantothenic acid, 423
riboflavin, 315
thiamin, 269
vitamin A and, 22
vitamin B,, 389
vitamin B,,, 529
Dilated cardiomyopathy (DCM), 650-651
Diphenadione, 250
Discoid lupus erythematosus, 200
Disease resistance. See also Immune system; Infections
vitamin E, 163, 167-168
Distemper, canine, 621
DNA repair and niacin, 357
DNA synthesis
folacin, 506, 549
vitamin BIZ, 549, 552
vitamin E, 168
Docosahexaenoic acid, 677, 679-680,
687-688,695,702,704, 706
Dogs
biotin deficiency, 466
carnitine
deficiency, 650-651
supplementation, 653-654
cholinedeficiency, 580, 585
requirements, 577
essential fatty acids (EFA)
deficiency, 700
requirements, 690
folacin
deficiency, 503
requirements, 492
niacindeficiency, 349, 369, 371
requirements, 358
toxicity, 380
pantothenic aciddeficiency, 436
requirements, 427
retinoid therapy, 73
riboflavindeficiency, 333
requirements, 322
thiamin
deficiency, 281,293-294
requirements, 277
vitamin A
deficiency, 33, 61
requirements, 40
storage, 27-28
supplementation, 72-73

769

INDEX
Dogs, vitamin A (cont.)
toxicity, 79
vitamin B,
deficiency, 403-404
requirements, 395
vitamin B,,
deficiency, 547
requirements, 536
vitamin C
deficiency, 620-621
supplementation, 630
vitamin D
deficiency, 127-128
requirements, 111, 114
toxicity, 141
vitamin E
deficiency, 198-200
requirements, 171
vitamin K
deficiency, 250
requirements, 241
Drummond, 16
Ducks
niacindeficiency, 368
requirements, 355, 358
pantothenic acid requirements, 427
riboflavin requirements, 322
vitamin B, requirements, 395
vitamin D requirements, 114
vitamin E
deficiency, 196
requirements, 171
vitamin K requirements, 241
Edemavitamin B, deficiency, 400
vitamin E deficiency, 194, 201, 205
Eggs and biotin deficiency, 445446,466469
Eicosanoids, 682-684
Eicosapentaenoic acid, 679, 682-683,
688,695
Eicosatrienoic acid, 677, 680, 684, 704
Eijkman, Christiaan, 11, 266
Encephalocoele, 508
Encephalomalacia, 156, 194-195
Endochrondral ossification defects (EOD),
126
Enterococcus hirae, 484
Epithelium and vitamin A, 34-35. See
also Keratinization
Equine degenerative myeloencephalopathy, 198

770

Equine motor neuron disease, 198
Ergocalciferol (vitamin D2), 91, 93, 94,
98, 115-116, 117. See also Vitamin D
Erythrocyte glutathione reductase (EGR),340
Essential fatty acids (EFA), 675-712
analytical procedures, 678
chemical structure and properties,
676-678
deficiency, 695-704
assessment of status, 702-704
cats, 700
dogs, 700
fish, 700-701
foxes, 701
humans, 702
laboratory animals, 701
poultry, 699
primates, nonhuman, 701
rabbits, 702
ruminants, 696-697
signs and symptoms, 695-696
swine, 697-699
history of, 675-676
introduction to, 675
metabolism and functions, 679-689
cancer and, 686, 688
cardiovascular disorders, 686-687,
688-689
eicosanoids, 682-684
fluidity, 685
immune functions, 682-684
mammary epithelium, 685-686
nervous system and, 687-688
prostaglandins, 682-684
surfactant, 687
water balance, 685
requirements, 689-693
influencing factors, 691-692
table of species, 690
triene-tetraene ratio, 689
sources, natural, 693-695
supplementation, 704-708
toxicity, 708
Estrogens and vitamin K, 232-233
Ethoxyquin, 193, 739
Etretinate, 73
Euglena gracilis, 528
Excretion
biotin, 449, 450
carnitine, 643
choline, 569
fat-soluble versus water-soluble vitamins, 4

INDEX
folacin, 487
niacin, 352-353
pantothenic acid, 439
phosphorus, 109
riboflavin, 317
thiamin, 271
vitamin A, 27
vitamin B,, 390-391
vitamin B,,, 532
vitamin C, 603-604
vitamin D, 102
vitamin E, 164
vitamin K, 234-235
Exotic animals, vitamin E deficiency, 203
Extrusion and vitamin stability, 717
Exudative diathesis, 193-194, 195, 196,
201
Fat color and carotene, 23
Fat-soluble vitamins, 4-5, 6. See also
Vitamin A; Vitamin D; Vitamin E;
Vitamin K
Fatty acid-binding proteins (FABPs),679
Fatty acids. See also Essential fatty acids(EFA);Lipid metabolism
carnitine and, 644, 647
choline and, 571
cis versus trans, 676-678
flavoproteins and, 320
polyunsaturated (PUFAs), 170-172,
179-180, 186,200,214-215,688,
697-698
vitamin B, and, 391
Fescue toxicity, 286, 302
Fibrin, 235, 253
Fishbiotindeficiency, 466
requirements, 453
carnitine deficiency, 651
carotene absorption, 23
cholinedeficiency, 580, 585
requirements, 577
essential fatty acids (EFA)
requirements, 690, 691
as source of, 695
folacin
deficiency, 503-504
requirements, 492
myo-inositol deficiency, 663
niacindeficiency, 370-371

requirements, 358
pantothenic aciddeficiency, 436-437
requirements, 427
riboflavindeficiency, 334
requirements, 322
thiamin
deficiency, 294
requirements, 277
thiaminase, 268-269, 293-294
vitamin A
deficiency, 61
requirements, 40
storage, 28
vitamin B,
deficiency, 404
requirements, 395
vitamin BIZ
deficiency, 547
intestinal microfloral synthesis, 537
requirements, 536
vitamin C
requirements, 609-610
supplementation, 631
vitamin D
deficiency, 128-129
requirements, 114
vitamin E
deficiency, 201
requirements, 171
toxicity, 216
vitamin K
deficiency, 250
requirements, 241
5-fluorouracil, 481, 483
Flavin adenine dinucleotide (FAD),311-321.
See also Riboflavin (vitamin B2)
Flavin mononucleotide (FMN), 3 11-321.
See also Riboflavin (vitamin B2)
Flavonoids, 668-669
Flavoproteins, 3 17-321
Folacin, 479-521
analytical procedures, 484
antagonists, 483-484, 491, 498
chemical structure and properties,
481-484
choline and, 572, 574, 581, 587
deficiency, 495-512
assessment of status, 510-512
cats, 503
dogs, 503
fish, 503-504

773

INDEX
Folacin, deficiency (cont.)
foxes and mink, 504
horses, 502
humans, 505-510
laboratory animals, 504-505
poultry, 499-502
primates, nonhuman, 505
ruminants, 496-497
swine, 497-499
functions, 487-490
history, 480-481
introduction to, 479-480
metabolism
absorption, 485-486
digestion, 485
excretion, 487
storage, 486-487
transport, 486
PABA and, 667-668
requirements, 487-490
table of species, 492
sources, natural, 494-495
stability, 494-495, 513-514
supplementation, 512-5 15, 729,
732-733,735,737
toxicity, 515-516
vitamin BIZ and, 533-534, 535, 548,
549,552-553
vitamin C and, 608
Folate-binding proteins (FBPs),486
Food, irradiation of, 93, 717
Foods and feeds. See also Sources of vitamins
insufficient analysis of, 718-719
irradiation of food, 93, 717
vitamin content of
biotin, 455
carnitine, 647
choline, 578
essential fatty acids (EFA), 694
folacin, 495
niacin, 361
pantothenic acid, 429
riboflavin, 325
thiamin, 279
vitamin A, 47
vitamin B,, 397
vitamin BIZ, 539
vitamin C, 612
vitamin D, 116
vitamin E, 174, 176
vitamin K, 242
vitamin variability in, 718-719

