NOAA Technical Memorandum NMFS-F /NEC- 25 

This TM series is used for documentation and timely communication of 

preliminary results. interim reports. or special purpose information; and 

has not received complete formal review. editorial control. or detailed 

editing. 

Histological Techniques 

for 

Marine Bivalve Mollusks 

Dorothy W. Howard and Cecelia S. Smith 

Oxford Lab., National Marine Fisheries Serv., Oxford, MD 21654 

U.S. DEPARTMENT OF COMMERCE 

Malcolm Baldridge, Secretary 

National Oceanic and Atmospheric Administration 

John V. Byrne, Administrator 

National Marine Fisheries Service 

William G. Gordon, Assistant Administrator for Fisheries 

Northeast Fisheries Center 

Woods Hole, Massachusetts 

June 1983 
Oxford La bora lory 

ii 
This manual is dedicated to Dr. Aaron Rosenfield and to the staff of the Oxford 

Laboratory, past and present. 

iii 
FOREWORD 

During the mid-1950s, massive and widespread oyster mortalities, particularly in the 

Delaware and Chesapeake Bay areas, served as a stimulus to establish a Federal oyster 

mortality study program. Consequently, Congress appropriated funds to the then 

Bureau of Commercial Fisheries (BCF) within tbe U.S. Department of the Interior. 

These appropriations enabled Federal, state, and academic institutions and laboratories 

to conduct research and management programs. These programs, either by contract, 

interagency agreement, or direct grant, would be designed to recognize and understand 

symptoms of disease, disease processes, their cause(s), and, furthermore, would help in 

the development of strategies that would prevent or control shellfish mortalities and the 

spread of disease agents. At that time, the Annapolis Laboratory of the BCF, under Mr. 

James B. Engle, was housed in an apartment building in Annapolis, Maryland, and the 

disease study was staffed by one biologist, Mr. Richard Burton, and two temporary 

summer technicians. I was one of those technicians, and started my career with strictly 

on-the-job training in histological and, later, pathological methods. In 1960 the Laboratory 

was moved to Oxford , Maryland, where a larger disease research and ecology program 

was initiated. 

Little was known at that time about oyster diseases, or even how to go about studying 

them. Standard vertebrate methods were used in the beginning, but it soon became 

obvious that many techniques needed modification and refinement. Because of the 

unique nature of invertebrate tissues, and their parasites, protocols were developed by 

Dr. Melbourne R. Carriker for the study of oyster disease in individual specimens and in 

populations. 

Since that time, over SOO,OOO sections of oysters and, more recently, other mollusks, 

crustaceans, and fishes have been processed. New and revised histological protocols 

and methodologies have been developed by the cooperative effort of several Laboratory 

staff and from information published in the scientific literature and in pertinent manuals. 

Handwritten laboratory notes and diaries, etc., have been collected over the past 20 

years and developed into this manual by the authors. The result is a working manual 

designed and written by technicians primarily for technicians, using very practical 

approaches and language which are comprehensible for someone just beginning, yet 

detailed and accurate enough to produce professional results in the hands of the 

experienced scientists should they elect to do the work themselves. 

I am proud to have been able to work with the authors. Over the past several years, 

their efforts and those of the senior staff of molluscan, crustacean, and fish pathologists 

resulted in the development of this manual. I feel that the authors' diligent efforts, both 

in the production of excellent histologic material and of this manual, will result in 

substantially enhancing the field of invertebrate pathology worldwide. 

c. AUSTIN FARLEY 

v 
ACKNOWLEDGEMENTS 

This manual is based upon the outlines, techniques, and guidelines of Virginia 

Liddell, Lucretia Krantz, and Helena Jones and we are grateful for their contributions. 

We particularly acknowledge our resident molluscan pathologists, Austin Farley and 

Fred Kern, for their support and professional assistance in preparation of this manual, 

and our Laboratory Director, Dr. Aaron Rosenfield, for his guidance and encouragement. 

We are grateful to Fred Kern for providing us with many of the photographs shown in 

this manual, as well as to Sara Otto for the use of her collection of specimens. 

We particularly thank Dr. Phyllis Johnson and Martin Newman for their help and the 

supervision given us while working out special stains and techniques. 

We appreciate the excellent anatomical illustrations created by Alice Jane Lippson. 

Our special thanks to Helen Lang and Susie Hines for their assistance in the lengthy 

literature searches, and to Muriel McNelis for her patience and understanding during 

the stages of preliminary typing. 

We are also grateful to Dr. Robert Lippson and Haskell Tubiash for their editorial 

assistance. 

And we sincerely acknowledge the late Betty Sanders of the National Cancer Institute 

for her encouragement and inspiration in our field of study. 

Last but certainly not least, we must give credit to the present and former staff 

members of the Oxford Laboratory. They provided us with ideas, techniques, approaches, 

and counsel. Their numbers are too numerous to mention but we acknowledge their 

contributions with gratitude. 

SPECIAL ACKNOWLEDGEMENT 

Due to the policy of this publication, it was not possible to recognize Mrs. Jane Swann 

as editor of this manual. We strongly feel that her talents and her editorial expertise 

should not be overlooked. We would like to express our appreciation to her through the 

following acknowledgement. 

A very special thanks is given to our editor, Jane Swann, for the endless hours of 

dedicated work on this manual. She transformed collections of rough drafts, procedures, 

and notes into a viable publication. During the continual changes and reorganizing of 

this manual, Jane's patience and perseverance went well beyond her normal duties. We 

the authors are indebted to her for her diligent editing and consistent review over a 

period of several years. Working with her was a pleasure and we are sincerely grateful to 

her. 
CONTENTS 

INTRODUCTION .. . . . . . . . . . . . . ........ . ... .... . . 

NOTES ON SAFETY ... ............... . . . .. . . . . . 

CHAPTER I. HANDLING OF LIVE ANIMALS 

A. Shipping Instructions for Shellfish 

B. Necropsy and Examination of Shellfish Specimens 

1. Processing Steps .. ... . . . 

2. Figures and Explanations ... .... . 

C. Processing of Specimens 

l. Oysters (Crassostrea and Ostrea spp.) 

2. Clams 

3. Mussels (Mytilus spp.) ...... . . 

4. Scallops (Argopecten and Placopecten spp.) 

5. Bivalve Larvae 

D. Notes . . . . ... . . . 

E. References . 

CHAPTER II. FROZEN TISSUE TECHNIQUE 

A. Description of Technique ..... .. . . . 

B. Reference ..... ... . .. .. . . . . ... .. . . 

CHAPTER III. FIXATION 

A. Formulas for Fixatives and Methods 

B. Special Note on Sea Water 

C. Notes on Fixation 

D. Problems and Resolutions 

E. Notes 

F. References . 

CHAPTER IV. EMBEDDING 

A. General Procedures 

B. Embedding Schedules 

C. Problems and Resolutions 

D. Notes. 

E. References . 

CHAPTER V. SECTIONING 

A. Techniques .. ...... .... .. . 

B. Preparation of Tissue Sections 

1. Blocks .... . ...... .. .... ... . 

2. Microtome . . . . . . . . . ...... .. . . 

3. Knives 

4. Sectioning . 

C. Problems and Resolutions 

D. Notes . 

E. References . 

CHAPTER VI. STAINING 

A. Methods and Use 

1. Standard Format for Paraffin-Embedded Sections .. ... . . . . 

2. Pre-Staining Techniques . . ......... . . 

3. Staining Methods ... . . .. . 

B. Problems and Resolutions 

C. Notes . . . . . . . . . . .. . .... . . 

D. References .... ... . . 

CHAPTER VII. COVERSLIPPING 

A. Tools and Techniques . 

B. Problems and Resolutions 

C. Notes 

D. Reference 

ix 

1 

.. .. .. 3 

5 

5 

5 

5 

5 

7 

7 

20 

20 

.... 21 

· .. 21 

. .... 21 

21 

· 33 

· . . . . . . . 33 

· .. 33 

35 

35 

37 

.. ... ... .. . 37 

· 37 

· . ....... . 37 

. .. .. 37 

· 39 

39 

39 

.39 

39 

39 

... .. . .. . . . . .. 42 

42 

42 

.. 42 

. ... 42 

· .. 42 

. . . .... 42 

· 43 

· . . . . . . . 43 

. . 43 

.. .... . 50 

. . . . . . . . 50 

. . . . . . . . . . . . . . . . . 53 

. ... .. 54 

55 

81 

81 

· 81 

.. .... . . ... 83 

· . 83 

83 

83 

83 
APPENDIX 

A. Special Techniques ............... . 

1. Culture of Perkinsus marinus ...... . . 

2. Decalcification 

3. Transfer of Tissue from Broken Slides . 

4. Transfer of Tissue from Paraffin to Plastic 

5. Record Keeping 

6. Filing and Slide Storage 

B. Notes 

C. References 

GENERAL REFERENCES 

INDEX 

x 

86 

.. 86 

86 

. . . . . . . . . . . 86 

87 

87 

............. 91 

. . . . . . . . . . . . ... . .. . ... . ... . . . . . .. . .. 91 

91 

91 

94 

95 
Histological Techniques 

For Marine Bivalve Mollusks 

Dorothy W. Howard and Cecelia S. Smith 

INTRODUCTION 

Investigators at the Oxford Laboratory diagnose and study finfishes, crustaceans, 

mollusks, and a variety of other marine and estuarine invertebrates. However, this 

manual is intended to serve as a guide for histological processing of shellfish, principally 

bivalve mollusks. Basically, the techniques included are applicable for histopathological 

preparation of all marine animals, recognizing however that initial necropsy is unique to 

each species. Several of the procedures described are adaptations developed by the 

Oxford Laboratory staff. They represent techniques usually based on principles established 

for histopathologic study of mammalian and other vertebrate tissues, but modified for 

marine and aquatic invertebrates. Although the manual attempts to provide adequate 

information on techniques, it is also intended to serve as a useful reference source to 

those interested in the pathology of marine animals. General references listed in the 

back of the manual will provide histological information on species not included in the 

text. 

The first chapter on handling live animals individually describes the procedures 

employed for oysters, clams, mussels, and scallops, from receipt to the point of fixation. 

Subsequent chapters provide information on the specific phases of histology with 

adaptions used by the Oxford Laboratory. The chapter on staining methods includes 

general tissue stains, special stains for specific tissue components, stains for frozen 

sections and for fresh and fixed blood smears. Variations in the duplicate stain protocols 

are given to correlate the pH of the stains with the fixatives and tissues. Most of the stains 

have been successfully used on tissues of mollusks, finfishes, and crustaceans. Special 

techniques and data sheets have been compiled in appendix form . 
NOTES ON SAFETY 

Safety cannot be overemphasized in a histology laboratory since volatile solvents and 

hazardous chemicals are used. Each employee should be made aware of the possible 

hazards involved. Laboratory supervisors must provide adequate instructions on the 

safe use of the chemicals involved, as well as storage and disposal, even though most 

manufacturers provide warning labels. 

The first supervisory priority is to be sure your working facility is adequately equipped 

for safe operation. The working environment must be free of toxic vapors. Fume hoods 

are essential, and are utilized from the initial steps of preparing fixatives, solvents, and 

noxious reagents to the final steps of coverslipping and cleaning the dried, stained slides. 

Protective stainless steel mesh gloves' are required for opening oysters and hard 

clams. Plastic gloves and disposable aprons should also be available and used as 

necessary. Pipette bulbs and disposable pipettes are used for contaminating and dangerous 

chemicals. Safety goggles and eyewashing facilities must be available. The laboratory 

should be equipped with a safety shower for emergencies. 

Safe operation of a microtome is essential. Locking levers on the hand wheel of the 

microtome should be kept in working order. When using the longer microtome knife, a 

piece of rubber hose is placed on the extended end as a safety guard. 

Gas line hoses to bunsen burners and the laboratory's fire extinguishers should be 

routinely checked, and the local fire marshal and Laboratory safety officer should 

perform periodic inspections. 

Extreme care should be used with highly explosive anhydrous picric acid salts. Dried 

ammoniacal silver salt solutions are also extremely explosive. These chemicals are used 

in special stain techniques. 

Technicians should become familiar with all aspects of the chemicals involved in 

histological procedures. Technicians who have not had the proper instruction in handling 

these dangerous chemicals risk serious injury to themselves and their co-workers. 

'Whiting & Davis Co., Plainville, Mass. 

3 
CHAPTER I. HANDLING OF LIVE ANIMALS 

A. SHIPPING INSTRUCTIONS FOR SHELLFISH 

Live animals must be shipped and received in good physiological condition to ensure optimal tissue analysis. If specimens 

cannot be shipped live in good condition, see Chapter III for 

instructions on using specific fixatives prior to shipment. The 

following instructions provide for shipment of live shellfish to 

ensure viable samples on arrival : 

• Pack live shellfish (e.g. oysters or clams) (sample of 50) in 

ambient salinity , seawater-soaked burlap, with cold packs, in a 

well-insulated, well-constructed waterproof shipping container. 

Sbipment sbould be made early in tbe week ~ Monday or 

Tuesday) to avoid arrival during tbe weekend period, with 

possible loss of samples because of improper storage conditions. 

• On outside of container, clearly label "LIVE SPECIMENS, 

REFRIGERATE BUT DO NOT FREEZE." For air freight, 

indicate "HOLD AT AIRPORT ON ARRIVAL AND CALL 

LABORATORY" (destination). 

• Wire or phone, immediately after shipment has been made , 

indicating where and when sent, estimated time of arrival, 

name of originating and , if possible, anticipated delivering 

carrier, flight number and, most importantly, the waybill number. 

• Ship by air freight (prepaid) directly to destination. 

• Provide background information with each sample, such as: 

name of requestor; to whom report should be sent, including 

address and phone number; pertinent collection data, such as 

temperature, salinity, pH, species, origin, history, sampling 

location , number of specimens in sample, and date collected. 

B. NECROPSY AND EXAMINATION OF 

SHELLFISH SPECIMENS 

Shellfish for histological processing should be sampled in 

the field by methods causing minimal, if any, damage to the 

specimens. They should be kept refrigerated , particularly in 

hot weather, until processed . Fixation should be as soon as 

possible after collection to reduce physiological or postmortem 

changes. 

1. ProceSSing Steps 

• Record field data and history of sample 

• Assign code 

• Scrub animals to remove fouling organisms 

• Measure shells 

Bill to hinge for oysters, mussels 

Across shell, side to side, for clams, scallops 

• Open shellfish 

Pry hinge open and cut adductor muscle to avoid damaging 

tissues 

• Record the following: 

Gaping bivalves that do not close 

Weak muscle or loss of muscle adhesion 

Lack of mantle reaction 

Any unsual odor 

Stabbing specimen when opening 

5 

Incision (cut) 

Laceration (tear). (Reaction to stabbing can cause acute 

inflammatory infiltration of hemocytes to localized 

areas.) 

• Record macroscopic information. (Shell abnormalities, 

parasites, and physiologically related conditions included 

represent only a few of the multitude associated with marine 

shellfish. The purpose of these examples is to show a spectrum 

of conditions seen during an examination of specimens. Any 

observations noted during the macroscopic examination will 

aid the investigators in their analyses.) 

Shell abnormalities. (Experience has shown that certain 

conditions are predictable from samples of a certain 

geographic area. Each condition and degree of prevalence 

has an effect on the animal and how the animal has adjusted 

to it.) 

Mantle recession (Fig. 1a) 

Recovery (Fig. Ib) 

Cliona sp. (boring sponge) penetrations, including estimated 

degree of infestation (Figs. 2a, b) 

Polydora sp. (mud worm) tunnels, including estimated 

degree of infestation (Figs. 3a, b, c) 

"Maladie du pied" (Fig. 4) 

Shell blisters (Fig. 5) 

Shell pustules (Fig. 6) 

Calcareous malformations (Fig. 7) 

Drill cases and/ or drill holes (Fig. 8) 

Obvious parasites 

Pinnotheres sp. (gill area) (Fig. 9) 

Mytilicola sp. (gut area) (Fig. to) 

Sphenophrya sp. (gill area) (Fig. 11) 

Bucephalus sp. 

Physiologically related conditions 

Fat (Fig. 12a) 

Medium (Fig. 12b) 

Watery (Fig. 12c) 

Pale digestive diverticulum (Fig. 13a); normal digestive 

diverticulum (Fig. 13b) 

Swollen tissue (edema); example, swollen heart (Fig. 14) 

Color 

Pigmentation (melanin) 

Green deposits (diapedesis) 

Other colorations 

Other gross abnormalities 

Watery cysts (Fig. 15) 

Abscesses and tumors 

Pearls (soft or hard) 

2. Figures and Explanations 

a. Shell Abnormalities 

1) Mantle Recession (Fig. 1 a) 

Heavy fouling of the inside shell margin indicates prolonged 

mantle recession. This condition usually indicates a sick oyster 

(Farley , 1968). 
2) Recovery (Fig. 1 b) 

The oyster has recovered from previous recessive growth 

(mantle recession), overlaying the fouled inner shell margin 

with new periostracum. "The periostracum is a film of organic 

material (scleroprotein called conchiolin), secreted by the cells 

located near the very edge of the mantle . ... the conchiolin 

appears as amorphous, viscous and transparent material which 

hardens shortly after being deposited" (Galtsoff, 1964). 

3) CHona sp. (boring sponge) penetrations (Figs. 2a, b) 

According to Galtsoff (1964), "Small round holes on the 

surface of mollusk shells indicate the presence of the most 

common animal associated with the oyster, the boring sponge." 

The sponge reaches the oyster tissue itself only in cases of old, 

heavy infestation. Usually, the holes made by the sponge are 

rapidly covered by a deposition of conchiolin. "However, should 

the deposition of shell material be delayed by adverse conditions, the sponge makes direct contact with the mantle and 

produces lysis of the epithelium and underlying connective 

tissue. Dark pigmented pustules fonn exactly opposite the holes 

in the shell" (Galtsoff, 1964). 

4) Polydora sp. (mud worm) tunnels (Figs. 3a, b, c) 

"P. websteri invades the shell cavity of the oyster, settles on 

the inner surface at a right angle to the edge , and builds a 

U-shaped mud tube with both orifices external. The structure 

is soon covered by a layer of conchiolin deposited by oyster 

and becomes a semitransparent blister" (Galtsoff, 1964). 

5) "Maladie du pied" (foot disease) (Fig. 4) 

"The disease affects the area of the attachment of the adductor muscle, primarily on the lower concave (left) valve, and in 

certain cases the upper, flat valve . .. . In advanced cases the 

muscle becomes detached from the valve and forms irregular 

cysts of horny and slightly elastic material. Later on when 

the cyst extends beyond the area of the muscle attachment, 

the cyst walls become covered with calcareous shell deposit" 

(Galtsoff, 1964). 

6) Shell blisters (Fig. 5) 

Blisters frequently are found on the inside of the shells near 

the adductor muscle. "Chambers found in C. virginica consist 

of irregular cavities containing mud or sea water. Such 

fonnations are called blisters. Blisters can be artificially induced 

by inserting a foreign object between the mantle and the shell. 