772

Fowl typhoid (Salmonellagallinarum), 38,
619, 722
Foxes
biotin deficiency, 467, 468
essential fatty acids (EFA) deficiency,
701
folacin
deficiency, 504
requirements, 492
niacindeficiency, 371
requirements, 358
pantothenic aciddeficiency, 437
requirements, 427
riboflavindeficiency, 334, 335
requirements, 322
thiamin
deficiency, 268, 282, 295
requirements, 277
vitamin A
deficiency, 33, 62
requirements, 40
vitamin B,
deficiency, 404
requirements, 395
vitamin C and, 623
vitamin D deficiency, 129
vitamin E deficiency, 201
vitamin K deficiency, 251
Free radicals, 165,204. See also
Antioxidants
Friction and vitamin stability, 715
Friedreich’s ataxia disease, 587
Fruitscholine content, 578
vitamin C and, 597, 599, 611-612
vitamin losses from, 717
Funk, Casimir, 12,266, 347-348
Fusariurn moniliforme, 276, 278
Fusarium roseum, 138
Geesevitamin A requirements, 40
vitamin D requirements, 114
Gerbils, myo-inositol deficiency, 663
Gerovital, 671
Gingivitis and vitamin C deficiency, 624,
625
Glossitis and riboflavin deficiency, 337
Glucosamine, 35
Glucose tolerance factor, 671

INDEX
Glutathione peroxidase, 169, 173, 606,
1658
Glycerophosphocholine, 566-567, 568
Goatsbiotin requirements, 453
cholinemetabolism, 570
requirements, 577
folacin requirements, 492
niacin requirements, 358
pantothenic acid requirements, 427
riboflavin requirements, 322
thiamin requirements, 277
vitamin A
deficiency, 54-55
requirements, 40
storage, 67
vitamin B, requirements, 395
vitamin B,, requirements, 536
vitamin D
deficiency, 121, 123
requirements, 114
vitamin E
deficiency, 185requirements, 171
vitamin K requirements, 241
Gossypol, 186-188
Grains. See also Foods and feeds
biotin content, 454-455, 471
choline content, 577
essential fatty acids (EFA)content, 704
folacin content, 514
niacin content, 361-362
pantothenic acid content, 429430,439
phytate content, 664
thiamin content, 278, 279
vitamin E content, 175, 176, 178
Growth hormone, choline requirements
and, 575
Guinea pigs. See also Laboratory animals
biotin deficiency, 468
carnitine deficiency, 643, 651
choline deficiency, 586
essential fatty acids (EFA)
deficiency, 701
requirements, 690
myo-inositol deficiency, 662
niacin deficiency, 372
pantothenic acid deficiency, 438
riboflavin deficiency, 334-335
thiamin deficiency, 295
vitamin A deficiency, 62
vitamin C

deficiency, 623
toxicity, 634
vitamin D
deficiency, 129
requirements, 114
vitamin E deficiency, 202
vitamin K
deficiency, 251
requirements, 241
Hamsters. See also Laboratory animals
biotindeficiency, 468
requirements, 453
choline deficiency, 586
folacin requirements, 492
myo-inositol deficiency, 662
thiamin deficiency, 295
vitamin A deficiency, 62
vitamin B, deficiency, 405
vitamin B,,
deficiency, 547-548
requirements, 536
vitamin D deficiency, 129
Hartnup’s disease, 360, 374, 378
Harvesting conditions and vitamin levels,
714
Hays. See also Alfalfa; Foods and feeds
carotene content, 44-45
thiamin content, 278
Heart disease. See Cardiovascular system
Heat stress, 53, 618-619, 629, 718. See
also Stress
Hemorrhages. See also Vitamin K, deficiency
flavonoids, 668-669
vitamin C deficiency, 614, 621, 623,
624,629
vitamin E deficiency, 192, 193, 194,
196,199
Hepatosis dietetica, 188
Herodotus, 91
High-density lipoprotein (HDL), transfatty acid depression of, 678
High-pressure liquid chromatography(HPLC). See Analytical procedures
Hippocrates, 11, 93
Histidine degradation and folacin, 489
History of vitamins, 7-13
biotin, 445-446
carnitine, 641
choline, 565-566
essential fatty acids (EFA), 675-676

773

INDEX
History of vitamins (coat.)folacin, 480-481
niacin, 347-349
pantothenic acid, 419-420
riboflavin, 3 11-3 12thiamin, 265-267
vitamin A, 16-18
vitamin B,, 385-386
vitamin BIZ, 523-525, 672
vitamin D, 91-94
vitamin E, 155-157
vitamin K, 227-229
vitamin-like substances, 659, 671-672
Holman, Dr. R.T., 676
Homocysteine, 489, 509, 511-512, 534,552-554,572
Hoof disorders and biotin, 456-459,
464-465
Hookworms, 37
Hopkins, Frederick, 12, 16
Horsesbiotindeficiency, 464-465
requirements, 453
supplementation, 464-465, 472
B vitamins, 7
carnitine deficiency, 650
carotene absorption, 23
choline requirements, 577
essential fatty acids (EFA)requirements,
690
folacin
deficiency, 502
requirements, 492
niacindeficiency, 368
requirements, 358
pantothenic aciddeficiency, 436
requirements, 427
riboflavindeficiency, 332
requirements, 322
thiaminabsorption, 269,274
deficiency, 282, 292-293
requirements, 277
supplementation, 304
vitamin A
deficiency, 33, 59-60
requirements, 40
vitamin B,
deficiency, 403

774

requirements, 395
vitamin BIZ
deficiency, 546
requirements, 536
vitamin C deficiency, 620
vitamin D
deficiency, 127
requirements, 114
toxicity, 142
vitamin E
deficiency, 197-1 98
requirements, 171
supplementation, 215
vitamin K
deficiency, 249
requirements, 241
toxicity, 258
vitamin supplementation, overview of,
730-732
Horsetail (Equisetum arvmse), 293
HumansP-carotene and, 38
biotindeficiency, 469-470
requirements, 453
status assessment, 471
supplementation, 473-474
carnitine
deficiency, 652-653
requirements, 645
supplementation, 654
toxicity, 655
carotene absorption, 23
cholinedeficiency, 586-588
requirements, 576, 577
supplementation, 591
toxicity, 592
essential fatty acids (EFA)
deficiency, 702, 703, 706-707
requirements, 690, 693
supplementation, 735
fatty acid transport, 679
folacinassessment of levels, 510-512
deficiency, 505-510
requirements, 491, 492-493
supplementation, 514-515, 735
toxicity, 5 15-5 16
myo-inositol, 663-664
niacindeficiency, 372-375
requirements, 358, 360

INDEX
supplementation, 378-379, 734-735
toxicity, 379-380
pantothenic aciddeficiency, 438-439
requirements, 427, 428-429
supplementation, 441
riboflavindeficiency, 337-339
requirements, 322, 323-324
supplementation, 340, 734
thiamin
deficiency, 281, 296-300
requirements, 276-277
supplementation, 304-305, 734
vitamin A
deficiency, 33, 63-65
requirements, 40
storage, 28
supplementation, 75, 734
toxicity, 81-82, 733
vitamin B,
deficiency, 405-408
requirements, 395, 396
supplementation, 41 1-412, 735
toxicity, 412-413
vitamin B,,
deficiency, 548-552
requirements, 536, 538
supplementation, 557, 735
vitamin C
deficiency, 624-626
excretion, 603-604
requirements, 610-61 1
storage, 603
supplementation, 632-633, 735
toxicity, 633-634
vitamin D
deficiency, 130-133
requirements, 114-115
supplementation, 139, 734
toxicity, 141, 733
vitamin E
deficiency, 203-206
plasma tocopherol levels, 207
requirements, 171, 174
supplementation, 215-216, 734
toxicity, 216
vitamin K
deficiency, 251-253, 733
requirements, 241
supplementation, 257-258, 734
vitamin supplementation, overview of,
732-735

Humidity and vitamin stability, 715
Huntington’s disease, 587
Hydralazine, vitamin B, antagonism, 385
Hydrogen peroxide, 165
Hydroperoxides, 169, 173
Hydroxyproline, 605
Hyperkeratosis, 63
Hypertrophic osteodystrophy, canine, 621
Hypothyroidism, vitamin A deficiency
and, 59
Immune system
autoimmune disease, 683, 684
p-carotene and, 76
fatty acids and, 699, 707
folacin deficiency, 490
pantothenic acid deficiency, 430,438
prostaglandins, 682-684
vitamin A and, 37-38,53
vitamin B, deficiency, 393-394, 406
vitamin C and, 606, 611, 619, 623
vitamin D and, 110
vitamin E and, 163, 167-168, 172-173,
180, 184, 191-192,198-199,203,
205,209-210
Infantile scurvy (Barlow’s disease), 600,
625-626
Infections
fatty acids and, 699, 700
thiamin requirements and, 276
vitamin A and, 30, 34-35, 37-38
vitamin C and, 619-621
vitamin E and, 167, 191, 198
vitamin requirements, effects on, 721
Inositol. See myo-inositol
Insulincholine and, 566
vitamin D and, 110
Interferons and vitamin C, 606
Interleukin (IL-2), 684
International chick unit (ICU), 95
Intestinal membrane Ca-binding protein(IMCal), 105
Intrinsic factor, 529-531, 551-552,
557-558,733
Iodopsin, 31
Ironantioxidant, 606
cobalt absorption and, 531
folacin deficiency, 51 1
phytate and, 665
riboflavin deficiency, 320
vitamin C and, 607-608,633-634

775

INDEX
Iron (cont.)
vitamin destruction by, 715
vitamin E and, 177, 193
Irradiation of food, 93, 717
Isoniazid, 387, 411
Isoprostanes, 683
Isotretinoin, 73
Japanese quailbiotin requirements, 453
choline requirements, 577
essential fatty acids (EFA) requirements,
690
folacin requirements, 492
niacin requirements, 358
pantothenic acid requirements, 427
riboflavin requirements, 322
thiamin requirements, 277
vitamin B, requirements, 395
vitamin B,, requirements, 536
vitamin D requirements, 114
Keratinization
retinoid therapy, 73
vitamin A deficiency, 28-30, 34, 48, 53,
54,58,59-60,62,64
Ketosis, 364, 377, 378, 648
Kidneycalcification, 141, 145
essential fatty acids (EFA) deficiency,
70 1
vitamin A storage, 27-28
vitamin D
effects on Ca and P reabsorption, 100,
107
metabolism, 99-100
Krebs cycle. See Citric acid cycle
Laboratory animals. See also Gerbils;
Guinea pigs; Hamsters; Mice; Ratsbiotin deficiency, 467-468
carnitine deficiency, 651
choline deficiency, 580, 586
essential fatty acids (EFA) deficiency,
70 1
folacin deficiency, 504-505
niacin deficiency, 371-372
pantothenic acid deficiency, 437-438
riboflavin deficiency, 334-335, 336
thiamin deficiency, 281, 295
vitamin A
deficiency, 62
requirements, 40

776

vitamin B, deficiency, 404-405
vitamin B,, deficiency, 547
vitamin C deficiency, 623
vitamin D deficiency, 129
vitamin E deficiency, 202
vitamin K deficiency, 251
Lactobacillus casei, 388, 480, 484
Lactobacillus citrovorum, 484
Lactobacillus lacatis, 524
Lactobacillus leichmannii, 528
Lactobacillus plantarum, 350, 422, 448
Laetrile, 670
L-Dopa, vitamin B, antagonism, 388
Least-cost formulations, 720
Lecithin, 566-569, 571, 576, 587-589.
See also Choline
Legumes, cobalt and, 540
Leucineleucine mutase, 532
toxicity, 375
tryptophan-niacin conversion, 354
Leuconostoc mesenteroides, 350
Leukotrienes, 683-684
Light. See Sunlight; Ultraviolet (W)light
Lind, James, 11, 599
Linoleic acid, 675-686, 689-694,
696-702, 704-708. See also Essential
fatty acids (EFA)
conjugated (CLA), 678, 686, 693-694,
706
Linolenic acid, 675-677, 679-682,
685-693,696,700,702, 704,
706-707. See also Essential fatty acids
Lipid metabolism. See also Essential fatty
acids (EFA)
biotin, 450-452
choline, 571
folacin, 487
myo-inositol and, 661
niacin and, 356
pantothenic acid, 424-425
vitamin B,, 391
Lipoic acid (thioctic acid), 669
Lipoproteinscholine, 571, 587
fatty acids, 678, 685, 706
vitamin C, 607
vitamin E, 162, 204, 216
Liverbiopsy, 28,29, 68-69
carn itin e
deficiency, 647

INDEX
synthesis, 643
cholinedeficiency, 566, 571, 580, 585-587
synthesis, 565, 569
fatty acid deposition, 679
folacin storage, 510
hepatosis dieterica, 188-189, 190
myo-inositol deficiency and, 662-663
retinol secretion, 25
thiamin phosphorylation, 290
vitamin A
storage, 27-28, 39, 67
toxicity and liver consumption, 79-80,81vitamin B,,
as source of, 523-525
storage, 531
vitamin D transformation, 98-99
vitamin E
deficiency, 202
storage, 163-164
vitamin K storage, 234
Lordosis, 621, 622
Low-density lipoprotein (LDL)
niacin, 378
vitamin A transport, 25
vitamin C, 607
vitamin E, 162,204, 216
Lumisterol, 93, 98
Lunin, N., 11
Lycopene, 19,78
Lysophosphatidylcholine, 566, 568
Lysyl oxidase, 107, 666
Magnesium
absorption, 105
deficiency, 124
Manganeseantioxidant, 606
vitamin E and, 177
Mastitis, 74
p-carotene supplementation and, 76
mastitis-metritis-agalacticasyndrome(MMA), 189-190
vitamin A and, 38, 52-53
vitamin E and, 210-212
Matrix Gla protein, 238, 239
McCollum, E.V., 12, 16, 17
Measles, vitamin A deficiency and, 75
Meat, carnitine content of, 646
Megablastic anemia. See Pernicious anemia
Mellanby, Sir Edward, 92

Membranes
carnitine and, 650
choline and, 566, 569, 571
fatty acids and, 682, 684, 685
myo-inositol and, 661
polyunsaturated fatty acid (PUFA)content, 172
riboflavin deficiency and, 321
selenium and, 173
vitamin C and, 606
vitamin E and, 165, 166, 173, 203,
213
Menadione (K3), 229,230,232-234,
256-258, 258. See also Vitamin K
Menaquinones (vitamin K2), 229, 230,
232-233, 242, 251. See also Vitamin K
Mendel, L. B., 12, 16
Metabolic acidosis, 135
Metabolism. See specific vitamins
Metalloenzymes, 606
Methionine
choIine and, 572,574-575,581-583,
584,587,589-591
folacin and, 487-490, 512
vitamin B, and, 396
vitamin B,, and, 534, 535
vitamin E and, 170
vitamin U (cabagin), 671
Methionine synthetase, 532, 549, 552
Methotrexate, 481, 483, 493, 591
Methylcobalamin, 532-533
Methylmalonyl CoA isomerase (mutase),
532,534-535,537-538,549,554
Mice. See also Laboratory animals
biotin deficiency, 468
choline deficiency, 586
essential fatty acids (EFA)
deficiency, 701
requirements, 690
folacin deficiency, 504-505
myo-inositol deficiency, 662
niacin requirements, 358
pantothenic aciddeficiency, 438
requirements, 427
pyrroloquinoline quinone (PQQ) deficiency, 666
riboflavindeficiency, 334
requirements, 322
thiamin
deficiency, 295
requirements, 277

777

INDEX
Mice (cont.)
vitamin A
deficiency, 62
requirements, 40
vitamin B,
deficiency, 405
requirements, 395
vitamin B,, deficiency, 547
vitamin D requirements, 114
vitamin E
deficiency, 202
requirements, 171
vitamin K deficiency, 251
Micro biological assays
biotin, 448
choline, 568
folacin, 484
myo-inositol, 660
pantothenic acid, 422
vitamin B,, 388-389
vitamin B,,, 528, 552
Microorganisms
biotin synthesis, 445,449, 452
choline synthesis, 579
fatty acid metabolism, 676, 696
folacin synthesis, 479480, 490,
496497,512
myo-inositol synthesis, 662
niacin synthesis, 368, 377
pantothenic acid synthesis, 419, 428,
431
phytase, 665
riboflavin biosynthesis, 322-323
thiaminase production, 269
thiamin synthesis, 269, 274-275, 283,
292
vitamin B, synthesis, 394
vitamin B,, synthesis, 523, 525,536-537,538,547
vitamin K synthesis, 227, 229, 239-240,
243-244,248-249
vitamins as growth factors for, 13
vitamin synthesis by, 6, 7
Milkcarnitine content, 646
choline content, 567, 591
essential fatty acids (EFA) content, 694,
706
folacin content, 491, 493, 494, 495
pantothenic acid content, 429,439
production and niacin supplementation,364-365,377
riboflavin content, 325