They are also caused by the invasion of shell cavity by Polydora 

or by perforations of the shell by boring sponges and clams" 

(Galtsoff, 1964). 

7) Shell pustules (Fig. 6) 

Related to Minchiniasis (MSX). "Oysters with raised yellowbrown conchiolin deposits on the nacreous surface of the 

6 

shell . .. These deposits contained a creamy yellow fluid which 

consisted of moribund M. nelsoni plasmodia, hemocytes, and 

cellular debris .... when present it was a reliable gross indication of the disease" (Farley, 1968). Farley termed a stage of the 

MSX disease "remission," when the shell pustules appeared as 

described above. 

8) Calcareous malformations (Fig. 7) 

These abnormalities are pathological. "They are associated 

with the disturbance of calcium metabolism which manifests 

itself in an overcalcification of selected parts of the organism . 

. They fall into two classes: 1) cysts and calcified cups; and 

2) calcified tumors and pearl-like structures attached to the 

valve or formed inside the soft tissues .. .. It is reasonable to 

assume that the described malformations represent a phenomenon analogous to the over-calcifications of various selected 

part of mammals described by Selye (1962) under the general 

term calciphylaxis" (Galstoff, 1969). 

9) Drill cases and/or drill holes (Fig. 8) 

The presence of oyster drill egg cases are recorded when 

attached to oyster shells in a sample since oyster drills are the 

most persistent predator of the oyster. The drill cases are 

tough, greenish leathery capsules in which oyster drill eggs are 

deposited . Small symmetrical holes in molluscan shells can be 

attributed to the action of oyster drills or other gastropods. 

b. Obvious parasites 

1) Pinnotheres sp. (gill area) (Fig. 9) 

These small oyster or pea crabs invade the mantle cavity of 

oysters and other bivalve mollusks. The female crab settles 

chiefly on the surface of the gills, and grows with the growth of 

the host. " ... the female crabs which have settled on the 

oyster erode its gills and impair their function. . .. Rapid 

regeneration of the damaged gills probably saves many oysters 

from death, but interference with the nonnal gill functions 

causes a relatively poor condition in many infested oysters" 

(Galtsoff,1964). 

2) Mytilicola sp. (gut area) (Fig. 10) 

This "parasitic copepod is found in the intestinal tract of 

bivalves and is easily recognized by its red color and relatively 

large size which makes it visible to the naked eye" (Galtsoff, 

1964). 

3) Sphenophrya sp. (gill area) (Fig. 11) 

These are ciliates which attack the gills of the oyster. "The 

disease is caused by a microparasite which has been tentatively 

placed in the genus Sphenophrya. The organism is a ciliate 

found in large cysts on the gill surface of diseased oysters" 

(Anonymous, 1965). 
4) Bucephalus sp. 

This trematode thrives in brackish water oysters and is 

inhibited by an increased salinity. "In cases of heavy infestation, 

the gonads and digestive diverticula are almost completely 

replaced by cercariae and by the long germ tubes of the 

sporocysts ... " (Galtsoff, 1964). 

c. Physiologically related conditions 

1) Fat, medium, or watery (Figs. 12a, b, c) 

During the initial necropsy, each shellfish is rated for its 

physiological condition as fat (Fig. 12a), medium (Fig. 12b), or 

watery (Fig. 12c). This condition reflects the oyster's general 

health and is affected by its environmental surroundings, stress, 

and seasonal and sexual changes. Fat oysters usually fiJI their 

shell; often they are sexually ripe and exhibit a creamy tan 

color. Medium oysters generally are smaller in their shell, 

flatter, and less rigid than fat oysters; there is more fluid inside 

the shell and their color is basically tan. Watery oysters are 

obviously bloated with water; they exhibit a grayish-tan color 

and the animal is slightly translucent. 

2) Pale digestive diverticulum (Figs. 13a, b) 

Oysters from the same location usually show uniform-colored 

digestive diverticula (reddish-brown, greenish-brown, yellowishtan , or blackish-brown) since they have been exposed to the 

same environmental food column. Occasionally, a pale digestive 

diverticulum occurs, and is so recorded. Many times the pale 

color can be associated with a health problem, diagnosed only 

after microscopic examination. 

3) Swollen tissue (edema), swollen heart ("cardiac vibriosis") 

(Fig. 14) 

During a survey for parasite distribution in Chesapeake Bay 

oysters, Tubiash et aJ. (1971) noted sporadic cases of greatly 

enlarged and edematous pericardia. "Examination of aseptically 

aspirated pericardial fluid showed heavy concentrations of 

gram-negative motile rods which proved morphologically and 

culturally compatible with Vibrio anguillarum, an organism 

implicated in diseases of fishes and larval bivalve mollusks. 

Except for pericardial enlargement, the animals appeared to 

be grossly and histologically normal" (Tubiash et al. , 1971). 

4) Color 

"Pigmentation may be considered as a certain phase of 

excretion. Green oysters of Long Island Sound develop dark 

green pigment as the result of absorption and storage of copper 

by the phagocytes ... " (Galtsoff, 1964). 

Other colorations sometimes. associated with mollusks are 

the result of pigments from the microscopic plants (algae) that 

mollusks consume. Some pink pigmentation of stored shellfish 

may be the result of the growth of so-called pink yeasts or 

pigmented bacteria (McCormack , 1956; Boon, 1972; Paparella, 

1973). 

7 

d. Gro .. abnormalHi .. 

1) Watery cysts (in the visceral mass, palp, or mantle) 

(Fig. 15) 

"The cysts, seen in the live oyster, are large conspicuous 

bubble-like cavities generally distended by internal pressure of 

accumulated fluid. Sections show these cysts to contain a 

central granular material usually in concentric layers . ... The 

cyst walls are formed of leucocytic cells which form themselves 

into an epithelium similar to that of the external epithelium of 

the mantle" (Mackin, 1962). 

2) Abscesses and tumors 

"Reports of tumors in bivalves are relatively common in 

comparison to other invertebrate groups. . .. a number of 

so-called tumors have been shown subsequently to be 

hyperplastic reaction~ to injury rather than true neoplasias" 

(Sparks, 1972). 

"Systematic surveys of molluscan populations offer a good 

method of obtaining extensive numbers of naturally occurring 

lesions in an attempt to classify molluscan tumors and specify 

criteria for distinguishing among true neoplasia, hyperplasia, 

and injury responses in these invertebrates" (Pauley, 1969). 

3) Pearls 

"In the case of pearl formation, Boutan (1923) has shown 

that calcareous deposits are formed by amoeboid cells which 

crawl through the mantle epithelium, while the latter secretes 

the concentric layers of the organic matrix (conchiolin)" 

(Galtsoff, 1964). 

C. PROCESSING OF SPECIMENS 

A random sample (preferably 30-50 specimens) of the 

population is collected for histological processing and study. A 

code is assigned for further in-house reference to the sample 

and for permanent filing. All pertinent data furnished with the 

sample should be included with the new code, along with the 

name of the requestor and the date(s) of collection (see 

Appendix , Molluscan Pathology Report). 

The initial steps employed in processing molluscan species 

are provided below. The discussion on processing oysters is 

presented completely in a stepwise manner. 

A separate section is provided for special opening and 

processing techniques for clams, mussels, and scallops. 

1. Oysters (Crassosfrea and Ostrea spp.) 

The oysters are scrubbed to remove sediment and fouling 

organisms, measured from the hinge to the bill (Fig. 16), and 

their size, in metric units, is recorded in numerical order. They 

are opened by inserting an oyster knife between the valves 

(Fig. 17) at the hinge arch and twisting to separate the hinge 

(Fig. 18). The adductor muscle is severed as close to the right 

valve as possible (Fig. 19), taking care not to injure internal 

tissue. The opened oysters are then examined (Figs. 20, 21) for 
color, condition (fat, medium, or watery), macroparasites, and 

abnormalities. The body is removed from the shell by severing 

the adductor muscle as close to the left valve as possible. The 

rectum is excised with scissors (Fig. 22) and placed in a correspondingly coded thioglycolate culture tube (Fig. 23) for determining the presence of Perkinsus (Labyrinthomyxa) marinus 

(=Dermocystidium marinum) (Ray, 1952). The animal is then 

washed in clean, filtered, ambient sea water to remove sand 

and placed on a clean No. 14 rubber stopper for cutting a 

cross-section with a sharp razor blade (Fig. 24). 

The first transverse cut is made where the palps and gills 

meet (see Figs. 25,28 for location). The second cut is made 1-2 

mm from the first cut, if frozen sections or tissue for electron 

microscopy study are desired. The third cut is made 4-5 mm 

beyond the second cut for histologic sectioning (on a very 

small specimen it may be just above the adductor muscle). 

Figure 29 illustrates oyster tissues which will be included in a 

4- to 5-mm transversely cut cross-section. A confusing point 

may be the crystalline style which is rarely seen during the 

processing of oyster tissues. According to Galtsoff (1964), 

8 

"The crystalline style is not a permanent structure. In oysters 

removed from the water and left in the air the style dissolves in 

a short time." 

The anterior portion containing the palps is placed in a 2-oz 

screw-top jar containing Milliporel-filtered sea water, correspondingly coded, and used for fresh squash examination (most 

parasites can be diagnosed at this point). The remaining 6- to 

7-mm-thick digestive diverticula section is placed in a Technicon2 

autopsy tray with its identifying code label (Fig. 26). The tray is 

immersed in a petri dish of sea water so the tissue will not dry 

out before fixing. The portion below the fourth cut from each 

animal is frozen for future reference, or discarded. The 1- to 

2-mm section of tissue to be used for frozen sectioning is 

placed in a labeled petri dish with the just-cut surface facing 

up, and is sectioned as described in the chapter on "Frozen 

Tissue Technique." When all the tissues in the sample are cut, 

the Technicon trays are placed in a jar of fixative (Fig. 27), 

coded , and processed as appropriate for the fixative. The 

scissors, razor blade. and stopper used for cutting should be 

rinsed between oysters to avoid carry-over contamination. 
Shellfish abnormalities. 

Fig. I. a, Mantle recession; b, recovery (specimen courtesy S. V. Otto). 

Fig. 2. a, Cliona; b, normal shell. 

9 
Fig 3. Sbellfisb abnormalities: 

a, Ligbt Polydora invasion; 

b, medium Polydora invasion; 

c, beavy Polydora invasion (specimen courtesy S.V. Otto). 

10 
Sbellfisb abnormalilies. 

Fig. 4. "Maladie du pied" (specimen courtesy S. V. Ouo). 

Fig. S. Sbell blislers wilb knife punclure marks. 

Fig. 6. Shell puslules (specimen courlesy C.A. Farley). 

Fig. 7. Calcareous abnormalities. 

11 
Fig. 8. Shellfisb abnormalilY: drill cases. 

12 
10 

Shellfish parasites. 

Fig. 9. Pinnotheres sp. 

Fig. 10. Mytilicola sp. lOX (photograph by C.A. Farley). 

Fig. II. Mulliple cysts on gill caused by Sphenophrya sp. 

(photograph by C.A. Farley). 

13 

11 
Fig. 12. Physiologically related conditions of shellfish: 

a, fat; b, medium; and c, watery. 

14 
Pbysiologically related conditions of sbellfisb. 

Fig. 13. a, Pale digestive diverticulum; and b, normal 

digestive diverticulum. 

Fig. 14. Swollen beart (specimen courtesy S.V. Otto). 

Fig. IS. Watery cysts (specimen courtesy S.V. Otto). 

15 
Processing of oyster, Crassostrea virginica . 

Fig. 16. Measuring oyster from bill to hinge. 

Fi!!. 17. Inserting knife in hinge. 

Fi!!. 18. Twisting knife to "pop" hinge. 

Fi!!. 19. Cutting adductor muscle. 

Fig. 20. Removing top shell for inspection. 

Fig. 2 \. Examining shell and oyster meat. 

16 
Processing of oyster. 

Fig. 22. Excising rectum of oyster. 

Fig. 23. Placing rectallissue in thioglycolate for culture. 

Fig. 24. CUlling cross·section. 

17 

Fi~. 25. 

Fi~. 26. 

Fig. 27. 

Oyster cut showing cross·section. 

Placin~ cross·section in tray with correspondin~ly coded 

label. 

Placing tray in fixative. 
He art -------------___ __ 

Perlcardlal a a c ---------__ 

An u a --------_ .... 

Adductor muacle 

(amooth) 

(8trlated) 

Cut edge of 

right mantle 

L eft man tI e --........ :-----"-'''''''';--0 

Inteatlne 

Digestive diverticula 

~...". _________ HI n g e 

Stomach 

1o!'ah'~....--''r--;'+1f1-------- E a 0 p " a g u a 

-~~,L-,~ ------- Lab I a I p a I p 8 

Style 

_____________ Go n ad 

Kidney 

G 1118 

Fig. 28. Gross anatomy of oyster. Tbe bold parallel lines sbow tbe 

location wbere tbe tissue for cross-section sbould be taken 

(illustration by AJ. l..ippson). 

18 
I n t est I n e-----!'off-iJ~!'__J4WJ 

'J---'6"IJ~~~lt-- Connective Ti ssue 

'~~ife4t4-Digest Ive dive rticu la 

Gonad---~~~~E 

Gill :>---------t;-Itl 

Fig. 29. Microscopic illustration of cross-section of oyster 

(illustration by 'A.J. Lippson). 

19 

Int es tine 

Kidney 

M ant Ie 
2. Clams 

Clams can be successfully opened with relative ease when 

the right equipment and techniques are employed . Different 

species and various sizes will dictate the appropriate technique. 

The following methods have been utilized at the Oxford 

Laboratory. 

The clams are held in a chilled environment until ready for 

processing. Chilling retards the metabolic rate and renders the 

clams sluggish and relaxed. 

a. Surf clams (Spisula solidissima) 

These large clams have an extensive muscular foot which 

prevents tight closure, and can be easily shucked . A long, 

sharp, flexible knife such as a fillet knife is used. Begin near the 

anterior adductor muscle, insert the knife behind the mantle 

next to the shell, and sever the muscle. Work the knife around 

the edge of the shell and cut the posterior adductor muscle . 

The same manipulation is used on the bottom shell. Immerse 

the clam in a bucket of ambient sea water and wash well, since 

these animals often contain much sand. The bucket of water 

should be changed frequently. 

b. Soft-shell clams (Mva arenaria) 

A disposable scalpel should be used when opening this species. 

The shells are fragile and brittle, so a light grip is recommended. Insert the blade along the shell just below the "snout" 

(siphon, Fig. 30), making sure the knife is positioned just below 

the mantle next to the shell, and cut around shell to hinge. 

Gently pry shell open and cut remaining tissue from the shell 

(Fig. 31). The longer it takes to insert the blade into the clam, 

the tighter the clam will close the shell (this is true for all 

bivalves). Therefore, insert the blade and cut the muscle as 

quickly as possible. Cut off the mantle and siphon (Fig. 32). 

Wash well and remove crystalline style if possible before making 

a cross-section (Fig. 33). 

c. Duck clams (Macoma balthica) 

These clams have paper-thin shells and are frequently 

damaged during collection. A quick and efficient method for 

opening these clams has been developed using a single-edged 

razor blade with a handle). The blade is extended from the 

holder and wrapped for support. The blade is both thin and 

flexible and can easily be maneuvered into and around the 

shell (Fig. 34), separating the tissue and shell without damage 

to the tissue (Fig. 35). Excise rectum if thioglycolate cultures 

are anticipated (Fig. 36). After making the cut for the crosssection (Fig. 37), remove the crystalline style and process as 

usual. 

d. Hard clams (Mercenaria and Arctica spp.) 

Clams with thick hard shells are the most challenging bivalves 

to open. Measure from side to side (Fig. 38). Using an oyster or 

clam knife, insert the tip (Fig. 39) into the clam just below the 

hinge which, interiorly, is just above the adductor muscle. A 

20 

hammer is necessary at times to penetrate the shell. Work the 

blade in for approximately one-half inch and twist. This will 

"pop" the hinge and occasionally the pressure may break the 

shell. However, the purpose is accomplished because the clam 

is exposed, making it easy to shuck without risking internal 

tissue damage. After the posterior adductor muscle is cut (Fig. 

40). carefully slide the knife along the edge of the shell to free 

the attached mantle (Fig. 41). Continue around the shell and 

cut the anterior adductor muscle. Use the same cutting action 

on the lower shell to free the clam from the shell (Fig. 42). 

Any variation of the above methods may be utilized as long 

as the clams are opened quickly, while avoiding any internal 

damage to the tissues. 

Once the clam has been carefully shucked and rinsed 

thoroughly (Fig. 43), cut a cross-section approximately 3- to 

5-mm thick (Fig. 45). Include as many major organs as possible. 

The anatomical drawing (Fig. 44) shows the different organs of 

a clam with a cutaway piece of mantle and gill section. For a 

successful cut, a very sharp knife blade is needed. Use a sawing 

motion, with very light pressure , to avoid displacing the soft 

tissues. 

Larger clams are difficult to cut uniformly. When the first 

cut is made, the clam retracts the muscular foot so tightly that 

it is hard to make a straight, thin, second cut. "Thinly" cut 

tissue sections are emphasized because the texture of the 

internal clam tissue is so soft that good fixative penetration is 

difficult to accomplish . 

The crystalline style in the clam is generally severed as the 

cross-section is taken. Remove the style when it is present, 

being careful not to damage the surrounding tissue. The 

crystalline style will harden during the embedment process, 

depending on the clam species, and may cause problems when 

sectioning (Macoma balthica is a good example). 

Since clams may have sand in the folds of tissue, care should 

be taken to remove it all. Wash well with filtered ambient sea 

water and inspect the interfolds of the mantle and gills prior to 

making the cross-section . If the sand has not been successfully 

removed before the animal is fixed and embedded, it will be 

incorporated with the tissue and will present an obvious problem 

when sectioning. Sand will leave saw-tooth nicks on the knifeedge , and is extremely destructive to the tissue as well, creating 

lines and torn ridges throughout the entire section. Broken 

chips of the shell and pearls can cause similar problems, so 

wash the tissue well before making the cross-section. Allow the 

live clams to remain in ambient sea water as long as possible so 

as to purge the sand and debris from their tissues. This will 

al.leviate many of the problems associated with sand in the gills 

and mantle tissue. 

3. Mussels (Mytilus spp.) 

Mussels from intertidal habitats often have many organisms 

within the byssus threads surrounding the shell ; therefore , all 

initial processing begins with a thorough scrubbing. This removes 

the possibility of transferring contaminating artifacts to the 

inside tissue . The length is then measured from bill to hinge 

and recorded . 

To open the mussel shell, insert a scalpel (Fig. 46) into the 

ventral byssal cavity and gently pry a small opening. Cut the 

mantle along the edge of the shell (Fig. 47). Take care to 

position the blade behind the mantle next to the shell, and cut 

in a clockwise motion around the bill to the hinge of the shell. 
Return the knife to the byssal cavity and, in the same fashion, 

cut to the top of the shell. The meat condition is recorded, 

along with any pertinent macroscopi c information . After the 

animal is washed in ambient sea water, the byssus is cut just 

into the byssal cavity tissue (Fig. 48). The mussel is then placed 

on a rubber stopper and a cross-section is taken (see Fig. 49), 

making sure all gill and mantle tissues are in place before the 

cut is made. Figure 50 shows the tissue orientation before the 

cross-section is placed in an autopsy tray (Fig. 51). The gross 

anatomy of the mussel is shown in Figure 52. The cross-section 

taken should include as many major organs as possible (see 

Fig. 53). 