778

thiamin content, 278-279
vitamin B, content, 398, 4 0 6 4 0 7
vitamin B,, content, 539, 540
vitamin C content, 615, 616
vitamin D content, 117, 132, 139
vitamin E content, 162-163, 175-176,
213
vitamin K content, 243, 252-253
Milk fever, 121-122, 134-136
Minkbiotindeficiency, 4 6 6 4 6 7
requirements, 453
essential fatty acids (EFA) requirements,
690
folacin
deficiency, 504
requirements, 492
niacin deficiency, 371
pantothenic acid deficiency, 437
riboflavindeficiency, 334
requirements, 322
thiamin
deficiency, 282, 295
requirements, 277, 302-303
vitamin A requirements, 40
vitamin B,
deficiency, 404
requirements, 395
vitamin B,, deficiency, 548
vitamin C and, 623
vitamin D deficiency, 129
vitamin E
deficiency, 201
requirements, 171
Molasses toxicity, 287
Molds. See also Mycotoxins
thiamin content and, 276, 278
vitamin D and, 138
Monkeys. See Primates, nonhumanMonosodium glutamate and vitamin B,,
388,411
Mulberry heart disease, 193
Muscular dystrophy, 156
serum enzyme levels, 206
vitamin E deficiency, 178-186,
188-189, 195-198,200-203
Mycotoxins, 722
folacin supplementation, 513
vitamin K and, 230,255
vitamin supplementation, 726
Myo-inositol

INDEX
chemical structure and properties, 660,
661
deficiency, 662-664
functions, 660-661
introduction to, 660
metabolism, 660
requirements, 662-663
sources, 661-662
Myopathy. See also Muscular dystrophy
vitamin B,, deficiency, 546
vitamin E deficiency, 180, 198-199
Naphthalenes, 53
National Research Council (NRC),
723-727
Neoplasia. See Cancer
Nervous systemcholine and, 587, 591
essential fatty acids (EFA), 687, 696,
701,702,706-707
folacin deficiency, 510
niacin deficiency and, 367, 374
pantothenic acid deficiency, 430-433,
436438thiamin and, 272-274,280-282,
283-288,292-299
vitamin B, and, 399, 400, 401, 402,
404-405,406,412413
vitamin B,, and, 533, 545, 546, 548,
549
vitamin E and, 170, 194-195, 198, 205
Neural tube defects
folacin and, 479,493, 508, 514
vitamin B,, deficiency, 552
Neurospora crassa, 568
Newcastle disease, 38, 256
Niacin (vitamin B3), 347-383
analytical procedures, 350-351
bioavailability, 719
chemical structure and properties, 350
deficiency, 362-376
assessment of status, 375-376
cats, 369
dogs, 369, 371
fish, 370-371
foxes and mink, 371
horses, 368
humans, 372-375
laboratory animals, 371-372
poultry, 367-368, 369, 370
primates, nonhuman, 372
rabbits, 372
ruminants, 363-365

cattle, beef, 363-364
cattle, dairy, 364-365
lambs, 365
swine, 365-367
functions, 355-357
history, 347-349
introduction to, 347
metabolism
absorption, 351-352
excretion, 352-353
storage, 352
tryptophan conversion, 353-355
requirements, 357-360
factors influencing, 358-360
table of species, 358
sources, natural, 360-362
stability, 379
supplementation, 376-379, 728-735,
737
toxicity, 379-380
vitamin B, and, 388, 390
Nicotinamide adenine dinucleotide(NAD), 350-352,355-357
Nicotinamide adenine dinucleotide phosphate (NADP), 350-352,355-357
Nicotinic acid. See Niacin (vitamin B3)
Night blindness (nyctalopia), 16-17, 19,
32-33,51,53-54,59-61,63-64,69
Nitrates and vitamin A nutrition, 42
Nitrogen metabolism
carnitine, 645
vitamin B,,, 534
Nomenclature, vitamin, 5-6
Novocaine, 671
Nucleic acids
folacin deficiency and, 489
vitamin B,, and, 533-534

Ocbromonas malbamensis, 528
Oilsessential fatty acids (EFA) content,
696-695
vitamin E content, 175, 177
vitamin K content, 243
Oleic acid, 676-678, 680-683, 689,
704-705
Opsin, 32
Oral contraceptives
folacin and, 493
vitamin B, and, 385, 396, 407, 722
vitamin C deficiency, 626
Orotic acid, 670
Osborne, T. B., 12, 16

779

INDEX
Osteocalcin, 133, 238, 248
Osteomalacia, 101, 106, 118, 121, 131:
132,133
Osteoporosis, 118
Oxythiamin, 267

PABA (p-aminobenzoic acid), 481,
667-668
Palmitoleic acid, 675-677, 680-681
Pangamic acid, 670
Pantothenic acid, 419-444
analytical procedures, 422-423
antagonist, 420
chemical structure and properties,
420422
deficiency, 430-439
assessment of status, 439
cats, 436
dogs, 436
fish, 436-437
foxes and mink, 437
horses, 436
humans, 438-439
laboratory animals, 437-438
poultry, 433-435
primates, nonhuman, 438
ruminants, 431
signs and symptoms of, 430-431
swine, 43 1-433
functions, 424-426
history, 419-420
introduction to, 419
metabolism, 423-424
requirements, 426-429
factors influencing, 426-429
table of species, 427
sources, natural, 429-430
stability, 421-422, 430, 440
supplementation, 439441, 729-730,
732, 737
toxicity, 441
vitamin B,, and, 535
Parasitic infestation, vitamin A supplementation and, 37
Parathyroid hormone (PTH), 100,
104-107, 109,121,136
Parkinson’s diseasevitamin B, and, 412
vitamin E and, 203
Parturient paresis (milk fever), 134-136
Pasteur, Louis, 10
Pauling, Linus, 611
Pellagra, 347, 348, 372-375, 378

780

Pelleting and vitamin stability, 715,
716-71 7
Penicillamine, 385, 41 1
Pernicious anemia, 523-524, 529, 532,
540,548452,557-558
Perosisbiotin deficiency, 460, 461, 463
choline deficiency, 566, 571, 583
folacin deficiency, 499, 501
niacin deficiency, 368
vitamin B,, deficiency, 545
Pet foods
vitamin A supplementation, 72-73
vitamin A toxicity, 79
vitamin E content, 214-215
Phalaris staggers, 543
Phenytoin, 484
Phosphatidylcholine, 566, 588. See also
Lecithin
Phospholipase, 568, 569
Phospholipids, 165, 166,203, 566, 569,
571, 573, 575, 682, 684
Phosphorus
absorption, 113-114
absorption by ruminants, 108-109
excretion, 109
phytates and, 664-665
rickets, 117-118
transport, 106
vitamin D, 100, 103intestinal effects, 104-106
renal reabsorption of, 107
requirements, effect on, 113
Phylloquinone (vitamin K), 229, 230,
232-234,242,249,254,257-258. See
also Vitamin K
Phytate and myo-inositol, 660, 662,
664-666
Picolinic acid carboxylase, 355, 356, 360
Pigeons and thiamin deficiency, 281, 291
Pigs. See Swine
Pinkeye, 48
Placenta, retained, 187
Placental transfer
vitamin D, 102
vitamin E, 162-163
Plants. See also Sources of vitamins
as carotene source, 43-47
riboflavin and, 324-325
vitamin D toxicity from, 142-145
Plasmalogens, 566, 571, 573, 587
Platelets, 166-167, 686, 687, 688-689,
702. See also Blood clottingINDEX
Polioencephalomalacia (PEM), 283-288,
301,302-304
Polyneuritis, 266, 276, 280, 290-292
Polyphenols, 668-669
Polyunsaturated fatty acids. See PUFAs(polyunsaturated fatty acids)
Poultry
biotindeficiency, 460-464
status assessment, 470-471
supplementation, 471, 472,473
carnitine deficiency, 649-650
cholinedeficiency, 580, 583-584
requirements, 575-576, 577
confinement rearing, 721
essential fatty acids (EFA)
deficiency, 699
supplementation, 704
fatty liver and kidney syndrome (FLKS),
464
folacin
deficiency, 499-502
requirements, 490-491,492
supplementation, 512, 513
niacindeficiency, 367-368, 369, 370
supplementation, 376, 379
NRC vitamin requirements, 726-727
pantothenic acid deficiency, 433-435
phytates, 664
riboflavindeficiency, 330-332
requirements, 322
supplementation, 340-341
thiamin
deficiency, 281, 289-292
requirements, 276, 277
supplementation, 302
UGFs (unidentified growth factors),
6 72
vitamin A
deficiency, 33, 37-38, 57-59
requirements, 40
vitamin B,
deficiency, 401-403
supplementation, 409-410
vitamin B,, deficiency, 545-546
vitamin C, 617-620
requirements, 609
supplementation, 629
vitamin D
deficiency, 101, 124-127

and embryonic development, 109-110,
126-127
supplementation, 138-1 39
vitamin E deficiency, 193-197
vitamin K
deficiency, 247-249
supplementation, 254-256
vitamin supplementation
elimination of, 727
overview of, 729
Pregnancy toxemia, carnitine deficiency
and, 648
Premixes, formulating vitamin, 741-742
Primates, nonhumanbiotin deficiency, 468-469
choline deficiency, 586
essential fatty acids (EFA)
deficiency, 701
requirements, 690
folacin deficiency, 505
niacin deficiency, 372
pantothenic acid deficiency, 438
riboflavin deficiency, 335
thiamin deficiency, 296
vitamin A
deficiency, 30, 33, 36, 62
requirements, 40
vitamin B, deficiency, 405
vitamin B,, deficiency, 548
vitamin C deficiency, 623
vitamin D deficiency, 129-130
vitamin E deficiency, 202
vitamin K deficiency, 251
Processing and storage effects on vitamins, 714-717
Propionic acid and vitamin B,,, 535,
537-53 8
Prostaglandins, 166, 682-684, 697
Protein metabolism
biotin, 450-451
folacin, 487, 488
niacin, 356
pantothenic acid, 424, 426
vitamin B,, 385, 391-394, 408
vitamin B,,, 533-534
Prothrombin, 235,245, 253-254
Provitamin A, 18, 19, 20, 43
Pseudovitamins, 659
Psoriasis and vitamin D, 110, 133, 140
Pteroylglutamic acid (PGA). See Folacin
PTH. See Parathyroid hormone (PTH)
PUFAs (polyunsaturated fatty acids),
179-180, 186,200,214-215. See also

78 1

INDEX
PUFAs, (cont.)
Essential fatty acids (EFA)
cancer and, 688
infections, modulating, 697-698
vitamin E and, 161, 170-172
Pumpkins as carotene source, 45
Purine synthesis
folacin and, 489
vitamin B,,, 533
Pyridoxal. See Vitamin B,
Pyridoxal phosphate (PLP), 386-393,
396,407-409. See also Vitamin B,
Pyridoxamine. See Vitamin B,
Pyridoxine-S’-P-D-glucoside(PNG),
398-3 9 9Pyrimidine synthesis
folacin and, 489
vitamin B,,, 533
Pyrithiamine, 267, 722
Pyrroloquinoline quinone (PQQ),
666-667
Rabbits
biotin deficiency, 469
cholinedeficiency, 580, 586
requirements, 577
coprophagy, 7
essential fatty acids (EFA)deficiency, 702
niacin deficiency, 372
riboflavin deficiency, 335
thiamin deficiency, 281, 296
vitamin A
deficiency, 36, 62-63
requirements, 40
vitamin B, deficiency, 405
vitamin B,,
deficiency, 548
requirements, 536
vitamin C and, 624
vitamin E
deficiency, 202-203
requirements, 171
vitamin K
deficiency, 251
requirements, 241
Radioimmunoassay. See Analytical proceduresRats. See also Laboratory animals
carnitine deficiency, 653
cholinedeficiency, 586
requirements, 577

782

choline toxicity, 592
coprophagy, 7
essential fatty acids (EFA)
deficiency, 701
requirements, 690
folacin
deficiency, 504
requirements, 492
myo-inositol deficiency, 662
niacindeficiency, 372
requirements, 358
pantothenic aciddeficiency, 437
requirements, 427
riboflavindeficiency, 334, 336
requirements, 322
thiamin
deficiency, 281, 295
requirements, 277
vitamin A
deficiency, 30, 33, 36, 62
requirements, 40
vitamin B,
deficiency, 385-386, 404-405
requirements, 395
vitamin B,,
deficiency, 547
requirements, 536
vitamin D
deficiency, 129
requirements, 114
vitamin E
deficiency, 202
requirements, 171
vitamin K
deficiency, 251
requirements, 241
Reproduction
P-carotene effects on, 76-77
biotin deficiency, 460,464,468
choline deficiency, 582-583, 584
cobalt deficiency, 541
fatty acids and, 696-697, 699, 700-702
folacin
deficiency, 479,498-499, 500-501,
505,506,508
requirements, 491, 493
supplementation, 512-515
pantothenic acid deficiency, 432-435,
437
riboflavin deficiency, 327-330, 331

INDEX
thiamin deficiency, 292
vitamin A, 36, 50-51, 54, 55-57, 60
vitamin A deficiency, 36
vitamin B,, 404-405
vitamin B,,, 544-545, 545
vitamin C, 608-609,616-617,
617-618, 623,624,629
vitamin D, 109-110,126-127
vitamin E, 155-156, 166, 179,
186-188, 191,193,196, 199,202
Requirements
biotin, 452-453
carnitine, 645
choline, 574-576,577
effects on
antagonists, vitamin, 722
antimicrobials, use of, 722-723
body reserves, 723
confinement rearing, 721
other nutrient levels, 723
physiological makeup, 720
production function, 720
stress and disease, 721-722
essential fatty acids (EFA),689-693
folacin, 490-493
myo-inositol, 662-663
niacin, 357-360
pantothenic acid, 426-429
riboflavin, 321-324
thiamin, 274-277
vitamin A, 39-43
vitamin B,, 394-396
vitamin BIZ, 535-538
vitamin C, 609-611
vitamin D, 111-115
vitamin E, 169-174
vitamin K, 239
Retinal degeneration and vitamin E deficiency, 199
Retinoic acid, 18. See also Vitamin A
collagenase inhibition by, 33
embryonic development and, 31
as hormone, 30
testosterone synthesis, 36
Retinoic acid receptors (RAR), 26-27,
30-3 1Retinoid therapy, 38, 73, 75
Retinol, 18-21,2447. See also Vitamin
A
Retinol-binding protein (RBP), 25-27, 48
Reye’s syndrome, 645
Rhodopsin, 31-32
Riboflavin (vitamin B2), 31 1-346

analytical procedures, 3 14-315
antagonists, 314
chemical structure and properties,
3 12-3 14
deficiency
assessment of status, 339-340
cats, 333-334
dogs, 333
fish, 334
foxes and mink, 334, 335
horses, 332
humans, 337-339
laboratory animals, 334-335, 336
poultry, 330-332, 333
primates, nonhuman, 335
rabbits, 335
ruminants, 326-327
swine, 327-330
food processing and storage; effects of,717
functions, 317-321
history, 311-312
introduction to, 3 11
metabolism
absorption, 315, 342
digestion, 315
excretion, 317
storage, 316-317
transport, 316, 342
requirements, 321-324
factors influencing, 323, 324
table of species, 322
sources, natural, 324-326
supplementation, 340-341, 729-730,
732, 734,737
toxicity, 342
Riboflavin-binding protein, 346
Rickets, 11, 17, 91-93, 98, 111, 739. See
also Vitamin D, deficiency
air pollution and, 112biological assay for vitamin D, 96
cats, 128
dogs, 127-128
foxes and mink, 129
horses, 127
humans, 130-132
laboratory animals, 129
poultry, 124-126
primates, nonhuman, 129-130
ruminants, 119-121
signs and symptoms, 117-119
swine, 123
Ringworm (Trichophyton verrucosum), 37