4. Scallops (Argopecten and Placopecten spp.) 

Scallops are relatively delicate animals and should be handled 

carefully prior to processing. Their muscle system is weak and 

easily detaches from the shell. They tend to gape and dehydrate 

quickly when out of the water. They are usually collected by 

dredging methods, which leaves the scallops full of sand and 

grit. Optimum treatment of these animals would be to immerse 

them in holding tanks of ambient sea water cooled to iOo-15°C 

for 24 hr (0 purge the sand from the tissues. If this is not 

possible , they may be wrapped in soaked burlap or seaweed 

and kept cool and damp with ambient sea water until processing. 

Begin by measuring the shell (Fig. 54). Then open with a 

scalpel on either side of the hinge (Fig. 55). Once the hinge is 

"popped ," the scalpel is gently slipped into and around the 

shell and the muscle is cut (Fig. 56). 

After the scallop is removed from the shell, wash thoroughly , 

but carefully, since fine sand sticks between the gill filaments 

(Millipore-filtered sea water is used to wash animals). When 

cutting the cross-section of the scallop as shown in Figure 57, 

the gills and mantle can easily detach from the animal and 

become disoriented in the cross-section. Figure 58 illustrates 

the gross anatomy of a scallop. If care is taken during processing, 

the organs seen in Figure 59 can be included in the crosssection taken for examination. For convenience, the piece of 

mantle may be eliminated unless specifically desired by the 

investigator. since the edge of the mantle is lined with sensory 

eyes which can cause extensive problems when sectioning the 

tissue . 

5. Bivalve larvae 

Most larvae are fixed in Davidson's fixative (see Table 1). 

The acetic acid aids in disintegrating the shell. Since the 

dissolved carbonate neutralizes the fixative , it should be changed 

frequently to maintain its effectiveness. Occasional agitation 

will enhance fixation. After the animals have been fixed, they 

should be washed and placed in verse nate (disodium ethylenediaminetetraacetate) (Sanders, 1972) to complete decalcification of shells (see Appendix, A.2. ). The time required for 

complete decalcification will vary with the size and species 

of animal. The versenate solution should be changed daily and 

the sample checked at that time . Once the end point of decalcification is determined, wash the larvae well and place in 70% 

ETOH until embedment. Bivalve larvae are processed by hand 

rather than mechanically with the Autotechnicon' . The solutions 

are pi petted onto and off the larvae at designated times, using 

processing schedules compatible to the size of the animals (see 

Table 3). The vacuum oven is often used to speed the process 

of dehydration and infiltration of paraffin into the larvae. Very 

21 

small animals can be stained with a light solution of eosin after 

the 70% ETOH rinse to help the technician keep track of them 

when performing the embedding process 

Seed-size bivalves, approximately 0.3-0.4 mm, are generally 

shucked out with a scalpel. Pry the hinge with a firm but light 

pressure to expose the muscle which, when cut, will allow 

inspection of the specimen. 

Smaller shells tend to chip when opening; therefore, once 

shucked, the animals are completely immersed in ambient sea 

water to wash out any shell debris. Each animal is encircled 

by the correspondingly coded label and carefully placed into 

an autopsy tray for fixation and later processing on the Autotechnicon. 

D. NOTES 

'Millipore Filter Corp., Bedford, Mass. 

2Technicon Instrument Corp .. Tarrytown, N.Y. 

"Edmond Scientific, Barrington, N.J. 

'Technicon Instrument Corp., Tarrytown, N.Y. 

E. REFERENCES 

Anonymous. 1965. New parasite disease under study. U.S. Bur. . 

Commer. Fish., Commer. Fish. Rev. 27(6): 44. 

Boon, D.B. 1972. The red pigment in discolored oysters and 

soft-shelled clams from the Chesapeake Bay. Chesapeake 

Sci. 13: 334-335. 

Farley, e.A. 1968. Minchinia nelsoni (Haplosporida) disease 

syndrome in the American oyster Crassoslrea virginica. 1. 

Protozool. 15: 585-599. 

Galtsoff, P.S. 1964. The American oyster Crassostrea virginica 

Gmelin . U.S. Bur. Com mer. Fish., Fish. Bull. 64. 480 pp. 

Galtsoff, P.S. 1969. Anomalies and malformations in the shells 

of Crassostrea virginica, pp. 575-580. 111 Neoplasms and 

Related Disorders of Invertebrate and Lower Vertebrate 

Animals (C.J. Dawe and J.e. Harshbarger, eds.), Natl. 

Cancer lnst. Monogr. 31. 

Mackin, 1.G. 1962. Oyster disease caused by Dermocystidium 

marinum and other microorganisms in Louisiana. Publ. 

lnst. Mar. Sci., Vol. 7, pp. 132-229. 

McCormack, G. 1956. Growth characteristics of the pink yeast 

that causes discoloration of oysters. U.S. Bur. Com mer. 

Fish., Commer. Fish. Rev. 18(11): 21-23. 

Paparella, M., ed. 1973. Red oysters. Information Tips, Univ. 

Maryland Mar. Products Lab., Crisfield, Md., pp. 2-3. 

Pauley, G.B. 1969. A critical review of neoplasia and tumorlike lesions in mollusks, pp. 509-539. In Neoplasms and 

Related Disorders of Invertebrate and Lower Vertebrate 

Animals (C.J. Dawe and 1.e. Harshbarger, eds.), Natl. 

Cancer Inst. Monogr. 31. 

Ray, S.M. 1952. A culture technique for the diagnosis of 

infections with Dermocyslidium marinum Mackin, Owen, 

and Collier in oysters. Science 116: 360-361. 

Sanders, B.J. 1972. Animal Histology Procedures of the 

Pathological Technology Section of the National Cancer 

Institute. Government Printing Office, Washington, D.e. 

Sparks, A.K. 1972. Invertebrate Pathology. Noncommunicable 

Diseases. Academic Press, New York. 

Tubiash, H.S., S.V. Otto, and R. Hugh. 1971 . Cardiac vibriosis, 

an infection of the American oyster, Crassoslrea virginica. 

Bacteriol. Proc. 1971, p. 47. 
Processing of soft clam, Mya arenaria. 

Fill. 30. Inserting knife. 

Fig. 31. Carefully cutting clam muscles on lower shell. 

Fig. 32. Cutting off siphon and connecting tissue. 

Fig. 33. Removing crystalline style. 

22 
Processing of duck clam, Macoma balthica. 

Fig. 34. Inserting knife. 

Fig. 35. Cutting lower adductor muscle. 

Fig. 36. Removing (excising) rectum. 

Fig. 37. Taking cross·section. 

23 
Processing of hard clam, Mercenaria mercenaria. 

Fig. 38. Measuring clam, side to side. 

Fig. 39. Inserting knife. 

Fig. 40. CUlling adductor muscle. 

24 

Fig. 41. 

Fig. 42. 

Fig. 43. 

Cutting muscles around edge of mantle. 

Removing clam from shell. 

Rinsing in amhient sea water to remove excess sediment in gills 

and mantle 
He art Cut edge of gll18 

Kid n e y ---------, 

P08terlor retractor 

mU8cle 

P08terlor adductor 

mU8cle 

Style 

HI n g e 

Stomach 

diverticula 

( a moo t h ) ---------,,L-.,r-.,.q{44oJ . .Y....:~ 

(atrlated) ~--" ....... --- Anterior retractor muacle 

An u a -----------+-+~..,..l1\\.:' 

Sip h 0 n a --------~1 

Gonada 

Fig. 44. Gross anatomy of hard clam. The bold parallel lines show the 

location where the tissue for cross-section should be taken 

(Illustration by A.I. Lippson). 

2-5 

Labial palpa 

Anterior adductor 

mU8cle 

(8mooth) 

(atrlated) 

Inte8t1ne 

Foot 
Pericardial cavJly 

Int estine 

,,;, Co':·. "~+--------- Ma nile 

~~~~~?~~~~~- Connective Tissue 

Stomach 

Gills----~ 