783

INDEX
Rodenticides, cholecalciferol, 146
Roundworms, 37
Ruminants. See also Cattle; Goats; Sheepbiotindeficiency, 4 5 6 4 5 7
supplementation, 472-473
calcium absorption, 107-108
carnitine deficiency, 648
cholinedeficiency, 579-581
metabolism, 569-570
requirements, 576, 577
supplementation, 590-591
cobalt supplementation, 556-557
essential fatty acids (EFA)supplementation, 696-697,704-705, 707-708
fatty acid metabolism, 676, 679
folacin
deficiency, 4 9 6 4 9 7
supplementation, 513
myo-inositol, 662
niacindeficiency, 363-365
requirements, 359
supplementation, 377
pantothenic acid deficiency, 431
phosphorus
absorption, 108-1 09
excretion, 109
phytates, 664
riboflavindeficiency, 326-327
requirements, 311, 323
thiaminabsorption, 269-270
deficiency, 275, 280-289
supplementation, 302-303
vitamin A
deficiency, 48-55
digestion, 22
storage, 67-68
toxicity, 78
vitamin B,
deficiency, 399
supplementation, 409
vitamin B,,
deficiency, 541-543
requirements, 536, 537-538
status determination, 554
supplementation, 555-556
vitamin C deficiency, 613-615
vitamin DD, versus D,, 117784

deficiency, 119-123
vitamin E
deficiency, 181-188
supplementation, 209-2 13
vitamin K deficiency, 244-245
vitamin requirements, 6
vitamin supplementation, overview of,
728-729

Saccharomyces auidinii, 447
Saccbaromyces cerevisiae, 448
Saccharomyces uvarum, 388
S-adenosylmethionine (SAM),549-550,
569,572,575
Schizophrenia, 374, 378, 611
Scoliosis, 205, 621, 622
Scurvy, 19-20. See also Vitamin C , deficiency
carnitine and, 643
clinical signs, 624
history, 597-600
infantile scurvy (Barlow’s disease), 600,
625-626
Seaweed, 539
Secadera, 289
Selenium (Se). See also Vitamin E
antioxidant, 606
assessment of, 207
deficiency in dogs, 199
deficiency in horses, 197-198
deficiency in poultry, 194-197
deficiency in ruminants, 181-1 87
deficiency in swine, 188-193
immune response and, 168
protection from toxic substances, 169
soil content, 181
supplementation, 734
vitamin E and, 156, 165, 169, 173
white muscle disease, 181-186
Sheepbiotin requirements, 453
calcium
absorption, 107transplacental movement, 102cholinemetabolism, 570
requirements, 577
cobaltdeficiency, 525, 537, 538, 541-543
supplementation, 557
essential fatty acids (EFA)requirements,
690
folacin requirements, 492

INDEX
niacindeficiency, 365
requirements, 358
pantothenic acid requirements, 427
phosphorus absorption, 108-109
pregnancy toxemia, 648
riboflavindeficiency, 327
requirements, 322
thiamin
deficiency, 280, 283-284, 286
requirements, 277
supplementation, 303
vitamin A
deficiency, 36, 53-54
requirements, 40
storage, 67
supplementation, 73
vitamin B, requirements, 395
vitamin B,,
deficiency, 541-543
requirements, 536
status determination, 554
supplementation, 555
vitamin D
deficiency, 121, 122-123
requirements, 114
vitamin E
deficiency, 181, 183-185
requirements, 171
supplementation, 163
vitamin K requirements, 241
Skin. See also Dermatitis/dermatosis
essential fatty acids (EFA) deficiency
and, 695-696,697-698,700-702
retinoids, 38
vitamin D production, 100, 111-113,
130, 140
Smoking and vitamin C requirements, 610
Solanum esuriale, 143
Solanum malacoxylon, 143, 144, 145
Solanum torvum, 143
Sources of vitamins
biotin, 454-455
carnitine, 646-647
carotene and carotenoids, 44-47
cobalt, 540
essential fatty acids (EFA), 693-695
flavonoids, 668-669
folacin, 494-495
myo-inositol, 661-661, 661-662
niacin, 360-362
pantothenic acid, 429-430

pyrroloquinoline quinone (PQQ), 666
riboflavin, 324-326
thiamin, 278-279
vitamin A, 43-48
vitamin B,, 397-399
vitamin B,,, 538-540
vitamin C, 611
vitamin D, 115-117
vitamin E, 174-178
vitamin K, 242-243
Soybeanscarnitine content, 655
choline content, 578, 589
phytates, 665-666
vitamin B, content, 398, 409-410
Spermatogenesis. See Reproduction
Sphingomyelin, 566, 568, 571, 576
Spiking syndrome, 722
Spina bifida, 479, 508
Spraddled-leg disease, 566, 581, 582
Squash as carotene source, 45
Stability of vitamins, 714-717, 739-740.
See also specific vitamins
Star thistle (Centaurea solstitialis), 293
Steatitis (yellow fat disease), 172,
200-201,201
Steenbock, 16
Storagecarnitine, 643-644
choline, 567
effect on vitamin requirements, 723
folacin, 486-487
niacin, 352
pantothenic acid, 423-424
riboflavin, 316-317
selenium, 173-1 74
thiamin, 270-271
vitamin A, 27-28
vitamin B,, 390
vitamin B,,, 531-532, 551
vitamin C, 603
vitamin D, 101-102
vitamin E, 163, 173-174
vitamin K, 234
Streptavidin, 447, 448, 722
Streptococcus faecalis, 388
Streptococcus lactis, 480
Stressmyo-inositol and, 663
vitamin C and, 603, 607, 610, 615-620,
627-628
vitamin E and, 167, 173, 193,210,211,
214

785

INDEX
Stress (cont.)
vitamin requirements, effects on, 721
Stuart-Prower factor, 235
Sudden infant death syndrome (SIDS),
470
Sulfonamides, 722-723
acetylation by coenzyme A, 420, 424
biotin deficiency, 448, 469
folacin and, 483-484,498, 503, 504
PABA, 668
vitamin K and, 230, 245, 255
Sulfurcholine and, 581, 584, 590
thiamin and, 284-286, 302, 303
Sunlight
exposure and vitamin D, 91-93, 97-98,
111-112, 122-124, 127, 130-132,
134,139-140
vitamin destruction by, 716
Superoxide anion radical, 165
Superoxide dismutase, 606
Supplementation, 713-745
biotin, 471-474
carnitine, 653
choline, 588-591
cost, 713
dietary vitamin intake, effects on,
714-720agronomic effects, 714
harvesting conditions, 714
least-cost feed formulation, 720
processing and storage, 714-717
reduced feed intake, 717-718
vitamin bioavailability, 719-720
vitamin variability, 718-719
essential fatty acids (EFA), 704-708
folacin, 512-515
forms of, 735-741
horses, 730-732
humans, 732-735
introduction to, 713-714
megadose, 632
niacin, 376-379
pantothenic acid, 439-441
poultry, 729
premixes, formulating, 741-742
riboflavin, 340-341
ruminants, 728-729
swine, 729-730, 731
synthetic vitamins, advantages of,
740-74 1
thiamin, 301-305
vitamin A, 69-75

786

intramuscular injection, 71-72
liquid, 70-71
in mineral mixture, 71
reasons for, 70
stability, 69-71
vitamin allowances, optimum, 723-727
vitamin B,, 409-412
vitamin B,,, 555-558
vitamin C, 627-633
vitamin D, 134-140
vitamin E, 207-216
vitamin requirements, effects on,
720-723
antagonists, vitamin, 722
antimicrobials, use of, 722-723
body reserves, 723
confinement rearing, 721
other nutrient levels, 723
physiological makeup, 720
production function, 720
stress and disease, 721-722
Surfactant, 687
Sweet clover poisoning, 230, 244
Sweet potatoes as carotene source, 45
Swinep-carotene supplementation, 77
biotinrequirements, 453
status assessment, 470-471
supplementation, 458, 471-473
carnitine
deficiency, 649
supplementation, 649, 653
cholinedeficiency, 580, 581-583
requirements, 574-575, 577
supplementation, 589, 590
toxicity, 592
confinement rearing, 721
essential fatty acids (EFA)
deficiency, 697-699
requirements, 690
supplementation, 707
folacin
deficiency, 497-499
requirements, 492
supplementation, 499, 512-513
hepatosis dietetica, 188-189, 190
mastitis-metritis-agalactica syndrome,189-190
niacindeficiency, 365-367
requirements, 358