~~~"'i=':::c-fj~~-ugestive dive rt i cula 

~~~~~------------Intestine 

Foot---~~---~ 

Fig. 45. Microscopic illustration of cross-section of hard clam 

(illustration by A.f. Lippson). 

26 
Processing of blue mussel, Mytilus edulis. 

Fig. 46. Inserling knife. 

Fig. 47. CUlling mande along shell 10 expose lissue. 

Fig. 48. CUlling byssus. 

27 

Fig. 49. 

Fig. 50. 

Fig. 51. 

Culling cross-seclion. 

Orienling lissue. 

Placing lissue wilh code in aulopsy Iray. 
He art 

Posterlo'!' retractor muscles Style 

Anus S tom a c h ----------~~ 

Digestive dlvertlcula_--zAj,,",~ "'\ ~JN.J03~-~:'Illr- P 0 s t e rio r add u c tor 

In t est I n e --------::~ilf'"JI~~~r-J 

E sop hag u s ------"77i~--Ji' 

Anterior 

m usc I e s --:---:----:-..-:~~~~~~~;:~~ Anterior adducto 

muscle 

Labial palP 

F 0 0 t 

Byssus-------------------__ ~U 

Fig. 52. Gross anatomy of mussel. The bold parallel lines show the 

location where the tissue for cross·section should be taken 

(illustration by A.J. Lippson). 

28 

muscle 

Gills 

Mantle 

Bys.al gland 
In t es tine ---------,,~~~.:.. ... 

Digestive di ve rticula ---4tr~~'-rI~"'..j 

St yle 

Kldney-------~~~a 

Plycat e gan -----t~"'!>:'IMf" t 

Musc Ie ---- --~~~-----.: 

Genita l follicles In 

mantle 

~'- a\ 1Ji:-¥T;~~-Plycat e organ 

~hrl------- Gill s 

''fttf-ffltlHt-f~;H;;:----If\;MlHf- By s sus g I and 

Bys sus musc Ie -----~~~rn ~~ W.~ 

Fig. 53. Microscopic illustration of cross-section of mussel 

(illustration by A.I. Lippson ). 

29 
Processing of sea scallop, A rgopeclen irradians. 

Fig. 54. Measuring scallop. 

Fig. 55. Inserting knife. 

Fig. 56. Cutting muscle. 

Fig. 57. Cutting cross-section. 

30 
H I n g e -------------t--~~!ffIilil!l~\~~\\ 

Dlge8tlve gland 

Stomach 

He art 

Add u c tor m u 8 c Ie 

(smooth) 

(striated) 

Kidney 

An u s ________ ---->,<'1111!1~ 

M.a n tie 

Fig. 58. 

Labial palp8 

~~t:iff;/ff7j~~~""--------- F 0 0 t 

...... ~~~~1x'e>_'t_'t_--- F e m a leg 0 n 8 d 

! 

~ ~J:ib~.~~!I~~lf~~=====-~' :n~t~e s tin e Style 

·: .. "'R~*~~?8st~~~+---- M 8 leg 0 n a d 

Gross analomy of scallop. T be bo ld parallel lines show Ihe 

local ion wbere Ibe lissue lor c ross-seclion should be laken 

(illuslralion by AJ. Lippson ). 

31 

G I" 8 
tomacn-----------~~~~*_-

Digestive divertl cula-l'HI&.l-:" 

Adducto r mu sc le----i---'--''7.'':l 

Go n ad. r'e m a I e ----l------\-N.:..sw.;-Ij.._+.~ 

Kidney-----------~~~~~ \N~ 

Intestine 

Gon ad . m a I e ---------------------'e;rl~ 

Fig. 59. Microscopic iIIuslralion of cross·seclion of scallop 

(illuslralion by AJ. Lippson J. 

32 

Gi II s 

Style 
CHAPTER II. FROZEN TISSUE TECHNIQUE 

A. DESCRIPTION OF TECHNIQUE 

Frozen tissue techniques enable a sampl.e to be quickly 

surveyed microscopically for obvious parasites or lesions. 

Specimens which exhibit suspicious macroscopic signs such as 

pale digestive diverticula, watery condition, mantle recession, 

or other observable abnormalities may be selected for frozen 

sectioning and quick diagnosis. 

The frozen tissue technique presented here may differ from 

methods used with vertebrate tissue primarily by its simplicity. 

Liddell (1967) outlined the following frozen sectioning method 

useful for most molluscan tissue: 

• When performing the necropsy, an extra thin cross-section 

is taken approximately 2 mm below the palps (see Fig. 28). 

• Tissue is quick-frozen to the object holder with its appropriate code (see Figs. 6Oa, b, c, d). 

• Object holder disc is then placed into the microtome chuck 

inside the cryostat. 

• Tissue is gradually advanced and rough trimmed until the 

full surface of the cross-section is exposed . 

• Knife blade and surrounding area are carefully brushed 

clean so that no ice chips or tissue residue will interfere with 

the surface of the tissue block. 

33 

• A frozen section is then cut at 6-8 pm. 

• Tissue is picked up with a clean glass slide or coverglass. 

• The section adheres immediately to the glass which is room 

temperature and the section melts into place. 

• Slide is now ready for staining or frozen storage. 

If immediate inspection of the tissue is desired, the section 

can be stained for a quick diagnosis. 

Successful cutting of frozen tissue on a cryostat or freezing 

microtome is accomplished by the same method that is used on 

a standard rotary microtome designed for sectioning paraffin 

blocks, the main difference being the consistency of the frozen 

tissue. Since molluscan tissue is soft and very moist, it should 

be quick-frozen while fresh. Before attempting to cut the tissue, 

make sure the knife and microtome have been chilled to -2SoC 

and the knife blade is sharp and clean. Consistently good results 

can be obtained with fresh frozen mol.luscan tissue. 

B. REFERENCE 

Liddell, V.A. 1967. Frozen-section technique in shellfish 

research. J. lnvertebr. Pathol. 9: 283-284. 
lLADE 

60b 60c 

Fig. 60. Frozen tissue tecbnique: 

a, tissue coded and frozen in object bolder; b, tissue positioned; 

c, cut; d, picked up on coverglass (illustration by R. Tolley). 

34 

60a 

60d 
CHAPTER III. FIXATION 

A. FORMULAS FOR FIXATIVES AND METHODS 

The choice of a fixative depends on the nature of the tissues, 

the interests of the investigator, and the purpose of the study. 

In most cases a general purpose fixative is preferable, especially 

for work in the field. Ultrastructural examinations demand 

specifically designed fixatives. If facilities are lacking for safe 

disposal of toxic substances like mercury, alternative formulations should be employed. In brief, the nature of the work and 

the facilities at hand determine the optimum fixative for the 

occasion. Table 1 lists formulas for fixatives and methods. 

TABLE 1. F/XA T/VE FORMULAS AND METHODS. 

Fixative 

BOUIN's (Coolidge and Howard, 1979) 

Saturated aqueous 

picric acid 

37-40% formaldehyde 

Glacial acetic acid. 

75 ml 

25 ml 

5 ml 

DAVIDSON's (Shaw and Battle, 1957) 

1 gal stock (J600 ml): 

Glycerin . 

37-40% formaldehyde 

95% ETOH 

Filtered ambient sea 

water (see III. B.) . 

. 400ml 

BOOml 

1200 ml 

. 1200ml 

Fixation time 

Optimum fixation 

time, 4-18 hr. Room 

temp.; pH, acid. 

Optimum fixation 

time, 24-48 hr. 

Refrigerate, 4°C; 

pH, acid . 

Preparation for 

sectioning 

Optimum tissue thickness, 

5-7 mm. To prepare for 

embedment, wash in many 

changes of 70% ETOH . May 

store larger tissue in Bouin's 

or70% ETOH (if necessary). 

Optimum tissue thickness, 

4-5 mm. To prepare for 

embedment, change to 50% 

ETOH. 2-hr minimum; 

change to 70% ETOH, 2-hr 

minimum. May store tissue 

in 70% ETOH or stock 

Davidson's without acetic 

acid (if necessary). 

FORMALIN ( 10% sea water) (Lillie, 1965; Farley, pers. commun.') 

37-40% formaldehyde .... 10 ml Optimum fixation Optimum tissue thickness, 

Filtered ambient sea water time, 24-48 hr. Room 4-5 mm. To prepare for 

(see III. B.) . 90 ml temp.; pH, neutral embedment, wash JO min to 

for best results. 4 hr, depending on size of 

tissue, to remove excess formalin. Store in 70% ETOH 

or 10% formalin. 

GLUTARALDEHYDE 12%) lFeng et al., 1971) 

50% glutaraldehyde .4 ml Optimum fixation 

Filtered ambient sea water time, 2-24 hr. Temp., 

(see Ill. B.) .96 ml 4°C; pH 7.2-7.4. 

Optimum tissue thickness, 

1 mm. To prepare for 

embedment, dehydrate as 

routine for desired plastic 

embedding medium. Wash 

in buffered sea water, pH 

7.2-7.4. Post-fix in buffered 

1 % osmium textroxide, pH 

7.2-7.4. Store in refrigerator, 

4°C. 

35 

Remarks 

Good fixative for protozoans or glycogen (large 

whole objects such as eyeballs). Not recommended for staining azure eosin or Feulgen reaction . 

Remove excess picric acid with many washes of 

70% ETOH or 70% ETOH saturated with lithium 

carbonate . Good for trichrome stains and special 

techniques, e.g. protargol silver. 

Note: Dry picric acid is dangerously explosive. 

Good standard fixative; general molluscan pathology; acceptable for most stains. Best results if 

stock fixative is made in order of ingredients in 

formula . Shake well before using stock fixative 

each time . 

Note: Add 1 part acetic acid to 9 parts stock 

Davidson's prior to fixation. 

Good general fixative , particularly good for field 

and cruise work because of simplicity. For seawater formalin, use filtered ambient sea water 

when fixing tissues in the field. 

Note: After opening a fresh bottle of commercial 

formalin, add a few marble chips or calcium carbonate to prevent onset of acidity. 

Recommended for electron microscopy. 
TABLE I. (conI.) 

Fixative Fixation time 

Preparation for 

sectioning Remarks 

1% GLUTARALDEHYDE/4% FORMALDEHYDE (IG4F) (McDowell and Trump, 1976; Farley, pers. commun.2) 

37-4(f'/0 buffered formalin stock: Optimum fixation Optimum tissue thickness, Good fixative for tissue to be viewed for light or 

Formaldehyde. . .. . 5 gal time , 24-48 hr for 2-3 mm. Wash well in electron microscopy. It is an excellent protein 

Na2HP04 (disodium penetration. See ambient sea water prior to fixative. 

phosphate) . ... . 284 gm original paper for embedment. Store tissue in Note: If not washed well, fixative will interfere 

Phenol red . . . .... 0.5 gm specific information. fixative at room temp. until with Schiffs reaction in PAS, MPAS, and FeulNaOH (sodium hydroxide) Room temp.; pH ready for embedment. Tissue gen stain. May want to use the aldehyde blocker 

. 1.2 gm 7.2-7.4 will tolerate long storage in suggested in staining procedures. Store working 

lG4F seawater fixative working solution: 

37-40% buffered formalin 

stock . .. .. . 120 ml 

50% glutaraldehyde ... . .. 20 ml 

Tap water . .... .. 360 ml 

Filtered ambient sea 

water (see III. B.) . . . 500 ml 

Note: For 10% buffered formalin,dilute 

120 ml of 37-40% buffered formalin 

stock with 88 ml of diluent (distilled 

or sea water, depending on tissue to 

be fixed). 

HELL Y's (Zenker formalin) (Coolidge and Howard, 1979) 

Zenker base . .95 ml Optimum fixation 

37-40% formaldehyde (add time, 6-18 hr. If 

just before using) . ..... .. 5 ml fixative becomes 

turbid, it has lost its 

effectiveness and 

should be discarded. 

Room temp.; pH, 

acid. 

ZENKER's (base) (LUlie, 1965) 

Mercuric chloride . .... .. 50 gm 

Potassium dichromate . . . 25 gm 

Sodium sulfate . . 10 gm 

Distilled water . 1000 ml 

Optimum fixation 

time, 6-24 hr. 

Note: Overfixing will 

dull nuclear detail. 

Room temp.; pH, 

acid. 

ZENKER's (acetic) (Coolidge aDd Howard, 1979) 

Zenker base . . .. 95 ml Optimum fixation 

Acetic acid (add just before time , 6-18 hr. Room 

using) . . 5 ml temp,; pH, acid. 

fixative. fixative at 4°C. Stable for 3 months. 

Optimum tissue thickness, See instructions for Zenker base. 

2-4 mm. To prepare for 

embedment, wash well in 

running water 6-24 hr, 

depending on thickness of 

tissue. Store (if necessary) 

in 70% ETOH. When turbid, 

change to fresh 70% ETOH. 

Optimum tissue thickness, 

2-4 mm. To prepare for 

embedment, wash well in 

running water 6-24 hr, 

depending on thickness of 

tissue. Store (if necessary) in 

70% ETOH. 

Optimum tissue thickness, 

2-4 mm. To prepare for 

embedment, wash wel1 in 

running water 6-24 hr, 

depending on thickness of 

tissue. Store in 70% ETOH. 

36 

Recommended for fixation of protein; gives exceptional cytologic detail in tissue. Remove mercury crystals before staining. 

Note: Good as a mordant for metachromatic 

staining. Use special caution with mercury disposal. A void metal containers. 

Good nuclear fixative. Remove mercury crystals 

before staining. 

Note : Good as a mordant for metachromatic 

staining. Use special caution with mercury disposal. A void metal containers. 
B. SPECIAL NOTE ON SEA WATER 

When sea water is not readily available, use either of the 

following as substitutes where indicated : 

Salt water. using "Forty Fathoms Bio-Crystals" J: 

Approximately 40 gm/liter = 30 0100 (normal saline) 

Approximately 20 gm/liter = 15 °100 (normal saline) 

Approximately 14 gm/liter = 8.5 °/ 00 (normal saline) 

Note: Check the salinity of each batch of sea water with 

a salinometer for best results. 

Salt water BSS (balanced salt solution) (Tripp et al., 1966): 

Sodium chloride (NaCl) ..... ...... 23.50 gm/ liter 

Potassium chloride (KCl) . . . 0.67 gm/liter 

Calcium chloride (CaCI 2) (anhydrous) 1.10 gm/liter 

Magnesium chloride (MgCI 2) (anhydrous) . 2.03 gm/liter 

Magnesium sulfate (MgS04) (anhydrous). 2.94 gm/liter 

Sodium bicarbonate (NaHCO)) . '. 0.02 gm/li.ter 

Potassium phosphate dibasic (K2HP04) 

(anhydrous) 0.19 gm/liter 

Glucose . . . . . . . . . 0.50 gm/liter 

Trehalose 0.50 gm/liter 

Phenol red 0.05 gm/liter 

Example: To make up 7 0100 sea water, use 7 gm of BSS 

in 1 liter of tap or distilled water; adjust pH to 7.2 or 

7.4 with buffers (see VI. A.3.e.). 

Note: This formula has been modified by deleting two 

sugars. Three percent of the above will make the 

equivalent of 30 0/00 ocean sea water (Farley, pers. 

commun .. ). 

C. NOTES ON FIXATION 

Mercury-based fixatives (Zenker and Helly's) have good 

penetration of tissue. They are excellent for protein fixation 

and preservation of nuclear detail if fixation times are observed. 

The mercury must be removed prior to staining. The procedure 

used in our laboratory to recover mercury from spent fixing 

fluids containing mercuric chloride was reported in Technicon 

Bulletin by D. L. Europa, Pathology Laboratories, Bellevue 

Medical Center, New York, N.Y. (Sanders, 1972): 

• Collect fixatives containing mercury in a jug and store in a 

hood or well-ventilated area. 

• Add 20 ml of 13% thioacetamide per liter of mercuric chloride 

waste. The thioacetamide can be stored in a capped glass 

container and is stable for approximately 1 yr. 

• Mix thoroughly, cap, and allow the solution to stand in a 

hood for 24 hr. 

• After 24 hr, filter in a hood with fluted filter paper. The 

residue containing mercuric salt can be stored in a glass jug 

indefinitely for approved disposa!. The clear filtrate can be 

discarded. 

Davidson's fixative is compatible with most of the stains 

used and penetrates through most tissue within 48 hr. 

The 1% glutaraldehyde/ 4% formaldehyde (lG4F) fixative is 

suitable for conversion of paraffin-embedded tissue to plastic 

embedment for use in light and electron microscopy. Penetration 

37 

is slower and less predictable with 1 G4F; therefore, care should 

be taken to cut tissues thinner during the initial necropsy. 

Ten percent seawater formalin or Bouin's is recommended 

for convenience in shipboard or field situations. No refrigeration 

is necessary and, once fixed, the tissues can be held in the 

fixative until returned to the laboratory. However. Bouin's is 

not compatible with the Feulgen-related stains. 

D. PROBLEMS AND RESOLUTIONS 

Table 2 includes some problems or artifacts resulting from 

fixation or inadequate fixation, as well as some means of 

resolving them. 

E. NOTES 

'Modified by C.A. Farley, National Marine Fisheries Service, 

Oxford, Md.; 1979. 

1 Modified by C.A. Farley, National Marine Fisheries Service, 

Oxford, Md.; 1965. 

J Marine Enterprises, Timonium, Md. 

• Modified by C.A. Farley, National Marine Fisheries Service, 

Oxford, Md.; 1979. 

5 Yevich, P.P. and C.A. Barszcz. Preparation of aquatic animals for histopathological examination. Mimeograph, U.S. 

Environmental Protection Agency, Narragansett, R.l.; 

1979. 

F. REFERENCES 

Coolidge, B.1. and R.M. Howard. 1979. Animal Histology 

Procedures of the Pathological Technology Section of the 

National Cancer Institute. 2nd ed. Nat!. Inst. Health Pub!. 

No. 80-275, Bethesda, Md. 

Feng, S.Y., 1.S. Feng, C.N. Burke, and L.H. Khairallah. 1971. 

Light and electron microscopy of the leucocytes of Crassostrea virginica (Mollusca: Pelecypoda). Z. Zellforsch. 

120: 222-245. 

Lillie, R.D. 1965. Histopathologic Technic and Practical Histochemistry. 3rd ed. McGraw-Hill, New York. 

Luna,L.G., ed. 1968. Manual of Histologic Staining Methods 

of the Armed Forces Institute of Pathology. 3rd ed. 

McGraw-Hili, New York. 

McDowell, E.M. and B.F. Trump. 1976. Histologic fixatives 

suitable for diagnostic light and electron microscopy. 

Arch. Patho!. Lab. Med. 100: 405-414. 

Mitchell, H.L. 1966. A simple method of permanent staining 

of intestinal parasites, using dimethyl sulfoxide. Tech. 

Bul!. Regist. Med. Techno!. 36: 45-46. 

Sanders, B.1. 1972. Animal Histology Procedures of the Pathological Technology Section of the National Cancer Institute. Government Printing Office, Washington, D.C. 

Shaw, B.L. and H.1. Battle. 1957. The gross and microscopic 

anatomy of the digestive tract of the oyster Crassostrea 

virginica (Gmelin). Can. J. Zool. 35: 325-347. 

Tripp, M.R., L.A. Bisignani, and M.T. Kenny. 1966. Oyster 

amoebocytes in vitro. 1. Invertebr. Patho!. 8: 137-140. 
TABLE 2. FIXATIVE PROBLEMS AND RESOLUTIONS. 

Problem or artifact 

Large, irregular black deposits 

throughout tissue. 

Small, black granular pigmentation in and 

around blood vessels. 

Tissue swells when put in water bath. 

Tissue too soft; not rigid in paraffin block. 

Possible causes 

Mercuric chloride remaining in tissue when fixed 

with mercury·based solutions. 

Formalin pigment from stock formaldehyde 

becoming too acid (age will decrease pH of 

formalin). 

Soft tissues are more difficult to penetrate with 

fixing agents. 

Improper penetration. 

38 

Solution, if any 

Wash tissue 6-24 hr before processing. Treat with 

Lugol's iodine and 2% sodium thiosulfate during 

hydration in staining (see VI. C.). 

Once formalin is opened , add marble chips 

periodically to help neutralize fixative. Wash 

tissue V2 hr to 4 hr, depending on size of tissue 

(Luna, 1968). 

24 hr after initial fixation, immerse tissues in fresh 

fixative and agitate periodically. Be sure tissues 

are cut thin during initial necropsy (Yevich and 

Barszcz, unpubI 5 ). 

For thorough fixation of soft tissue such as shell fish 

"gapers;' immerse whole specimen in fixative of 

choice for V2 hr to 1 hr. Wash thorough Iy ; process 

as usual. This procedure should firm-up tissue 

before necropsy. Larger molluscan tissues should 

be immersed in fresh fixative after 24 hr in the initial fixative and the specimens agitated periodically during the day to ensure good penetration of 

fixative. When the fixative has adequately preserved the specimen, wash well and follow routine embedment process (Yevich and Barszcz, 

unpubI.S). 
CHAPTER IV. EMBEDDING 

A. GENERAL PROCEDURES 

The embedding sequence for molluscan tissue is comparable 

to that used for human tissue. Once fixation is attained, the 

tissues must be dehydrated . They are then hand-changed from 

the fixative through 50% and 70% ETOH, or washed according 

to specific fixative requirements (see Table I). The remaining 

dehydration, clearing in xylene, and infiltration of paraffin is 

by hand or on an automatic tissue processor such as Autotechnicon' or Ultratechnicon2 (see Table 3). If the entire mollusk 

has been fixed (a procedure not recommended because of 

penetration problems), that portion which is to be used for 

fixation and sectioning should be cut to proper size and immersed 

in fresh fixative. 

Paraffin (ParaplastJ) with a melting point of 56. 6°C is a most 

satisfactory product for year-round use and is kept ready for 

embedding in a paraffin dispenser. The processing basket, 

containing eight trays of tissues, is removed from the tissue 

processor and placed in a paraffin bath. A smaller paraffin 

bath is used for ease in handling individual trays of tissues. 

Commercially made aluminum molds and plastic embedding 

rings are employed. The rings are labeled with a code 

corresponding to the tissues to be embedded. In order for the 

paraffin blocks to be released from the molds easily , the molds 

are dipped before use in a release dip (95 ml of 95% ETOH and 

5 ml of glycerin, Lab Tek4). Heated forceps are used for 

transferring tissues from the tray to the mold to prevent cooling 

of the paraffin and attachment of the forceps to the tissue. 

Once all the equipment is in order, embedding may begin. 

Slowly remove one tray from the basket and place in the small 

paraffin bath. With heated forceps, remove a label from one 

tissue in the tray. Find the corresponding embedding ring and 

choose a mold of appropriate size. Holding the mold by its 

handles with thumb and middle finger, cover the bottom of the 

mold (no more than one-fourth full) with heated paraffin from 

the dispenser. With heated forceps, turn the tissue over and 

gently squeeze to release any trapped air bubbles (Fig. 61). 

Quickly orient the tissue as desired (Fig. 62) and place the 

embedding ring onto the base mold with the correct identification number and hold in place with index finger. Fill the mold 

to the top with paraffin (Fig. 63) and place in a shallow container 

of ice water to cool for approximately 10 min (Fig. 64). This 

process should be completed as rapidly as possible to prevent 

the paraffin from cooling, entrapment of bubbles, and layering 

of the paraffin within the block. After the block has cooled, use 

gentle pressure to remove it from the mold, and place it on a 

tray to be filed and kept in the refrigerator prior to sectioning. 

This embedding procedure yields perfectly shaped blocks cast 

into plastic mounting rings, which can be quickly oriented in 

the microtome vise for sectioning. 

39 

The Technicon basket and trays should be placed in a bath 

of used xylene and soaked overnight to dissolve attached 

paraffin. For cleaning, rinse them in 50:50 ETOH and xylene, 

then in 100% ETOH, and place on paper toweling to dry. Trays 

will then be ready for use. Note: Tissues too large for aluminum 

molds may be embedded utilizing commercially cast lead "L's" 

and copper plates, and mounted on fiber blocks for sectioning. 

B. EMBEDDING SCHEDULES 

The schedules shown in Table 3 are recommended for specific 

types of tissue. The best results for dehydration, clearing, and 

infiltration are obtained only by allowing the tissue adequate 

exposure time to reagents, combined with frequent agitation 

of tissue. 

Since molluscan tissues are soft and dense, a maximum of 

5-mm thickness is recommended for complete infiltration. 

Fresh reagents are used for each step of embedment to prevent 

evaporation and carry-over of reagents. The paraffin baths 

should be checked regularly to prevent the possibility of 

overheating and breaking down of the paraffin. The temperature 

of the paraffin should not exceed the recommended degree 

range (e.g. Paraplast range, 56°-62°C). 

C. PROBLEMS AND RESOLUTIONS 

Table 4 describes problems that could result from incomplete 

dehydration, clearing, or infiltration during the embedment 

process. 

D. NOTES 

1,2 Technicon Instrument Corp., Tarrytown, N.Y. 

J Lancer, St. Louis, Mo. 

4 Miles Laboratories, Inc. , Naperville, Ill. 

E. REFERENCES 

Luna, L.G., ed. 1968. Manual of Histologic Staining Methods of 

the Armed Forces Institute of Pathology. 3rd ed. McGrawHill, New York. 

Sheehan, D.C. and B.B. Hrapchak. 1973. Theory and Practice 

of Histotechnology. c.y. Mosby Co., St. Louis. 
TABLE 3. EMBEDDING SCHEDULES FOR SPECIFIC TYPES OF TISSUE. 

19·hr routine embedding schedule Ion Autotechnicoo) from 70% 

ETO" for molluscan tissue approximately S-mm thick 

7-hr short schedule for embedding small tissue such as larvae by band 

80% ETOH . . 112 hr 

80% ETOH . I hr 95% ETOH . . Y2 hr 

1st 95% ETOH . 

2nd 95% ETOH 

1st 100% ETOH .. 

2nd 100% ETOH 

3rd 100% ETOH 

1st xylene . 

.2 hr 

. . 2 hr 

. ... . 2 hr 

2 hr 

95% ETOH . . . . Y2 hr 

100% ETOH . . ... Y2 hr 

100% ETOH . . . . . . . . . . . I hr 

100% xylene 

Xylene 

Xylene 

· . Y2 hr 

2nd xylene .. 

Y2 xylene/ Y2 paraffin .. 

1st paraffin . 

. I hr 

.. . I hr 

. .... 2 hr 

.2 hr 

. . .. 2 hr 

. . . 2 hr 

Yz xylene/ liz paraffin .. 

Paraffin. 

· . Y2 hr 

. ... Y2 hr 

. Y2 hr 

· . Y2 hr 

Paraffin . 1Y2 hr 

2nd paraffin. Note: Use vacuum oven for infiltration of paraffin . 

Hold tissue in fresh paraffin bath until embedment 

2-hr sbort scbedule for embedding small specimens by tbe Ultra· 

4-hr embedding schedule 100 Ultratechoicon, under vacuum) for 

molluscan tissue not to exceed S·mm thick 

tecbnicon or vacuum oven 

80% ETOH 

1st 95% ETOH. 

2nd 95% ETOH . 

1st 100% ETOH . 

2nd 100% ETOH . 

3rd 100% ETOH 

4th 100% ETOH 

. .. 5 min 

. .. . 10 min 

. ..... . 20min 

.. 15 min 

... 20 min 

20 min 

.30min 

1st xylene . . 15 min 

2nd xylene . . . . . . . . . . .. 20 min 

Y2 xylene/Y2 paraffin . 30 min 

1st paraffin . 20 min 

80% ETOH 

95% ETOH 

95% ETOH 

100% ETOH 

100% ETOH 

100% ETOH 

100% ETOH 

Xylene. 

Xylene. 

Y2 xylene/ Y2 paraffin 

Paraffin. 

Paraffin. 

.. .. 5 min 

.. 5 min 

.. 5 min 

. .. 10 min 

. . . 10 min 

. ... . 10min 

. 10 min 

. 10 min 

. . .. 10 min 

. . . 10 min 

. 20 min 

. .15 min 

2nd paraffin . . . . 35 min 

Hold tissue in fresh paraffin bath until embedment; maximum time 

Note: If ETOH is not available . isopropyl alcohol in combination with 

toluene (instead of xylene) may be used for processing tissues. 

to hold. 3 hr. 

TABLE 4. EMBEDMENT PROBLEMS AND RESOLUTIONS. 

Problem or artifact 

Tissue pulls apart while sectioning with 

large empty spaces; loss of tissue orienta· 

tion. 

Tissue too soft; the block scrapes against 

blade rather than cutting cleanly. 

Crumbling or tearing of section. 

Tissue smells of xylene 

and still too soft. 

Round holes in tissue areas. 

Possible causes 

Improper infiltration. dehydration. clearing. and 

impregnation (usually large tissues or whole 

mollusks). 

Soft tissues are not only difficult to fix. but 

infiltration takes longer. 

Incomplete dehydration. clearing. and/or 

infiltration. Paraffin too hot during infiltration 

and/or embedding. 

Too much carry·over of xylene; paraffin has not 

completely impregnated the tissue. 

Air bubbles trapped under tissue while embedding 

in paraffin. 

40 

Solution, if any 

Large tissues should be processed on a longer embedment schedule, with the aid of a vacuum 

system (Sheehan and Hrapchak, 1973). 

Soft tissues should be cut thin and processed 

quickly for complete fixation. Adequate time 

should be allowed for impregnation of paraffin. 

Consider size and tissue density when 

planning processing of tissue. Always check 

temperature of paraffin (Luna, 1968; Sheehan 

and Hrapchak , 1973). 

Immerse tissue in a clean paraffin bath and agitate, 

while extending the processing time (Sheehan and 

Hrapchak,1973). 

Once tissue is immersed in paraffin, roll it over 

once and orient tissue as desired; this should 

permit air bubbles to escape. 
Embedding. 

Fig. 61. Placing tissue in mold. 

Fig. 62. Orienting lissue. 

FiR. 63. FillinR mold after adding coded ring. 

Fig. 64. Blocks hardening in ice waler. 

41 
CHAPTER V. SECTIONING 

A. TECHNIQUES 

Successful sectioning of tissue for histological examination 

can only be accomplished with patience and acquired skill. 

Sheehan and Hrapchak (1973) in Theory and Practice of 

Histotechnology pose four major requirements for producing 

good histological slides: (1) a skilled technologist, (2) a sharp 

microtome knife, (3) a proper microtome, and (4) properly 

prepared material. Additionally, practice, experience, and 

knowledge of the tissues are necessary to learn the art of 

microtomy. 

Once the paraffin is firm, the blocks are aligned in consecutive order according to the code. They should then be 

refrigerated for 2 hr (5°C) before sectioning. Individual blocks 

should be kept cool until ready for sectioning since a warm 

bl?ck will not section properly. Carefully mount the ring in the 

nucrotome vise, orienting the tissue giUs or identification number 

to the right. Also, orient the head of the microtome so that the 

bottom surface of the block is parallel to the knife-edge. The 

microtome is adjusted to cut 5- to 6-jim sections, the thickness 

usually employed in our studies. A knife angle of between 6° 

and 8° is recommended for the AO Spencer 820 microtomel . 

Using the manual advance lever, trim the block until the tissue 

is fully exposed. Slight angling of the block may be necessary to 

ex.pose all o~ the tissue without cutting the block to a dangerous 

thmness. Lift the ribbon of paraffin sections that has been 

created by cutting with continual, even strokes (Fig. 65). The 

ri~bon of sections is floated on a heated water bath (Fig. 66) 

adjusted to a temperature of approximately 42°C. The water 

bath should be changed frequently and kept clean and clear of 

air bubbles to avoid histological artifacts. Ribbons are preferred 

to individual sections since the thickness of the cut is more 

uniform throughout. 

Pre-cleaned microscope slides with etched ends are made 

ready for tissue sections by placing one drop of Mayer's al bumin 

fixative (Gray, 1954) on a slide; a second slide is placed face 

down on the first and the two are slid apart, evenly distributing 

the fixative on each slide. A thin albumin coating is desired, 

since if it is too thick it will adversely affect the results. The 

coating causes sections to adhere to the slide during staining. 

Before placing the tissue on the slide, and to ensure accurate 

identity, scratch the code number of each tissue on the reverse 

side of the etched end of the slide, using a diamond pen. Dip 

the properly prepared and coded slide , etched side up, into the 

wat.er bath und~r t~e best tissue section on the ribbon (Fig. 67). 

GUide the sectIOn mto place on the slide with a small camel's 

hair brush. Gently raise the slide from the water and, with the 

handle of the brush, draw down each side of the slide gently to 

cu~ the section from the excess paraffin of the ribbon (Fig. 68). 

Usmg the brush or a probe, orient the tissue centrally on the 

slide, leaving at least one-eighth-inch clearance at the end of 

the slide. This will prevent the coverslip from coming in contact 

with the groove in the storage box. Place the slides in a rack to 

drain excess water, then into a drying oven set at 42°C for 

approximately 12 hr, or until the sections are completely dry. 

Note: The temperature of the water bath and oven should not 

exceed 42°C for tissues embedded in Paraplast (melting point 

56°- 57°C). Melting tissues on the slides can cause distortion 

of cells in the tissue. 

The completely dry slides with affixed tissue sections are 

42 

removed from the drying oven and placed in a staining holder. 

Grasp the holder so the handle is to the left. The slides are 

placed in the holder vertically, etched end up and facing away 

from you, front to back, in consecutive numerical order. The 

slides are now ready for staining. 

The step-by-step preparation of slides follows. 

B. PREPARATION OF TISSUE SECTIONS 

1. Blocks 

• Gradually chill before cutting (this cuts down on thick 

compressed tissue sections); extreme cold will crack the paraffin. 

The blocks are kept refrigerated until ready to section, and 

during cutting they are kept in an ice bath in intended cutting 

sequence. 

• Block should be parallel to the knife-edge. Before cutting, 

the blocks can be trimmed for different slide requirements 

(example, serial sections) or for proper orientation of tissue. 

2. Microtome 

• Know your microtome; keep it properly serviced and adjusted. 

(I) For the AO Spencer microtome, a knife angle of 6°_8° 

is recommended. 

(2) When knives become too short in width (because of 

frequent sharpening) to fit in the microtome holder, a 

pencil can be used to lift the blade uniformly higher. 

Diamond etching pens are perfect for this purpose 

because they are sturdy and remain straight. 

(3) Keep microtome well oiled and lubricated to assure 

optimum operation. 

(4) Tighten set screws on knife holder and block vise 

before cutting. 

3. Knives 

• Hand honing of knives usually gives the very best results for 

cutting tissue sections. A combination of Ivory soap and Lava 

soap is recommended on a Belgian honing stone. Figure 69 

illustrates the honing action. 

• The AO automatic knife sharpener2 has been very satisfactory 

for sharpening new or reconditioned knives. B. Coolidge and 

R. Howard (pers. commun.J) suggested the combination of an 

abrasive (Metadi diamond abrasive·) and a lubricant (Shandon 

microsharp lubricating fluidS). Best results have been attained 

by using only the coarse setting on the machine. 

4. Sectioning 

• Tissue sections are cut routinely at 5-6jim, selecting the best 

section within a ribbon for uniform thickness. 

• Oyster, clam, scallop, and mussel tissue ribbons are generally 

cut and transferred to a water bath. The bath temperature 

should not exceed 42°C. The sections should be picked up on 
slides quickly to reduce the possibility of tissue breakdown. 

The water bath should be cleaned daily to eliminate albumin 

build-up and bacterial growth in the water. Filling the water 

bath with room temperature water will reduce the accumulation 

of air bubbles in the bottom of the bath when it has heated 

to 42°C. 

• A warming tray set at 42°C is used when doing serial sections 

of small tissue or when sectioning problem tissue. Two 

techniques follow for special serial section studies. 

Small tissue for serial sections (Fig. 70): 

Shellfish larvae or seed oysters must often be embedded 

whole because of their size. The tissue should still be oriented 

for the cross-section. However, to ensure that all desired tissues 

are exposed for study, serial sections may be requested. 

(1) Hand trim the block as close to the tissue as possible. 

(2) Make sure the block is parallel to the knife after 

trimming to ensure better serial sections. 

(3) Line up an estimate of the number of albuminized 

slides necessary, e.g. 10 at a time, coded consecutively. 

(4) Cover the slides with a small pool of distilled water. 

(5) Trim to the tissue with microtome and begin saving 

the ribbons. 

(6) Count the number of sections which will fit under 

the intended coverslip size, lay sections down directly 

on the slide, top to bottom, and ribbons across left to 

right. 

(7) Continue this procedure uniformly for the number of 

slides necessary to cut through the tissue. 

(8) Carefully line up ribbon strips and drain excess water. 

(9) Wipe the back of the slide dry. 

(10) Place on warming tray at 42°C to allow sections to 

flatten evenly. 

(11) Eliminate air bubbles, folds, or rolls by teasing out 

with gentle manipulation of two wet brushes. 

(12) Place slides onto racks and put in oven at 42°C to 

continue drying. 

Note: For problem tissue that tends to split or break apart, 

continue to ice block in between ribbons. Lay sections 

down in sequence on wax-coated paper (photographic 

paper liners are good for this procedure) and pick up with 

a wet brush to transfer to consecutive slides. 

Special blocks for serial sections (Fig. 71): 

To fulfill requests for study slides for instructional purposes, it 

may be necessary to produce a large number of slides from the 

same block. Since blocks may be valuable examples of a specific 

disease , parasite, or shellfish condition , care is taken to cut a 

minimum of the tissue from the block. 

(1) Albuminize and label desired number of slides. 

(2) Tape down black paper to the counter top. Use the 

highly wax-coated photographic liner paper so the 

sections do not stick. 

43 

(3) Carefully align the block in the chuck of the microtome. 

(4) Line the slides up consecutively, but not touching, 

and add a small pool of water (avoid overflowing the 

slides). The counter surface should be flat for easier 

slide manipulation . 

(5) Begin sectioning the chilled block; lay the ribbons on 

the paper in the order of cutting in a bottom-ta-top 

sequence. Try to keep as much of a continuous cutting 

motion as possible so that sections will be cut in 

uniform thickness. 

(6) Separate the sections by cutting between each with a 

sharp razor blade. 

(7) Lift cut sections with a wet brush and float in sequence onto the coded slides. 

(8) Orient tissue properly and drain water from the slides. 

(9) Place slides on warming tray until sections stretch out. 

Warning: For ripe oysters, clams, or very soft tissues, 

reduce time on the warming tray since soft tissues will 

sometimes break down after long exposure to heat. 

(10) Place slides onto racks and put in oven at 42°C to 

complete drying. 

C. PROBLEMS AND RESOLUnONS 

Normally an inexperienced technician will encounter many 

problems when sectioning tissue. The ability to recognize the 

source of the difficulty and to remedy it is important. A list of 

commonplace problems, their causes, and possible solutions is 

shown in Table 5. 

D. NOTES 

1,2American Optical Corp., Buffalo, N.Y. 

3Animal Histology Unit, National Cancer Institute, Bethesda, 

Md.; 1977 

'Buehler Ltd., Evanston, Ill. 

5Shandon Southern Instruments, Inc., Sewickley, Pa. 

E. REFERENCES 

Gray, P. 1954. The Microtomist's Formulary and Guide. The 

Blakiston Co., New York. 

Luna, L.G., ed. 1968. Manual of Histologic Staining Methods 

of the Armed Forces Institute of Pathology. 3rd ed. 

McGraw-Hili, New York. 

Sheehan, D.C. and B.B. Hrapchak. 1973. Theory and Practice 