INDEX
supplementation, 366, 376-377
pantothenic aciddeficiency, 431-433
requirements, 427, 428
production function and vitaminrequirements, 720
restricted feeding and vitamin intake,718
riboflavindeficiency, 327-330
requirements, 321-322, 323
supplementation, 340-341
thiamin
deficiency, 282, 289, 290
requirements, 277
supplementation, 302
UGFs (unidentified growth factors), 672
vitamin A
deficiency, 36, 38, 39, 55-57
requirements, 40
storage, 68
supplementation, 72
vitamin B,
deficiency, 400-401
requirements, 395
vitamin B,,
deficiency, 543-545
requirements, 536
vitamin C
deficiency, 615-617
requirements, 609
supplementation, 629
vitamin D
deficiency, 123-1 24
requirements, 114
supplementation, 138
toxicity, 141
vitamin E
deficiency, 156, 188-193
requirements, 171, 172
supplementation, 215
vitamin K
deficiency, 245-246
requirements, 241
supplementation, 246, 256
vitamin supplementationin disease conditions, 725-726
elimination of, 727
overview of, 729-730, 731
Tachysterol, 98
Takaki, K., 11, 266
Tall fescue toxicosis, 286, 302

Temperature and food consumption, 718
Testosterone and retinoic acid, 36
Tetrahydrofolate (THF), 487, 533-534.
See also Folacin
Thiaminases, 267-269, 287-289, 292,
293-294, 717, 722
Thiamin pyrophosphate (TPP), 270,272,
273,301
Thiamin (vitamin B,), 265-310
analytical procedures, 269
antagonists, 267-269, 276, 287, 299,
303,722
chemical structure and properties, 267,
268
deficiency
assessment of status, 300-301
beriberi, 265-267,280, 296-299
cardiovascular disorders, 280, 289,
294,296-298
cats, 281, 293-294
clinical signs, table of, 280-281
crocodiles, 296, 297
dogs, 281,293-294
fish, 294
foxes and mink, 282, 295
horses, 282, 292-293
humans, 281,296-300
laboratory animals, 281,295
neurological disorders, 280, 283-288,
292-299
polioencephalomalacia, 283-288, 301,
302-304
polyneuritis, 290-292
poultry, 281, 289-292
primates, nonhuman, 296
rabbits, 281, 296
ruminants, 280-289
swine, 282,289,290
food processing and storage, effects of,717
functions, 271-274
decarboxylations, 271-272
nervous system and, 272-274
history, 265-267
introduction to, 265
metabolism
absorption, 269-270,274
digestion, 269
excretion, 271
phosphorylation, 270
storage, 270-271
transport, 270
requirements, 274-277

787

INDEX
Thiamin, requirements (cont.)
factors influencing, 274-277
table of species, 277
sources, natural, 278-279
stability, 278-279, 302, 304
supplementation, 301-305, 728-734,
737
toxicity, 305
Thioctic acid, 669
Thrombin, 235, 253
Thyrocalcitonin (calcitonin). See Calcium
Thyroid hormone
retinoic acid and, 30-31
riboflavin and, 316, 324
Tocopherol, 607-608. See also Vitamin E
Toxicity. See specific vitamins
Transaminases, 391-392
myo-inositol and, 663
vitamin B, and, 408
Transcalciferin, 101
Transcobalamins, 530
Transcription regulation, 103
Transforming growth factor (TGFb), 684
Transketolase, 272, 301
Transportbiotin, 449-450
calcium, 105-106
carnitine, 642-643
choline, 569
fatty acid, 679
folacin, 486
phosphorus, 106
riboflavin, 316, 342
thiamin, 269
vitamin A, 24-27
vitamin B,, 390
vitamin BIZ, 530, 551
vitamin C, 603
vitamin D, 99-100, 101
vitamin E, 162
vitamin K, 233
Transthyretin, 25
Tricarboxylic acid (Krebs) cycle. See Citricacid cycle
Trichuris suis, 38
Trimethylamine, 569, 570, 588-589
Trisetum flavescens, 143, 145
Tryptophan
niacin and, 347, 349, 353-355,
357-360, 358-360,363-377
vitamin B, and, 396,406
Turkeysbiotin788

deficiency, 460-464
requirements, 453
cholinedeficiency, 580, 583
requirements, 577
essential fatty acids (EFA) requirements,
690
folacin
deficiency, 499-501
requirements, 492
niacindeficiency, 367-368, 370
requirements, 358
pantothenic aciddeficiency, 435
requirements, 427
restricted feeding and vitamin intake,718
riboflavindeficiency, 331, 333
requirements, 322
thiamin requirements, 277
vitamin A requirements, 40
vitamin B,
deficiency, 402-403
requirements, 395
vitamin BIZrequirements, 536
vitamin C supplementation, 629
vitamin D
deficiency, 124, 125
requirements, 114
vitamin E
deficiency, 196
requirements, 171
supplementation, 212
vitamin K requirements, 241
Tyrosine and vitamin C, 607
Ubiquinones, 669-670
Ultraviolet (W)light, 93, 97-98,
111-112, 124, 130,145, 716
Unidentified growth factors (UGFs),
671-672
Urinary calculi, 55, 407-408, 411,
633-634
Vedder, Edward B., 266
Vegetables. See also Foods and feeds
as carotene source, 45choline content, 578
folacin content, 495
vitamin C and, 599, 611-612
vitamin K content, 242

INDEX
vitamin losses from, 717
Vegetarian diets and vitamin B,,, 540,
550-551,558
Very-low-density lipoprotein (VLDL),
571
Visionnight blindness (nyctalopia), 16-17, 19,
32-33,51,53-54,59-61,63-64,
69
vitamin A functions, 31-34
xerophthalmia, 12, 16-17, 19, 33, 51,
58,61-64
Vitamin A, 15-90
analytical procedures, 21-22
chemical structure and properties,
18-21
cis versus trans forms, 18
units of activity, 20-21
deficiency, 48-69
bone development and, 36, 52, 58
cats, 60-61
causes of, 65-66,74
dogs, 33, 61
drought and, 66
fish, 61
foxes, 33, 62
horses, 33, 59-60
humans, 33,63-65
immune system and disease, 37-38,
53,64
incidence and relationship to bodystores, 33, 65-68
infections and, 34-35, 37-38
keratinization, 28-30, 34-35, 48, 53,
54,58,59-60,62,64
laboratory animals, 30, 33, 36, 62
night blindness, 16, 17, 32-33, 51, 53,
54,59, 60,61,63-64
poultry, 33, 37-38,57-59
primates, nonhuman, 62
rabbits, 36, 62-63
reproduction and, 30, 36, 50-51, 54,
55-57,60
reproductive effects, 36
ruminants, 48-55
cattle, 33-34, 36, 38, 50-53
goats, 54-55
sheep, 36, 53-54
signs of, 49
swine, 36,39,55-57
xerophthalmia, 16, 17, 33, 51, 58, 61,
62,63-64
food processing and storage, effects of,717

functions, 28-38
bone development, 36
epithelium, maintenance of normal,
34-35
immunological response and disease,3 7-3 8overview of, 28-31
reproduction, 30, 36
vision, 31-34
history, 16-18
introduction to, 15
metabolism
absorption, 23-24
digestion, 22
excretion, 27
storage, 27-28, 39, 67
transport, 24-27
nuclear retinoic acid receptors (RAR),
26-27,30-31
receptor, 103-104
requirements, 39-43
factors influencing, 41-42
table of species, 40
sources, natural, 43-48
stability of, 46-48, 69-71, 715, 716
supplementation, 69-75, 728-732, 734,
736
toxicity, 78-82
vitamin E and, 170
vitamin K and, 241
Vitamin A2, 18-19
Vitamin allowances, optimum, 723-727
Vitamin B,. See Thiamin (vitamin B,)
Vitamin B,. See Riboflavin (vitamin B,)
Vitamin B,. See Niacin (vitamin B,)
Vitamin B,, 385-417
analytical procedures, 388-389
antagonists, 387-388, 722
bioavailability, 719
carnitine and, 643
chemical structure and properties,
3 86-3 87
deficiency, 399-409
assessment of status, 408-409
cats, 404
dogs, 403-404
fish, 404
foxes and mink, 404
horses, 403
humans, 405-408
laboratory animals, 404-405
poultry, 401-403
primates, nonhuman, 405