of Histotechnology. c.Y. Mosby Co., St. Louis. 
TABLE 5. SECTIONING PROBLEMS AND RESOLUTIONS. 

Problem or artifact 

Grooves through tissue; tissue torn and 

separated. 

Grooves through tissue, continuing from 

one block to another. 

Entire section splits, including clear 

paraffin. 

Tissue rubbed or smudged, sometimes 

pulled apart; thickened areas indistinct. 

Compression (tissue section noticeably 

smaller than block). 

Shearing (horizontal parallel folds in tissue 

sections). 

Tissue has raised round areas. 

Moth-eaten appearance. 

Sections are alternately thick and thin, 

usually with compression of thin sections. 

Lifting of sections on upstroke. 

Block lifts ribbon. 

Tissue with rows of round raised areas. 

Tissue looks distorted. Paraffin and tissue 

appear melted. 

Tissue separates quickly on water bath. 

Ribbon curved. 

Possible causes 

Sand, shell, miscellaneous debris left in tissue prior 

to fixation. Unique to species: scallop-sensory 

eyes; mussels- byssus; clams-crystalline style; 

and various bivalves-pearls. 

Knives need reconditioning. 

Nick in knife blade. 

Diny knife; lint, hair, other debris, or nick in knife 

blade. 

Dirty knife, probably coated with paraffin on the 

back. 

Block too warm. Section too thin. Knife angle too 

close to vertical. Dull knife. 

Dull knife or wrong knife angle. 

Air bubbles trapped under tissue. 

Rough cutting tissue. 

Set screws are loose on knife holder or the block 

is not tight. Block of tissue is too hard or the tilt of 

the knife is insufficient to clear base. This also 

may occur when the block is too large or too warm. 

Too vertical a knife angle or dull knife. 

Ribbon charged by static electricity (check by 

testing whether or not ribbon sticks to everything 

else). No clearance angle. Upper edge of block has 

fragments of wax on it. Edge of knife (either front 

or back) has fragments of wax on it. 

Shearing when section is put on water bath or 

warming tray. 

Water bath too hot , warming tray too hot, or slide 

oven too hot. 

Water bath too hot, breaking down paraffin and 

tissue. 

Edges of block not parallel; knife not uniformly 

sharp, causing more compression on one side of 

block than other. 

44 

Solution, if any 

Inspect tissue during preparation before cutting 

cross-section. Check knives under dissecting 

scope for damage after sectioning problem tissue. 

Use disposable blades for problem tissue. 

Move to new area of knife ; resharpen. If not removed by sharpening, recondition knives (Luna, 

1968). 

Clean back and front of knife carefully; move to 

another area on knife or resharpen knife (Luna, 

1968). 

Clean back of knife after rough trimming of each 

block. 

Sharpen knife. Strive to have tissue section comparable in size with exposed tissue in block. Keep 

tissue chilled before sectioning (Luna, 1968). 

Sharpen knife. Check angle of knife. 

Make sure water bath is bubble-free. Lay ribbon 

in bath gently to avoid large bubbles or folds. 

Bubbles can be gently teased out of tissue area by 

brush manipulation while tissue is floating on bath 

(Gray, 1954). 

Select tissue in middle of 6-flm ribbon. Several 

6-flm sections should be taken after rough trimming of block to avoid problem (Luna, 1968). 

Check microtome before cutting. Adjust angle. 

Cool block [or at least Y2 hr before sectioning 

(Gray , 1954; Luna , 1968; Sheehan and Hrapchak, 

1973) 

Check knife angle. Sharpen knife (Sheehan and 

Hrapchak, 1973). 

Increase room humidity. Alter knife angle to give 

clearance angle. Scrape under surface of block 

with safety razor blade. Clean knife with xylene 

(Gray , 1954). 

Avoid selecting sections with shearing lines that 

extend into tissues. 

Check temperature of paraffin routinely before a 

tissue sample is processed; maintain warming tray 

and water bath temperature at 42°C. 

Maintain 42°C in water bath; transfer tissue to 

slides quickly. 

Trim block evenly. Try another portion of knifeedge or resharpen knife. Let block cool (Gray, 

1954 ; Sheehan and Hrapchak , 1973), 
TABLE 5. SECTIONING PROBLEMS AND RESOLUTIONS. (Continued) 

Problem or artifact 

Sections distorted. 

Sections bulge in middle. 

Tissue breaks away from paraffin or is 

shattered by knife. 

No ribbon forms because : paraffin 

crumbles; sections, though individually 

perfect, do not adhere; or sections roll 

into cylinders. 

Sections appear wrinkled. 

Chatter of tissue. 

Possible causes 

Blunt knife. Ribbon too stretched when laid down 

on water bath. Tissues not properly hardened 

before embedding. 

Paraffin cool in center, warm on outside. Only 

sharp portion of knife is that which cuts center 

of block. Object may be too hard for paraffin 

embedment, or some clearing agent remains in 

object. 

If tissue appears chalky and shatters under knife, 

it is not impregnated with paraffin. If tissue 

shatters under knife, but is not chalky, it is too hard 

for paraffin sectioning. If tissue pulls away from 

paraffin, but does not shatter, the wrong dehydrant, clearing agent, or paraffin has been used . 

Paraffin contaminated with clearing agent. Knife 

angle wrong. Dull or dirty knife. Warm block. 

Blunt knife used for cutting. Water bath used for 

flattening too hot, so that folds in sections are 

fused into position. 

Occurs frequently with very ripe oyster tissues. 

The gonads are soft tissue and, therefore, difficult 

to infiltrate. 

45 

Solution, if aoy 

Use suitable knife. Handle ribbons in short 

lengths; lay down gently. Use more suitable 

fixative or fix longer. Take extra care in dehydrating, clearing, and embedding (Gray , 1954). 

Let block cool evenly in ice bath. Try another 

portion of knife-edge or resharpen knife. Reembed object in appropriate media or melt paraffin to remove excess clearing agent (Gray, 1954). 

Discard block and start again. If tissue is irreplaceable, try dissolving off paraffin, re-dehydrating, re-clearing, and re-embedding. Avoid xylene 

in clearing muscle tissue (Gray, 1954). 

Re-embed. (Note: Paraffin very readily absorbs 

hydrocarbon vapors) (Gray, 1954). If the section 

is cut very slowly, and the edge of the section held 

flat with a brush, ribbons may sometimes be 

formed. Adjust knife angle. 

None. Prevent by sharpening knife and cut new 

sections. Control temperature of water bath used 

for flattening. Make sure that slide is clean , so that 

water flows uniformly over it. 

Allow block to warm to room temperature and 

try to ribbon. Be sure knife is sharp and tight in 

holder. 
Seclioning. 

Fig. 65. Lifting a ribbon of oysler lissue seclions. 

Fig. 66. Laying seclions down on waler balh. 

Fig. 67. Direcling sec lion onlo slide. 

Fig. 68. Culling e~cess paraffin ribbon from edge of slide. 

46 
A 

B 

Fill. 69. Action used in hand boning 01 knife blades 

(illustration by R. Tolley). 

47 
Fig. 70. Serial sec lions of small lissues. 

48 
Fig. 71. Serial sections of lar~e tissues. 

49 
CHAPTER VI. STAINING 

A. METHODS AND USE presented as notes, and the principal reference. 

The collection of staining methods described in this chapter 

provides routine general tissue stains, as well as a variety of 

special stains for the demonstration of specific tissue elements. 

The format for each stain description includes the staining 

procedure, its results, necessary reagents, additional information 

To provide for ease of use of these stain descriptions, Table 

6 gives a synoptic listing of these stains, keyed to the pages on 

which their descriptions occur, and Table 7 provides a synoptic 

listing of the tissues and tissue elements to be stained for, 

keyed also to the appropriate pages. 

TABLE 6. SYNOPTIC LISTING OF STAINS. 

Name of stain Acronym Specific for Approll. time Page no. 

Alcian blue AB Acid mucopolysaccharides 3 hr 20 min 55 

Alizarin red S ARS Calcium I hr 45 min 56 

Azure-eosin/ Giemsa AZE Blood 1 hr. fresh blood 57 

Azure-eosin/ Giemsa AZE Blood ; gram positive I hr 30 min. fixed 58 

bacteria tissue 

Feulgen gram Bacteria; gram positive I hr 50 min 60 

Feulgen picromethyl FPM DNA; collagen, connective tissue I hr 35 min 61 

blue 

Gram Bacteria; gram positive and I hr 10 min 62 

negative 

Grocott's methenamine GMS Fungi 3 hr 15 min 63 

silver 

Harris' hematoxylin HHE) General tissue I hr 30 min 64 

and eosin (progressive) 

Harris' hematoxylin HHE:! General tissue 1 hr 10 min 65 

and eosin (regressive) 

Heidenhain's iron IHE Nuclear detail 1 hr 3D min 66 

hematoxylin and eosin 

Mallory's hematein Copper metals 1 hr 67 

Mallory's trichrome Collagen, connective I hr 68 

(modified) tissue 

Malt periodic acid MPAS Glycogen digestion and PAS 2 hr 20 min 69 

Schiff positive substances 

Malt periodic acid MPAS-AB Neutral and acid 5 hr 70 

Schiff-alcian blue m ucopolysaccharides 

Oil red 0 ORO Fats 10 min, fresh 71 

frozen tissue 

Periodic acid Schiff PAS) Glycogen; fungi; neutral I hr 15 min 72 

mucopolysaccharides 

Periodic acid Schiff PASz Glycogen; fungi; neutral I hr 45 min 73 

mucopolysaccharides; good 

nuclear detail 

Perls' Prussian blue PPB Ferric iron 2 hr 74 

50 
TABLE 6. SYNOPTIC LISTING OF STAINS. (Continued) 

Name of stain Acronym Specific for ApproK. time Page no. 

Sudan black B SBB Phospholipids 2 days 75 

Toluidine blue a TBO Hemocytes 30 min, fresh blood 76 

Von K6ssa Calcium 2 hr 77 

Wilder's reticulum WR Connective tissue, 1 hr 78 

reticulum 

Ziehl and Harris' ZHH Acid-fast spores I hr 45 min 79 

hematoxylin 

Ziehl and methylene Acid-fast bacteria I hr 30 min 80 

blue 

TABLE 7. SYNOPTIC LISTING OF TISSUES AND TISSUE ELEMENTS TO BE STAINED FOR. 

Tissues and 

tissue elements Acronym Name of stain ApproK. time Page no. 

Acid-fast: 

(a) Bacteria Ziehl and methylene blue 1 hr 30 min 80 

(b) Spores ZHH Ziehl and Harris' 1 hr 45 min 79 

hematoxylin 

Bacteria: 

(a) Acid-fast (see acid-fast) 

(b) Gram positive Feulgen gram; I hr 50 min 60 

AZE azure-eosin/ Giemsa I hr 30 min 58 

(c) Gram positive Gram 1 hr 10 min 62 

and negative 

Blood: 

(a) Differentiation AZE Azure-eosin/Giemsa 1 hr, fresh blood 57 

AZE Azure-eosin/Giemsa 1 hr 30 min , fixed 58 

tissue 

(b) Hemocytes TBO Toluidine blue 0 30 min , fresh blood 76 

Calcium 

ARS Alizarin red S I hr 45 min 56 

Von K6ssa 2 hr 77 

Connective tissue : 

(a) Collagen Mallory's trichrome I hr 68 

(b) Collagen FPM Feulgen picromethyl blue 1 hr 35 min 61 

(c) Reticulum WR Wilder's reticulum 1 hr 78 

Copper metals Mallory's hematein I hr 67 

DNA FPM Feulgen picromethyl blue 1 hr 35 min 61 

Fats and 

phospholipids: 

(a) Fats ORO Oil red 0 10 min, fresh frozen tissue 71 

(b) Phospholipids SBB Sudan black B Allow 2 days 75 

51 
TABLE 7. SYNOPTIC LISTING OF TISSUES AND TISSUE ELEMENTS TO BE STAINED FOR. (Continued) 

Tissues and 

tissue elements 

Fungi: 

(a) Mycelia and hyphae 

(b) Walls 

General tissue: 

(a) Progressive 

hematoxylin 

(b) Regressive 

hematoxylin 

Glycogen: 

(a) See fungi (PAS) 

(b) Glycogen 

digestion 

Iron 

Mucopolysaccharides: 

(a) Acid 

(b) Neutral (see PAS, MPAS) 

(c) Neutral and acid 

Nuclear detail 

Acronym 

GMS 

PAS) 

PAS2 

HHE) 

HH ~ 

MPAS 

PPB 

AB 

MPAS-AB 

IHE 

Name of stain Approx. time 

Grocott's methenamine 3hrl5min 

silver 

Periodic acid Schiff I hr IS min 

Periodic acid Schiff with I hr 45 min 

Heidenhain's counterstain 

Harris' hematoxylin and I hr 30 min 

eosin 

Harris' hematoxylin and I hr 10 min 

eosin 

Malt periodic acid Schiff 2 hr 20 min 

Peds' Prussian blue 2 hr 

Alcian blue 3 hr 20 min 

Malt periodic acid 5 hr 

Schiff-alcian blue 

Heidenhain's iron I hr 30 min 

hematoxylin and eosin 

52 

Page no. 

63 

72 

73 

64 

65 

69 

74 

55 

70 

66 
1. Standard Format for Paraffin-Embedded 

Sections 

To eliminate the repetitive cycle of solutions and times, our 

stain format for paraffin-embedded sections will read as fo.llows: 

deparaffinize, hydrate to water, stain protocol, dehydrate, clear, 

and mount. Blot slides after immersion in each solution to 

avoid carry-over to next solution. Some stain protocols indicate 

a fixative of choice for best results; therefore, it is essential for 

the requestor to designate the fixative. 

a. Deparaffinize 

Immerse slides in two changes of xylene for 5 min each for 

sections cut from 5-6 11m thickness. Change xylene regularly to 

ensure complete deparaffinization. 

Thicker sections will require three to four changes of xylene 

and longer immersion times. 

b. Hydrate to Water 

Carry slides through descending concentrations of ethyl 

alcohol to water as follows: 

100% ETOH . 2 changes 5 min each 

95% ETOH . 1 change 3 min each 

70% ETOH . 1 change 3 min each 

50% ETOH . 1 change 3 min each 

30% ETOH .. 1 change 3 min each 

Distilled water 2 min each 

The 100% ETOH should be renewed frequently depending 

on the number of slides involved to avoid carry-over of xylene. 

Obvious xylene contamination in 100% ETOH will leave an 

insoluble scum on slides when carried into diluted alcohols. 

The solutions should flow smoothly off slides before descending 

to the next alcohol. 

53 

c. Removal of Mercury 

Tissues preserved in a fixative containing mercury (Helly's, 

Zenker's, or Zenker acetic) must be treated with Lugol's iodine 

and sodium thiosulfate before staining to prevent mercury crystal artifacts. 

To prepare for stain, deparaffinize slides and hydrate to 

water. Add Lugol's iodine solution (iodine should be approximately 10% of solution) to the first 70% ETOH and place slides 

in solution for 10 min. Rinse slides in 70% ETOH, change to 

50% ETOH, and follow the 30% ETOH with 2% sodium thiosulfate for 5 min. Wash in running tap water 10 min, then return slides to distilled water. Continue with staining procedure 

(Mitchell, 1966). 

Lugol's iodine: Potassium iodide . 

Iodine .. 

Distilled water .. 

6.0gm 

4.0 gm 

100.0 ml 

2% sodium thiosulfate: 2 gm sodium thiosulfate per 

100 ml distilled water 

d. Dehydrate 

When the stain and counterstain are complete, the slides are 

taken through ascending ethyl alcohols: 

95% ETOH .. . . . . . 

100% ETOH 

100% ETOH ..... .. . . 

e. Clear 

6 dips or 3 min 

)liz min 

3 min 

Immerse slides in two changes of xylene for 5 min each. 

Change xylenes regularly to avoid alcohol carry-over. 

f. Mount 

Slides are ready to coverslip with the mounting medium of 

choice. 
2. Pre-Staining Techniques 

a. Aldehyde Blocking Technique 

This technique precedes Schiff (Feulgen)-type stains where 

nonspecific aldehyde staining has occurred. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Place slides in a saturated solution of dimedone' 

overnight 

(4) Rinse thoroughly in running tap water - Y2 hr 

(5) Proceed with desired Schiff (Feulgen)-type stain 

Reagent: 

'Dimedone (5.5-dimethyl-l ,3-cyclonexanedione) 

saturated solution 

Dimedone ..... .. . .. . .... ... .. .... . ...... 0.5 gm 

Distilled water . . ...... 100.0 ml 

Stir or shake intermittently for at least 5 hr and 

filter before use. 

Note: "Completeness of blockade can be tested by transferring 

a treated . rinsed slide directly to the Schiffs reagent, 

followed by the usual metabisulfite and water rinses. There 

should be no staining if the blockade of aldehyde groups is 

complete ... " (Feder and O'Brien, 1968). 

Reference: Feder. N. and T.P. O'Brien. Am. J. Bot. 55: 123-142; 

1968. 

b. Celloidin Coating for Slides 

Use this technique when tissues tend to come loose from 

slides. 

Procedure: 

( 1) Deparaffinize 

(2) 100% ETOH - 2 changes - 5 min each 

(3) Place in 0.5% celloidin' - 1 min 

(4) Drain 

(5) Place in 70% ETOH 

(6) Hydrate to water 

(7) Stain as usual 

Reagent: 

'I % celloidin stock solution 

Parlodion ...... ........ . . . . . .. . . . 

100% ETOH ..... .... .... . 

Let stand till parlodion is dissolved. 

l.Ogm 

100.0 ml 

54 

0.5% celloidin working solution 

1 % celloidin stock 

Ethyl ether (anhydrous) 

(Danger. Avoid open flame or electrical 

discharges_ Use hood. Avoid breathing vapor.) 

20.0 ml 

20.0ml 

Note: (1) For Feulgen picromethyl blue (FPM), remove celloidin 

after Schiff's (Feulgen), with one change of 70% ETOH 

and two changes of 100% ETOH, then hydrate to 70% 

ETOH and to water, and continue staining with picromethyl 

as usual. (Butanol will not remove celloidin; therefore. the 

celloidin must be removed in 100% ETOH before 

completing the stain.) (2) Do not use celloidin with alcian 

blue staining. The celloidin will pick up alcian blue and 

will mask the entire slide. (3) Use extreme caution when 

handling or storing celloidin. It is highly explosive. 

Reference: Coolidge, B.J. and R.M. Howard. Animal Histology Procedures of the Pathological Technology Section 

of the National Cancer Institute. 2nd ed . Natl. Inst. Health 

Publ. No. 80-275, Bethesda, Md.; 1979. 

c. Acid Clean Glassware 

Dichromate-acid cleaning solution: 

Potassium dichromate ........ . 

Distilled water . _ ...... . 

Dissolve with low heat. 

Add slowly: 

Sulfuric acid 

200gm 

2000 ml 

200ml 

Extreme care must be taken in preparation of this solution 

and its use. Take the following precautions: 

(I ) Slowly add the acid to the aqueous solution. Place 

container in stone sink while pouring acid. 

(2) Wear protective face mask, acid-resistant apron and 

gloves. Check gloves for holes each time they are 

used. 

(3) Potassium dichromate has allergenic properties and 

should be handled with care. 

(4) Store in acid-proof container. 

Clean glassware is submerged in the cleaning solution for 2 

hr or more, rinsed well in running water and several changes of 

distilled water. Dry and store in a dust-free cabinet. 

To dispose of used acid bath, safety waste containers are 

recommended. Do not pour down the drain_ 

Reference: Coolidge, B.J. and R.M. Howard. Animal Histology 

Procedures of the Pathological Technology Section of the 

National Cancer Institute. 2nd ed. Natl. Inst. Health Pub!. 

No. 80-275, Bethesda, Md.; 1979. 
3. Staining Methods 

a. Alcian Blue (AB) (3 hr 20 min) 

Specific for acid mucopolysaccharides. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water and remove mercury crystals, if 

present (see VI. A.l.c.) 

(3) Distilled water - 2 min 

(4) Alcian blue' - 30 min 

(5) Rinse briefly in running water-4-6 dips 

(6) 0.5% alcoholic ammonium hydroxidel-2 hr 

(7) Rinse briefly in running tap water-4-6 dips 

(8) Counterstain in nuclear fast redJ (Kernechtrot)-5 

min 

(9) Rinse briefly in distilled water (swish through in large 

container) - 6 dips 

(10) Dehydrate 

(11) Clear 

(12) Mount 

Results: 

Acid mucopolysaccharides- brilliant blue 

Nuclei-bright red or shades of blue 

Background - pink to red 

Other basophilic substances-various shades of blue or 

purple 

Other cytoplasmic elements-shades of pink 

Reagents: 

'0.1 % alcian blue in 0.01 M HCI (hydrochloric acid) (pH 2.2) 

- Helly's fixed material (Johnson, 1980) 

Alcian blue 8GX 

1 NHCI . 

Distilled water 

1.0 gm 

10.0 ml 

990.0 ml 

0.1 % alcian blue in 3% acetic acid - Davidson's or formalinfixed tissue (Johnson, 1980) 

Alcian blue 8GX 

Distilled water . 

Glacial acetic acid 

0.1 gm 

.. 97.0ml 

. 3.0ml 

Note: Check expiration date of alcian blue stain. 

20.5% alcoholic ammonium hydroxide (Johnson , 1980) 

Ammonium hydroxide .. 3.0 ml 

95% ETOH 597.0 ml 

JNuclear fast red (Kernechtrota) 

5% aluminum sulfate (AI2(S04h ): 

AI2(S04)J· ...... . ... . . . 

Distilled water. 

0.1 % nuclear fast red: 

Kernechtrot nuclear fast red .... . .. .. . 

5% AliS04h. 

Note: Refrigerate; good till loses color. 

25.0 gm 

500.0 ml 

. .. 0.5gm 

500.0 ml 

55 

References: Pearse, A.G.E. Histochemistry, Theoretical and 

Applied. 2nd ed. Little, Brown and Co., Boston ; 1960. 

(Modified by P.T. Johnson. Histology of the Blue Crab, 

Callinectes sapidus. A Model for the Decapoda. Praeger, 

New York; 1980.) 
b. Alizarin Red S (ARS) (1 hr 45 min) 

Specific for calcium (Lillie, 1965). Use a positive control test 

slide. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) 0.1 % alizarin red S' - 1 hr 

(4) Rinse in several dips of tap water 

(5) Counterstain in 1 % light greenl - count to 15 quickly 

(6) Distilled water - 6-10 dips 

(7) Dehydrate 

(8) Clear 

(9) Mount 

Results: 

Calcium salts - intense reddish-orange 

Background - pale green 

Reagents: 

'0.1 % alizarin red S 

Alizarin red S . 

Distilled water . 

21% light green (tends to overstain) (Dahl and 

Dole, 1952) 

Light green, SF yellowish . 

Distilled water . 

Glacial acetic acid 

. 0.1 gm 

100.0 ml 

1.0 gm 

100.0 ml 

1.0 ml 

Reference: Luna, L.G., ed. Manual of Histologic Staining 

Methods or the Armed Forces Institute of Pathology. 3rd 

ed. McGraw-Hill, New York; 1968. 

56 
c. Azure-Eosin/Giemsa (AZE) (1 hr) 

Specific for fresh blood smears fixed in methanol. 

Procedure: 

(I) Giemsa 1, pH 6.8-7.1 - 1 hr 

(2) Rinse in buffered distilled water2 

(3) Blot individual slides to prevent water spots 

(4) Air-dry 

Results: 

Nuclei and basophilic substances - blue 

Cytoplasm and acidophilic substances - pink 

Reagents: 

lGiemsa 

Giemsa (azure B type) (American Scientific 

Cat. No. S-7712-\b) 

Buffer (will·vary with blood species; see 

McIlvaine-Lillie buffer table, Vl.A.3.e.). 

Distilled water 

2 Distilled water 

Buffer to same pH of stain used 

5.0 ml 

10.0 ml 

185.0 ml 

Note: (1) If coverslips are required , allow the blood smears to 

air-dry completely, then dip in clean xylene and mount. 

(2) For old fish blood smears, immerse slide in the following 

solution for 5-10 min: 

100% ETOH .. ... . 60.0 ml 

Glacial acetic acid 10 drops 

Rinse in distilled water and continue with Giemsa stain 

procedure (Drury and Wallington, 1967). 

Reference: Lillie, R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed. McGraw-Hill, New York; 1965. 

57 
d. Azure-Eosln/Giemsa (AZE) (1 hr 30 min) 

Metachromatic stain for fixed tissues. Specific for blood 

studies and gram positive bacteria. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Giemsa' (see table for pH. VI.A.3.e.) - 1 hr 

(4) Drain 20 sec 

(5) Acetone - IV2 min 

t6) Acetone - 3 min 

(7) 50:50 acetone/ xylene - 1 V2 min 

(8) Clear 

(9) Mount 

Results: 

Nuclei - blue to violet 

Cytoplasm - varying shades of light blue or pale pink 

Muscle - bright pink 

Bacteria - blue 

Reagent: 

'Giemsa 

Giemsa (azure B type) (American Scientific 

Cat. No. S-7712-1b) . 5.0 ml 

Buffer (McIlvaine-lillie buffer table. 

VI.A. 3.e.) 10.0 ml 

Acetone . . . . . . . . . . . . 25.0 ml 

Distilled water 160.0 ml 

Note: Use fresh Giemsa stain for each set of slides. The 

stain starts to deteriorate after 1 hr. 

References: Lillie. R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed. McGraw-HiII. New York; 1965. 

(Modified by M.W. Newman . National Marine Fisheries 

Service. Oxford. Md.; 1971.) 

58 
e. Fixative Guide for Giemaa stain 

The following table developed by Martin Newman (pers. 

commun.) provides the recommended pH for buffers to be 

used with various fixatives. After the preferred pH has been 

determined, one should consult the McIlvaine-Lillie buffer 

table also shown below: 

Fixative 

Formalin . 

Hollande . 

Formalin, followed by decalcifier 

Zenker formalin 

Davidson's . 

Susa . 

Helly's . 

Bouin's 

.... 4.5 

· 4.75 

..... 4.8 

· 4.95 

4.95 

· 4.95 

. . 5.1 

... .. 5.5 

McIlvaine-Lillie buffer table (Lillie, 1954) 

Stock solutions: 

MilO citric acid 

Citric acid . . . ...... . . 2.1 gm 

25% methanol (25 ml methanol + 75 ml 

distilled water) . 100.0 ml 

MIS disodium phosphate (sodium 

phosphate dibasic) 

Disodium phosphate .. . ... . 

25% methanol . 

.... 2.84 gm 

100.0 ml 

Stock solutions (M 110, M 15) made in 25% methanol; pH values 

are for final aqueous dilution of 1:25: 

2!! 

2.5 

2.6 

2.65 

2.7 

2.75 

2.8 

2.9 

3.0 

3.05 

3.1 

3.2 

3.3 

3.45 

3.6 

3.75 

3.95 

4.1 

4.3 

4.5 

4.75 

4.95 

5.3 

5.5 

5.7 

6.0 

MIlO citric 

acid (ml) 

20.0 

19.5 

19.0 

18.5 

18.0 

17.5 

17.0 

16.5 

16.0 

15.5 

15.0 

14.5 

14.0 

13.5 

13.0 

12.5 

12.0 

11.5 

11.0 

10.5 

10.0 

9.5 

9.0 

8.5 

8.0 

MIS disodium 

phosphate (mI) 

0.0 

0.5 

1.0 

1.5 

2.0 

2.5 

3.0 

3.5 

4.0 

4.5 

5.0 

5.5 

6.0 

6.5 

7.0 

7.5 

8.0 

8.5 

9.0 

9.5 

10.0 

10.5 

11.0 

11.5 

12.0 

59 

MIlO citric MIS disodium 

2!! acid (m]) phosphate (ml) 

6.1 7.5 12.5 

6.3 7.0 13.0 

6.4 6.5 13.5 

6.5 6.0 14.0 

6.6 5.5 14.5 

6.8 5.0 15.0 

6.9 4.5 15.5 

7.0 4.0 16.0 

7.1 3.5 16.5 

7.2 3.0 17.0 

7.3 2.5 17.5 

7.4 2.0 18.0 

7.5 1.5 18.5 

7.7 1.0 19.0 

8.0 0.5 19.5 

8.3 0.0 20.0 

Reference: M.W. Newman, National Marine Fisheries Service, Oxford, Md.; 1970. 
f. Feulgen Gram (1 hr 50 min) 

Specific for gram positive bacteria. Use a positive control 

test slide. 

Procedure: 

(I) Deparaffinize and hydrate to water 

(2) Hydrolyze in 5 N HCI' (hydrochloric acid) - 30 min 

(3) Distilled water - I min (change frequently to avoid 

carry-over) 

(4) Distilled water - I min 

(5) Blot well before entering Schiff's (Feulgen) stain 

(6) Schiff's (Feulgen)l stain (refrigerated) - IS min 

(7) Distilled water - 2 changes - 1 min each 

(8) Running tap water - 2 min 

(9) Crystal violet3 - 2 min 

(10) Wash in running water 

(II) Lugol's iodine' - 2 min 

(12) Wash in running water 

(13) Decolorize in 95% ETOH until "blue" does not bleed 

off slide 

(14) Wash in running water 

(15) 0.03% picromethyl blueS - 3 min 

(16) Destain in I part 100% ETOH to 9 parts butanol - 6 

dips (change for each rack) 

(17) Butanol - I V2 min 

(18) Butanol - 3 min 

(19) Xylene 

(20) Mount 

Results: 

Nuclei - red 

Muscle - yellow 

Gram positive - blue 

Collagen - light blue 

Background - green 

Reagents: 

'5 N HCi (Lillie, 1965) 

Distilled water .. .. . . . . . . 

Concentrated HCI sp. gr. 1.19 . 

2 Schiff's (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin (stir) 

Cool to 50°C (filter optional) 

. .. 583.5 ml 

. 416.5 ml 

Add 20 ml of normal HCI (916.5 ml of distilled water and 

83.5 ml of concentrated HCI sp. gr. 1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium metabisulfite (Na2S20 s) 

Shake vigorously 

Store in a dark place; reagent takes about 2 days to turn a 

straw color 

Add charcoal (approx . 112 gm!200 m!) ; shake and filter 

Refrigerate before use 

Note : See note for Feulgen picromethyl blue stain , VI.A.3.g. 

3 Crystal violet (Brown and Brenn , 1931) 

Crystal violet . 

Sodium bicarbonate 

Distilled water 

Note: Prepare fresh before use. 

. 0.8gm 

1.0gm 

100.0 ml 

60 

, Lugol's iodine (Sanders, 1972) 

Iodine 

Potassium iodide . .. 

Distilled water 

50.03% picromethyl blue (Farley, pers. commun.) 

Methyl blue (Roboz Surgical Instrument 

1.0 gm 

.. 2.0 gm 

100.0 ml 

Cat. No.1 A240C) . . .. 0.15 gm 

Saturated aqueous picric acid (Fisher 

Scientific Cat. No. CS-311-ld) 500.0 ml 

References: Brown , lH. and L. Brenn. Bull. Johns Hopkins 

Hosp. 48: 69-73; 1931. 

(Modified by C.A. Farley, National Marine Fisheries 

Service, Oxford, Md.; 1980.) 
g. Feulgen Picromethyl Blue (FPM) (1 hr35 min) 

Specific for DNA and collagen in connective tissue. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Distilled water - 2 min 

(4) Hydrolyze in 5 N HCI' (hydrochloric acid) at room 

temp. - 30 min 

(5) Distilled water - 1 min (change frequently to reduce 

carry-over) 

(6) Distilled water - 1 min 

(7) Blot well before entering refrigerated Schiff's 

(Feulgen)l stain - 15 min 

(8) Distilled water - 2 changes - 1 min each 

(9) Running tap water - 2 min 

(10) Distilled water - 1 min 

(11) 0.03% picromethyl blue) - 3 min 

(12) Destain in 1 part 100% ETOH to 9 parts butanol ~ 6 

dips (change for each rack) 

(13) Butanol - IV2 min 

(14) Butanol - 3 min 

(15) Clear 

(16) Mount 

Results: 

DNA - red 

Nucleoli and muscle and other acidophilic 

substances - yellow 

Connective tissue - blue 

Cytoplasmic protein - green 

Reagl!l1ts: 

IS N Hel (Lillie, 1965) 

Distilled water 

Concentrated HCl sp. gr. 1.19 

2 Schiff's (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin (stir) 

Cool to 50°C (filter optional) 

, , , , 583.5 ml 

416.5 rot 

Add 20 ml of normal HCl (916.5 ml of distilled water and 

83.5 ml of concentrated HCI sp. gr. 1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium metabisulfite (Na2S20 S) 

Shake vigorously 

Store in a dark place; reagent takes about 2 days to turn a 

straw color 

Add charcoal (approx. V2 gm!200 ml); shake and filter 

Refrigerate before use 

Note: For Schiff's (Feulgen), suggest basic fuchsin from 

Fisher Scientific Co. (P-389), Certified Biological Stain 

Color Index No. 42500. It is very important to get a good 

lot of basic fuchsin. Store Schiff's in refrigerator in a 

tightly capped bottle; always use a test slide to check 

stain. When the Schiff's takes on a purple tone, it will no 

longer stain effectively; discard. Also, when making up 

solution, do not double or triple batch. Make separate 

200-ml portions for best results and store in separate 

containers. 

61 

J 0.03% picromethyl blue 

Methyl blue (Roboz Surgical Instrument 

Cat. No. lA240C ) 

Saturated aqueous picric acid (Fisher 

Scientific Cat. No. CS-31Hd) . 

..0.15gm 

500.0 ml 

Reference: Farley, CA. Natl. Cancer Inst. Monogr. 31: 541-

555; 1969. 
h. Gram (1 hr 10 min) 

Specific for gram positive and gram negative bacteria. Use 

positive and negative control test slides. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Sterling's crystal violet' - 2 min 

(4) Wash in running water - 2 min 

(5) Lugol's iodine2 - 2 min 

(6) Running water - 2 min 

(7) Decolorize in 95% ETOH until "blue" does not bleed 

off slide 

(8) Running tap water - 2 min 

(9) Safranin OJ - 2 min 

(10) Running tap water - 2 min 

(11) Picromethyl blue' or picric acidS (optional) - 3 min 

(12) Differentiate in 1 part 100% ETOH to 9 parts butanol 

- 6 dips (or until safranin 0 does not bleed off slide) 

(13) Butanol - 1112 min 

(14) Butanol - 3 min 

(15) Clear 

(16) Mount 

Results: 

Nuclei - red 

Gram positive - blue 

Gram negative - red 

Background - yellow or green, depending on counterstain 

Reagents: 

'Sterling's crystal violet (Conn et aI. , 1962) 

Crystal violet (mortared in 10 ml of 

100% ETOH) . . . . . . . . . . . . . 5.0 gm 

Aniline oil . . . . . . . . . . . . . . . . . . . . . . . . 2.0 ml 

Distilled water . . . . . . . . . . . . . 88.0 ml 

Note: Aniline oil extremely hazardous. Avoid breathing 

vapor and contact with skin. 

2Lugol's iodine (Sanders, 1972) 

Iodine . 

Potassium iodide . 

Distilled water 

JSafranin 0 (Conn et al., 1962) 

Safranin 0 dissolved in 10 ml of 

1.0gm 

. .. 2.0gm 

100.0 ml 

95% ETOH ..... .. . . . . .. . . 0.25 gm 

Distilled water .. . . . .. . .. . 

4().03% picromethyl blue (Farley, pers. commun.) 

Methyl blue (Roboz Surgical Instrument 

. 90.0ml 

Cat. No. lA240C ) . . . . ....... . .. . .. . 0.15 gm 

Saturated aqueous picric acid (Fisher 

Scientific Cat. No. CS-311-1 d) ......... . ... 500.0 ml 

SPicric acid (saturated aqueous) 

Distilled water 

Picric acid ... ... . . . . . . .. . 

.. . . . . .. .. .... 20.0 mJ 

. ..... 2.0gm 

62 

References: Brown , J.H. and L. Brenn. Bull. Johns Hopkins 

Hosp. 48: 69-73 ; 1931. 

(Modified by C.A. Farley, National Marine Fisheries 

Service , Oxford, Md. ; 1980.) 
I. GrocoH'. Methenamine Silver (GMS) (3 hr 15 

min) 

Specific for fungi. Use a positive control test slide and acidcleaned glassware. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Oxidize in 4% chromic acid 1 - 1 hr 

(4) Wash in tap water - few seconds 

(5) 1% sodium bisulfite2 - 1 min (to remove any residual 

chromic acid) 

(6) Wash in running water - 5-10 min 

(7) Rinse in distilled water - 3-4 changes 

(8) Place in freshly mixed methenamine silver nitrate3 

working solution in incubator or water bath at 58°-

60°C for 60 min or until section turns yellowishbrown 

(9) Rinse in distilled water - 6 changes 

(10) Tone in 0.1 % gold chloride4 - 2-5 min 

(11) Rinse in distilled water 

(12) Remove unreduced silver with 2% sodium 

thiosulfate (hypo)5 - 2-5 min 

(13) Wash thoroughly in tap water 

(14) Counterstain in 0.2% light green" - 30-45 sec 

(15) Dehydrate 

(16) Clear 

(17) Mount 

Results: 

Fungi - sharply delineated in black 

Mucin - taupe to dark gray 

Inner parts of mycelia and hyphae - old rose 

Background - pale green 

Reagents: 

14% chromic acid 

Chromic acid .. ... . . . 

Distilled water 

21 % sodium bisulfite 

Sodium bisulfite 

Distilled water 

3Methenamine silver nitrate working solution 

Prepare the following solutions: 

(a) 5% silver nitrate 

Silver nitrate 

Distilled water 

(b) 3% methenamine 

Methenamine 

Distilled water 

(c) 5% borax 

Borax .......... . . 

Distilled water. . ....... . 

(d) Methenamine silver nitrate stock solution 

.. . 4.0gm 

100.0 ml 

1.0gm 

100.0 ml 

. 0.25 gm 

5.0ml 

3.0gm 

100.0 ml 

5.0gm 

100.0 ml 

5% silver nitrate (a) 5.0 ml 

3% methenamine (b) . 100.0 ml 

Note: A white precipitate forms in the methenamine 

silver nitrate stock solution, but immediately dissolves on shaking. Clear solution remains usable 

for months. Store in refrigerator. 

63 

Combine for working solution as follows: 

Methenamine silver nitrate stock (d) 

Distilled water . 

5% borax (c) 

Note: Make working solution fresh. 

40.1 % gold chloride 

1 % gold chloride ... . .. . . 

Distilled water . 

Note: This solution may be 'Used repeatedly. 

52% sodium thiosulfate (hypo) 

Sodium thiosulfate ...... . . 

Distilled water ...... . 

"0.2% light green 

Light green .. ... . . .. . 

Distilled water 

25.0ml 

25.0 ml 

2.0ml 

10.0 ml 

9O.0ml 

2.0gm 

100.0 ml 

.. 0.2gm 

100.0 ml 

Note: Acid-cleaned glassware for preparation of solutions in 

the above silver procedure is recommended, as well as 

acid-cleaned coplin jars. The silver will precipitate on the 

slide very easily. Keep all solutions away from metal, and 

be sure to coat forceps with paraffin before immersing in 

silver solution . 

Reference: Luna, L.G., ed. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. 

McGraw-Hili, New York; 1968. 
j. Harris' Hematoxylin and Eosin (HHE1

) (Progressive Stain) (1 hr 30 min) 

General tissue stain used with tissues fixed in Helly's or 

Zenker's. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water and remove mercury crystals, if 

necessary (Vl.A.l.c.) 

(3) Harris' hematoxylin' - 2 min 

(4) Distilled water - 6 dips 

(5) Running tap water - 5 min 

(6) Distilled water - 6 dips 

(7) 95% ETOH - 2 min 

(8) Eosin bluish2 - 20 sec-2 min (may vary with tissue 

and fixative used) 

(9) Dehydrate 

(10) Clear 

(11) Mount 

Results: 

Basophilic substances - blue 

Acidophilic substances - pink 

Reagents: 

'Harris' hematoxylin 

Harris' hematoxylin (VWR Scientific Cat. 

No. AL33050-3e ) .. 

Glacial acetic acid ... . . . . . 

2Eosin bluish 

Eosin bluish (Fisher Scientific Cat. No. 

E-514d) 

95% ETOH . 

Note: Age eosin bluish 1 month before using. 

. 200.0 ml 

14.0 ml 

1.0gm 

100.0 ml 

Reference: Johnson, P.T. Histology of the blue crab, Callinectes 

sapidus. A model for the Decapoda. Praeger, New York; 

1980. 

64 
k. Harris' Hematoxylin and Eosin (HHE2) (Regressive Stain) (1 hr 10 min) 

General tissue stain for tissues fixed in Davidson's, formalin, 

and Bouin's. 

Procedure: 

(I) Deparaffinize 

(2) Hydrate to water and remove mercury, if necessary 

(VI.A.l.c.) 

(3) Harris' hematoxylin l - 8 min 

(4) Distil.led water - 2 min (use fresh distilled water for 

each rack) 

(5) Acid alcohoF - 1 dip 

(6) Running tap water - 5 min (tissue should " blue" 

here) 

(7) Distilled water - 3 min 

(8) 95% ETOH - 3 min 

(9) Eosin yJ - 3 min 

(10) 95% ETOH - 6 dips 

(11) Dehydrate 

(12) Clear 

(13) Mount 

Results: 

Basophilic substances - blue 

Acidophilic substances - pink 

Reagents: 

lHarris' hematoxylin 

Harris' hematoxylin (VWR Scientific 

Cal. No. ALJ3050-3e) . 

Glacial acetic acid. 

2 Acid alcohol 

70% ETOH. 

Concentrated hydrochloric acid (HCI) 

. .... 200.0 ml 

14.0 ml 

. 99.0 ml 

sp. gr. 1.19 . . .. .... . . . 1.0 ml 

JEosin y 

Eosin y 

Distilled water 

100% ETOH . 

Just before use, add concentrated 

. 0.2gm 

75.0 ml 

25.0 ml 

HCI 1 drop!200 ml 

Note : If precipitate does not dissolve on stirring, eosin may 

stain too orange. 

Reference: Johnson, P.T. Histology of the blue crab, Callinectes sapidus A model for the Decapoda. Praeger, New 

York; 1980. 

65 
I. Heidenhain's Iron Hematoxylin and Eosin 

(IHE) (1 hr 30 min) 

Specific for good nuclear detail. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Place in Lang's mordant' - 10 min 

(4) Wash in running water - 5 min 

(5) Dip in distilled water 

(6) Hematein2 or hematoxylin3 - 10 min 

(7) Dip in distilled water 

(8) Destain in picric acid' until only nuclei remain stained 

- 10-20 min (check with microscope) 

(9) Wash in running tap water - 5 min 

(10) Dehydrate in alcohols through 95% ETOH as usual 

(11) Counterstain in eosin bluishs - 20 sec-2 min 

(12) Dehydrate 

(13) Clear 

(14) Mount 

Results: 

Nuclei - black 

Muscle - pink 

Background - gray 

Reagents: 

'Lang's mordant stock solution (Conn et aI. , 1962) 

Distilled water 1000.0 ml 

Glacial acetic acid . 10.0 ml 

Sulfuric acid. . . . . . . . . . . . . . 1.2 ml 

Ferric ammonium sulfate . . . . . . . . . . 30.0 gm 

Lang's mordant working solution 