789

INDEX
deficiency (cont.)
rabbits, 405
ruminants, 399-409
swine, 400-401
functions, 391-394
history, 385-386
introduction to, 385
metabolism
digestion, absorption, and transport,
389-390excretion, 390-391
storage, 390
niacin synthesis and, 358
requirements, 394-396
dietary protein and, 396
table of species, 395
riboflavin and, 320, 339
sources, natural, 397-3951
stability, 397-398, 410-411
supplementation, 409-412, 729-733,
735,737
toxicity, 412-413
Vitamin B,,, 6, 523-563
analytical procedures, 528-529
antagonists, 526, 528
chemical structure and properties,
526-528
choline and, 574, 581
deficiency, 540-554
assessment of status, 552-554
bats, 546
cats, 546-547
dogs, 547
fish, 547
horses, 546
humans, 548-552
laboratory animals, 547-548
mink, 548
poultry, 545-546
primates, nonhuman, 548
rabbits, 548
ruminants, 541-543
swine, 543-545
folacin and, 486-487, 489, 508
food processing and storage, effects of,717
functions, 532-535
history, 523-525, 672
introduction to, 523
metabolism
absorption, 529-531, 551
digestion, 529

790

excretion, 532
tissue distribution and storage,
531-532,551
transport, 530, 551
in plant tissues, 7
requirements, 535-538
table of species, 536
sources, natural, 538-540
stability, 526, 528
supplementation, 555458,738
toxicity, 558-559
vitamin C and, 608
vitamin E and, 170
Vitamin B,, (orotic acid), 670
Vitamin B,, (pangamic acid), 670
Vitamin B,, (laetrile), 670
Vitamin C (ascorbic acid), 597-640
analytical procedures, 601-602
antagonists, 601
bioavailability, 719
carnitine and, 643
chemical structure and properties,
600-601
deficiency, 613-627
assessment of status, 626-627
cats, 621
dogs, 620-621
fish, 621-623
folacin and, 51 1
horses, 620
humans, 624-626
laboratory animals, 623
poultry, 617-620
primates, nonhuman, 623
rabbits, 624
ruminants, 613-615
swine, 615-617
food processing and storage, effects of,717
functions, 604-609
antioxidant, 605-607
collagen synthesis, 604-605
immune system, 606
list of, 607-609
history, 597-600
introduction to, 597
metabolism, 602-604
requirements, 609-611
sources, natural, 611
stability, 601, 612-613, 630-631
supplementation, 627-633, 738
cattle, 628

INDEX
fish, 631
humans, 632-633
poultry, 629
stress and, 627-628
swine, 629
toxicity, 633-634
vitamin E and, 170
Vitamin D, 9 1-153
analytical procedures, 95-97
antagonists, 95
chemical structure and properties,
94-95
deficiency, 117-133
assessment of status, 133
cats, 128
dogs, 127-128
effects, general, 117-119
fish, 128-129
foxes and mink, 129
horses, 127
humans, 130-133
laboratory animals, 129
poultry, 124-127
primates, nonhuman, 129-130
ruminants, 119-123
swine, 123-124
functions, 103-111
calcium absorption by ruminants,
107-108
calcium and phosphorus homeostasis,104
bone effects, 106-107
intestinal effects, 104-106
kidney effects, 107
chick embryonic development,
109-110phosphorus absorption by ruminants,
108-109
history, 91-94
introduction to, 91
metabolism, 97-102
absorption, 97
conversion from precursors, 97-98
conversion to active forms, 98-101
excretion, 102
storage and placental transfer,101-102transport, 101
phytate and, 664
receptor, 103-104
requirements, 111-115
calcium-phosphorus ratio, 113

sunlight exposure, 111-112
table of species, 1 11
signal transduction, 104
sources, natural, 115-1 17
stability, 136-137
supplementation, 134-140, 728-734,
736
toxicity, 140-146
plants, 142-145
safe upper dietary levels, 145-146
vitamin E and, 170
Vitamin D,, 111-113, 618
Vitamin D-binding protein (DBP), 101, 103
Vitamin E, 155-225
analytical procedures, 160-161
chemical structure and properties,
157-160
deficiency, 178-207
assessment of status, 206-207
cats, 200-201
dogs, 198-200
exotic animals, 203
fatty acids and, 708
fish, 201
foxes and mink, 201
horses, 197-198
humans, 203-206
laboratory animals, 202
overview, 178-181
poultry, 193-197
chickens, 194-196
turkeys, 194-1 96
primates, nonhumans, 202
rabbits, 202-203
ruminants, 181-188
swine, 188-193
food processing and storage, effects of,717
forms of, 157-160
functions, 164-170
antioxidant, 164-166
blood clotting, 166-167
disease resistance, 167-168
electron transport and DNA, 168
membrane structure and prostaglandin
synthesis, 166protection from toxic substances, 169
selenium, relationship with, 169
history, 155-157
introduction to, 155
metabolism
absorption and transport, 161-162

791

INDEX
Vitamin E, metabolism (con.?.)placental and mammary transfer,162-163
storage and secretion, 163-164
requirements, 169-174
PUFAs and, 170-172
table of species, 171
sources, natural, 174-178
stability, 159, 160, 177-178, 208, 716
supplementation, 207-216, 728-734,
736
forms available, 207-209
needs for, 213-214
ruminants, 209-213
toxicity, 216
vitamin A and, 80
Vitamin H3 (gerovital), 671
Vitamin K
analytical procedures, 231-232
antagonists, 230-23 1, 255, 722
chemical structure and properties,
229-230
deficiency
antibiotics and, 246, 248, 252, 255
assessment of status, 253-254
cats, 250
dogs, 250
fish, 250
foxes, 251
horses, 249
humans, 251-253
laboratory animals, 251
poultry, 247-249
primates, nonhuman, 251
rabbits, 251
ruminants, 244-245
swine, 245-246
food processing and storage, effects of,717
functions, 2 35-2 39
blood clotting, 235-236
bone development, 238-239
carboxylation reactions, 236-238
history, 227-229
introduction to, 227
metabolism
absorption and transfer, 232-234
excretion, 234-235
storage, 234
requirements, 239-242
table of species, 241
sources, natural, 242-243

792

stability, 243, 256-257
supplementation, 254-258, 729-730,
732-734, 736
toxicity, 258-259
Vitamin-like substances, 659-674
coenzyme Q, 669-670
flavonoids (polyphenols), 668-669
glucose tolerance factor, 671
history of, 659, 671-672
lipoic acid (thioctic acid), 669
myo-inositol, 660-666
p-aminobenzoic acid (PABA), 667668
pyrroloquinoline quinone (PQQ),
666-667
vitamin B13 (orotic acid), 670
vitamin B,, (pangamic acid), 670
vitamin B,, (laetrile), 670
vitamin H, (gerovital), 671
vitamin U (cabagin), 671
Vitamins. See also specific vitamins
classification of, 4-5
definition of, 3dietary intake, factors influencing
agronomic effects, 714
harvesting conditions, 714
least-cost feed formulation, 720
processing and storage, 714-71 7
reduced feed intake, 717-718
vitamin bioavailability, 719-720
vitamin variability, 718-719
history of, 7-13
nomenclature, 5-6
occurrence, 7requirements, 6-7
Vitamin U (cabagin), 671
Von Bunge, C., 11-12
Von Euler, 16-17
Von Liebig, Justus, 10
Warfarin, 231, 237, 238, 245, 250,
255
Water-soluble vitamins, 4-5, 6
Wernicke’s syndrome, 299
White liver disease, ovine, 543
White muscle disease, 181-1 86
Wine and flavonoids, 669
Xanthine oxidase, 3 17, 3 19
X disease (hyperkeratosis), 53
Xerophthalmia, 12, 16-17, 19, 33, 51,
58, 61-64

INDEX
Yeast
niacin content, 360, 361
pantothenic acid, 420,429
thiamin content, 278, 279
Zenker’s degeneration, 178
Zincantioxidant, 606

folacin digestion and, 485
phytate and, 664
vitamin A and, 48-49, 66-67
vitamin destruction by, 715
vitamin E and, 177
Zoo wildlife species, vitamin E deficiency,
203793