Lang's mordant stock. 

Dimethyl sulfoxide (DMSO) (Mitchell, 

1966) 

... 200.0 ml 

... . 20.0 ml 

2Hematein stock solution (Farley, pers. commun.) 

100% ETOH 200.0 ml 

Hematein ... . . .. .. . .. ... 20.0 gm 

Hematein working solution (Mallory, 1968) 

Hematein stock 

Distilled water 

DMSO 

3Hematoxylin stock solution (Lillie, 1965) 

10.0 ml 

190.0ml 

20.0 ml 

100% ETOH . . 100.0 ml 

Hematoxylin 10.0 gm 

Note: Allow hematoxylin stock solution to age 4-6 weeks 

before use. 

Hematoxylin working solution 

Distilled water 

Hematoxylin stock 

DMSO 

190.0 ml 

10.0 ml 

20.0ml 

66 

Note: Both hematein and hematoxylin are included for the 

above procedure for Heidenhain's iron hematoxylin. The 

hematoxylin is purer but takes 4-6 weeks to ripen, while 

hematein may be used immediately. Do not contaminate 

hematoxylin with mordant; stain will become ineffective. 

When using DMSO, avoid contact with skin. 

'Picric acid (saturated aqueous) 

Distilled water .............. . . 

Picric acid ...... . . 

SEosin bluish 

Eosin bluish (Fisher Scientific Cat. No. 

E-514d) 

95% ETOH 

.20.0ml 

... 2.0gm 

1.0gm 

100.0 ml 

Note: Allow eosin bluish to age 1 month before use. 

References: Conn, H.J. , et al. Staining Procedures Used by the 

Biological Stain Commission. 2nd ed. The Williams & 

Wilkins Co., Baltimore; 1962. 

(Modified by C.A. Farley, National Marine Fisheries 

Service, Oxford , Md .; 1979.) 
m. Mallory'. Hematein (1 hr' 

Specific for copper and other metals. Use a positive control 

test slide. 

Procedure: 

(I) Deparaffinize 

(2) Hydrate to water. but do not permit slides to remain 

in water 

(3) Continue immediately 

(4) Hematein ' with dimethyl sulfoxide (DMSO) - 5 min 

(5) Running tap water to "blue" slides - 5 min 

(6) Dip in distilled water 

(7) 1 % safranin 0 2 - 2 min 

(8) Running tap water - 2 min 

(9) Destain in 1 part 100% ETOH to 9 parts butanol -

1-6 dips 

(10) Butanol - 2 min 

(II) Butanol - 5 min 

(12) Clear 

(13) Mount 

Results: 

Background - bright red 

Special contaminants. e.g. copper - deep clear blue 

Reagents: 

'0.5% hematein stock solution (Farley. pers. 

commun.) 

Hematein . 

Distilled water ....... . 

Hematein working solution 

1.0gm 

200.0 ml 

0.5% hematein stock . . . . . . . . . 10 parts 

DMSO 1 part 

Note: Be careful not to contaminate hematein; a dark color 

change will indicate the stain is no longer usable. Make up 

immediately prior to use. It will not keep overnight after 

DMSO is added. Avoid contact of DMSO with skin. 

21 % safranin 0 (Conn et al. . 1962) 

Safranin O .. 

Distilled water 

1.0gm 

100.0 ml 

References: Lillie. R.D. Histopathologic Technic and Practical Histochemistry. 3rd ed. McGraw-Hill. New York; 1965. 

(Modified by C.A. Farley. National Marine Fisheries 

Service. Oxford. Md.; 1980.) 

67 
n. Mallory's Trichrome (Modified) (1 hr) 

Specific for coHagen in connective tissue. Use a positive 

control test slide. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) 0.5% acid fuchsin 1 - 7 min 

(4) Dip twice in tap water 

(5) Blot slide well 

(6) Aniline blue-orange G2 - 15 min 

(7) Rinse in tap water - 2 quick dips 

(8) Transfer to several changes of 95% ETOH (quickly) 

(9) Dehydrate (quickly, because slides can lose too much 

aniline blue in this process) 

(10) Clear 

(II) Mount 

Results: 

Nuclei - red 

Collagen fibers - intense blue (also reticulum, pale blue) 

Elastic fibers - pale pink, pale yeHow, or unstained 

Muscle - red-purple to blue (depending on specimen) 

Erythrocytes and myelin - yel.low 

Reagents: 

10.5% acid fuchsin 

Acid fuchsin ......... . . 

Distilled water 

. 0.5 gm 

100.0 ml 

Note: Acid fuchsin stain gives best results after aging a few 

days. 

2Aniline blue-orange G (best for tissues fixed in Davidson's 

formalin) 

Dissolve in this order: 

Aniline blue ....... . . . 

Orange G 

Phosphomolybdic acid 

Distilled water ......... . . 

. ... 0.5gm 

... . .. . 2.0gm 

1.0gm 

100.0 ml 

Aniline blue-orange G (best for Helly's or Zenker's fixed 

tissue) (Johnson, 1980) 

Aniline blue .......... . . 

Orange G 

Phosphotungsic acid ...... . 

Distilled water ..... . . 

Glacial acetic acid ...... . . . . 

. ... 0.5gm 

... 2.0gm 

.... 2.0gm 

100.0 ml 

. . .. 2 drops 

References: Pauley, G. B. 1. Invertebr. Pathol. 9: 268-269; 1967. 

(Modified by P.T. Johnson. Histology of the Blue Crab, 

Callinecles sapidus. A Model for the Decapoda. Praeger, 

New York; 1980.) 

68 
o. Malt Periodic Acid Schiff (MPAS) (2 hr 20 

min) 

Specific for PAS positive substances other than glycogen. 

Procedure: 

(I) Deparaffinize 

(2) Hydrate to water 

(3) Immerse in malt' (diastase) - I hr at 40°C 

(4) Wash well in running tap water - 10 min 

(5) Distilled water 

(6) Oxidize in 1 % periodic acid2 at room temp. - 5 min 

(7) Wash in running tap water - 5 min 

(8) Blot well and place in refrigerated SchiffsJ (Feulgen) 

reagent - 5 min 

(9) Wash well - 10 min 

(10) Hematoxylin· of choice 

(11) "Blue" in tap water - 5 min 

(12) Dehydrate 

(13) Clear 

(14) Mount 

Results: 

Glycogen is digested 

Other PAS positive substances - pink-magenta 

Basophilic substances - blue-black 

Reagents: 

'Malt for MPAS (Lillie, 1965) 

Diastase malt. 

Buffer, pH 5.5-6.0 . 

Distilled water 

Buffer for malt (Lillie, 1965) 

.. 0.2gm 

. . . 20.0ml 

180.0 ml 

Boil distilled water . . 1000.0 ml 

Sodium phosphate monobasic 

(NaH2P04.H20) . . 1.97 gm 

Sodium phosphate dibasic (Na2HP04) . . . . . . 0.28 gm 

Note: pH 5.5 to 6.0; check pH meter. Phosphate buffer 

same as Sorenson's. Add 1 thymol crystal when cool to 

prevent mold , and refrigerate. 

21% periodic acid (Drury and Wallington, 1967) 

Periodic acid . 

Distilled water 

3Schiffs (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin and stir 

Cool to 50°C (filter optional) 

I.Ogm 

100.0 ml 

Add 20 ml of normal hydrochloric acid (HC!) (916.5 ml of 

distilled water and 83.5 ml of concentrated HCl sp. gr. 

1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium meta bisulfite (Na2SPs) 

Shake vigorously. Store in a dark place ; reagent takes 

about 2 days to turn a straw color 

Add charcoal (approx. V2 gm1200 m!); shake and filter 

Refrigerate before use 

Note: See note for Feulgeo picromethyl blue stain, VI.A.3.g. 

69 

4Harris' hematoxylin 

Harris' hematoxylin (VWR Scientific 

Cat. No. AL33050-3e) . . .. . . . .. . 200.0 ml 

Reference: Lillie, R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed. McGraw-Hill, New York ; 1965. 
p. Malt Periodic Acid Schiff-Alcian Blue (MPASAB) (5 hr) 

Specific for differentiation of acid and neutral mucopolysaccharides. Use a positive control test slide. 

Procedure: 

(I) Deparaffinize 

(2) Hydrate to water and remove mercury, if present 

(see VI.A.1.c.) 

(3) Immerse in malt I (diastase) - 1 hr at 40°C 

(4) Wash well in running tap water 

(5) Oxidize in 1 % periodic acid2 at room temp. - 5 min 

(6) Wash in running water - 5 min 

(7) Blot well and place in refrigerated SchiffsJ (Feulgen) 

reagent - 5 min 

(8) Wash well in running water - 10 min 

(9) Harris' hematoxylin· - 2 min 

. (10) "Blue" in running tap water - 5 min 

(11) Rinse in distilled water 

(12) Alcian bluel - 30 min 

(13) Rinse briefly in running tap water 

(14) 0.5% alcoholic ammonium hydroxide6 - 2 hr 

(IS) Rinse in running tap water 

(16) Dip in distilled water 

(17) Dehydrate 

(18) Clear 

(19) Mount 

Results: 

Acid mucopolysaccharides - bright intense blue 

Neutral mucopolysaccharides - magenta 

PAS positive tissue - magenta 

Nuclei - blue 

Reagents: 

IMalt (Lillie , 1965) 

Diastase malt ... 

Buffer, pH 5.5-6.0 .. 

Distilled water 

Buffer for malt (Lillie, 1965) 

.0.2gm 

.. 20.0ml 

180.0 ml 

Boil distilled water . . 1000.0 ml 

Sodium phosphate monobasic 

(NaH2PO •. H20) . 1.97 gm 

Sodium phosphate dibasic (Na2HP04) . .... .. . 0.28 gm 

Note: pH 5.5 to 6.0; check pH meter. Phosphate buffer 

same as Sorenson's. Add 1 thymol crystal when cool to 

prevent mold, and refrigerate. 

21 % periodic acid (Drury and Wallington, 1967) 

Periodic acid. 

Distilled water 

JSchiffs (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin and stir 

Cool to 50°C (filter optional) 

1.0gm 

100.0 ml 

Add 20 ml of normal hydrochloric acid (HCl) (916.5 ml of 

distilled water and 83.5 ml of concentrated HCI sp. gr. 

1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium meta bisulfite (Na2S20 s) 

70 

Shake vigorously 

Store in a dark place; reagent takes about 2 days to turn a 

straw color 

Add charcoal (approx. 112 gm!200 ml); shake and filter 

Refrigerate before use 

Note: See note for Feulgen picromethyl blue stain, VI.A. 3.g. 

'Harris' hematoxylin 

Harris' hematoxylin (VWR Scientific Cat. 

No. AL330S0-3e) . 200.0 ml 

SO.l % alcian blue in om M HCI (pH 2.2) - Helly's or Zenker's 

fixative (Johnsun, 1980) 

Alcian blue 8GX . 

1 N HCI 

Distilled water 

1.0 gm 

10.0 ml 

.. 990.0 ml 

0.1 % alcian blue in 3% acetic acid - Davidson's or formalinfixed tissue (Johnson, 1980) 

Alcian blue 8GX . . .. . .. .. . 

Distilled water 

Glacial acetic acid . 

. .. . 0.1 gm 

... .. . . 97.0 ml 

.3.0ml 

Note: Check expiration date of alcian blue stain. 

00.5% alcoholic ammonium hydroxide (Johnson , 1980) 

Ammonium hydroxide . . . . . 3.0 ml 

95% ETOH . . . . . . . . . . . . 597.0 ml 

References: Lillie, R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed . McGraw-Hill, New York; 1965. 

Pearse, A.G.E. Histochemistry, Theoretical and Applied. 

2nd ed. Little, Brown and Co., Boston; 1960. 

(Modified by P.T. Johnson. Histology of the Blue Crab, 

Callinectes sapidus. A Model for the Decapoda. Praeger, 

New York; 1980.) 
q. 011 Red 0 (ORO) (10 min) 

Specific for fat in frozen sections. 

Procedure: 

(1) Frozen sections, air-dried or fixed (if fixed, wash Qut 

fixative) 

(2) Rinse in dilute 49.5% isopropyl alcohoP 

(3) Oil red 0 2 - 5 min 

(4) Rinse in dilute 49.5% isopropyl alcohol 

(5) Rinse in distilled water 

(6) Mount in glycerin jelly 

Results: 

Fat - bright orange-red 

Reagents: 

149.5% isopropyl alcohol 

99% isopropyl alcohol 

Distilled water .. 

. . . . . .. . . . 50.0 ml 

50.0 ml 

JOil red 0 stock solution 

Supersaturated: 

Oil red O . 

99% isopropyl alcohol. 

Oil red 0 working solution 

Saturated: 

Oil red 0 stock .. ... . . .. ... . . ..... .. . . . . 

Distilled water . 

. 0.5 gm 

125.0 ml 

12.0 ml 

. . 8.0ml 

Mix. allow solution to stand for 5 min, and filter. 

Note: Use working solution until precipitate begins to form. 

Reference: Thompson, S.W. Selected Histochemical and Histopathological Methods. Charles C. Thomas Publ., Springfield , Ill.; 1966. 

71 
r. Periodic Acid Schiff (PAS1) (1 hr 15 min) 

Specific for glycogen, fungi, and neutral mucopolysaccharides. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Oxidize in 1% periodic acid l at room temp. - 5 min 

(4) Wash in running tap water - 5 min 

(5) Blot well and place in refrigerated Schiff's2 

(Feulgen) reagent - 5 min 

(6) Rinse immediately in running tap water - 10 min 

(7) Distilled water - 6 dips 

(8) Weigert's hematoxylinJ - 30 sec 

(9) Running water - 6 dips 

(10) Distilled water - 6 dips 

(11) Dehydrate 

(12) Clear 

(13) Mount 

Results: 

Glycogen - magenta 

PAS positive substances - red-pink 

Basophilic substances - blue-black 

Reagents: 

11 % periodic acid (Drury and Wallington, 1967) 

Periodic acid .. 

Distilled water 

2Schiff's (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin (stir) 

Cool to 50°C (filter optional) 

Add 20 ml of normal hydrochloric acid (HC!) 

(916.5 ml of distilled water and 83.5 ml of 

concentrated HCI sp. gr. 1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium metabisulfite (Na2S20 s) 

Shake vigorously 

1.0gm 

100.0 ml 

Store in dark place; reagent takes about 2 days to turn a 

straw color 

Add charcoal (approx. 112 gm!200 ml); shake and filter 

Refrigerate before use 

Note: See note for Feulgen picromethyl blue stain, VI. 

A.3.g. 

JWeigert's hematoxylin for PASI (Lillie, 1965) 

Solution A: Hematoxylin . 

95% ETOH. . ... .. . 

Solution B: Ferric chloride 

(FeCiJ .6H20) . 

Distilled water 

HC!. 

1.0 gm 

100.0 ml 

. 2.5gm 

.99.0ml 

. .. . 1.0 ml 

Mix equal parts of solutions A and B immediately before use. 

Note: When using Weigert's hematoxylin in this procedure, 

work quickly: rinse in running water to avoid overstaining 

and precipitation of stain on slide; acid-cleaned slides will 

prevent stain precipitation. This stain will not keep; make 

fresh each time. 

72 

Reference: Lillie, R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed. McGraw-Hill, New York; 1965. 
s. Periodic Acid Schiff (PAS2) (Heidenhain's 

Hematoxylin Counterstain) (1 hr 45 min) 

Specific for nuclear detail, fungi, glycogen, and neutral 

mucopolysaccharides. 

Procedure: 

(I) Deparaffinize 

(2) Hydrate to water 

(3) Oxidize in 1 % periodic acid l at room temp. - 5 min 

(4) Wash in running tap water - 5 min 

(5) Blot well and place in refrigerated Schiffs2 (Feulgen) 

reagent - 5 min 

(6) Go directly to running tap water - 10 min 

(7) Distilled water - 6 dips 

(8) Mordant in 3% ferric ammonium sulfate3 - 5 min 

(9) Wash in running tap water - 2 min 

(10) Heidenhain's hematoxylin' - 2-5 min, according to 

intensity desired, at room temp. 

(11) Differentiate in picric acid) until only nuclei remain 

stained - 10-20 min (check with microscope) 

(12) Wash in running tap water - 2 min 

(13) Dehydrate 

(14) Clear 

(IS) Mount 

Results: 

PAS positive substances - red-pink 

Glycogen, collagen, reticulum, mucin, and neutral mucopolysaccharides - red-pink 

Basophilic substances - blue to black 

Reagents: 

11 % periodic acid (Drury and Wallington, 1967) 

Periodic acid .. 

Distilled water 

2Schiff's (Feulgen) (Sanders, 1972) 

Boil 200 ml of distilled water 

Add 1.0 gm of basic fuchsin (stir) 

Cool to 50°C (filter optional) 

1.0gm 

100.0 ml 

Add 20 ml of normal hydrochloric acid (HCI) (916.5 ml of 

distilled water and 83.5 ml of concentrated HCI sp. gr. 

1.19) 

Cool to 25°C (room temp.) 

Add 1.0 gm of sodium meta bisulfite (Na2S20 s) 

Shake vigorously 

Store in a dark place; reagent takes about 2 days to turn a 

straw color 

Add charcoal (approx. 112 gm!200 ml); shake and filter 

Refrigerate before use 

Note: See note for Feulgen picromethyl blue stain, VI.A.3.g. 

33% ferric ammonium sulfate 

Ferric ammonium sulfate 

Distilled water 

'Heidenhain's hematoxylin (Mitchell, 1966) 

.. 3.0gm 

100.0 ml 

0.5% hematein stock (VI.A.3.m.) 10 parts 

Dimethyl sulfoxide (DMSO) . . I part 

Note: Do not contaminate hematoxylin with mordant; stain 

will become ineffective. Avoid contact of DMSO with 

skin. 

73 

SPicric acid (saturated aqueous) 

Distilled water . . . . ... . . . . . .. 20.0ml 

Picric acid . . . .. . 2.0 gm 

References: Lillie, R.D. Histopathologic Technic and Practical 

Histochemistry. 3rd ed. McGraw-Hili, New York; 1965. 

Conn, H.J., et al. Staining Procedures Used by the Biological 

Stain Commission. 2nd ed. The Williams & Wilkins Co., 

Baltimore, Md.; 1962. 
t. Pert.' Prussia" Blue (2 hr) 

Specific for ferric iron. Use a positive control test slide and 

acid-cleaned glassware. 

Procedure: 

(l) Deparaffinize 

(2) Hydrate to water 

(3) Expose sections to a fresh mixture of equal parts of 

2% potassium 1 or sodium ferrocyanide2 and 2% 

hydrochloric acid (HCl)3 - 30-60 min (if longer 

period is used, change solution after first 30 min) 

(4) Wash in distilled water 

(5) Counterstain nuclei with 1 % neutral red' - 3 min 

(6) Wash in running tap water 

(7) Dehydrate 

(8) Clear 

(9) Mount in Canada balsam (a neutral mounting medium) 

Results: 

Ferric iron - Prussian blue to green 

Nuclei - red 

Background - pink 

Reagents: 

12% potassium ferrocyanide 

Potassium ferrocyanide . 

Distilled water 

22% sodium ferrocyanide 

Sodium ferrocyanide ....... . 

Distilled water 

32% HCl 

. Concentrated HCl sp. gr. 1.19 

Distilled water .......... . 

41% neutral red 

Neutral red ....... . . . . ... . 

Distilled water 

2.0gm 

100.0 ml 

2.0gm 

100.0 ml 

.... . ... .... 2.0ml 

. . .... . . . .. ... 98.0 ml 

1.0 gm 

100.0 ml 

Note: Keep solutions away from metal to avoid contamination. 

Acid fixatives interfere with iron. Versenate decalcification 

interferes with results (Drury and Wallington, 1967). 

Reference: Drury, R.A.B. and S.A. Wallington. Carleton's 

Histological Technique. 4th ed. Oxford Univ. Press; 1967. 

74 
u. Sudan Black B (SBB) (allow 2 days) 

Specific for phospholipids. Use duplicate slides. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Post-chrome in 3% potassium dichromate 1 - 1-2 hr 

(1 hr 15 min for Helly's fixative) 

(4) Wash in running water - 10 min 

(5) Dehydrate to 70% ETOH and stain in 0.3% SBB2 at 

room temp. - 16 hr (time varies for specific tissue) 

(6) Rinse in 70% ETOH and wash-5-1O min (one set of 

test slides goes into acetone for 7V2 hr and is washed 

in running water for 5-10 min, counterstained, and 

mounted for comparison response of results) 

(7) Counterstain in 1 % neutral red3 - 1 min 

(8) Wash in running water - 5-10 min 

(9) Dip in distilled water 

(10) Mount in glycerin jelly 

Results: 

Phospholipids - black-blue, black 

Background - red 

Bound lipids and remaining neutral lipids - black, bluegray, or brownish 

Other tissue components - pink to red or unstained 

Following the acetone extraction, lipid components are 

unstained 

Reagents: 

13% potassium dichromate 

Potassium dichromate. 

Distilled water 

20.3% SBB 

SBB . 

70% ethyl alcohol ... 

31 % neu tral red 

Neutral red 

Distilled water 

. . ... . ... .. 3.0gm 

100.0 ml 

. 0.3gm 

100.0 ml 

l.Ogm 

100.0 ml 

References: Pearse, A.G.E. Histochemistry, Theoretical and 

Applied. 2nd ed. Little, Brown and Co., Boston; 1960. 

(Modified by P.T. Johnson. Histology of the Blue Crab, 

Callinectes sapidus. A Model for the Decapoda. Praeger, 

New York; 1980.) 

75 
v. Toluidine Blue 0 (TBO) (fresh blood, 30 min) 

Specific for hemocytes. 

Procedure: 

(1) Prepare slide with fresh blood 

(2) Fix in 1 % glutaraldehyde/4% formaldehyde (1 G4F) 

fixative (made with ambient sea water) - 2 min 

(3) Wash in running tap water - 1 min 

(4) Distilled water - 6 dips 

(5) 0.5% toluidine blue 01 - 5 min 

(6) Running tap water - 6 dips 

(7) Destain in 1 part 100% ETOH to 9 parts butanol - 6 

dips 

(8) Butanol - 2 min 

(9) Butanol - 4 min 

(10) Clear 

(11) Mount 

Results: 

Nuclei - blue 

Granules - purple or blue 

RBCs - green 

Reagent: 

10.5% toluidine blue 0 

Toluidine blue 0 . 

Borax . 

Distilled water 

.. . . 0.5 gm 

. . . O.Sgm 

100.0 ml 

Heat gently to dissolve the stain, then filter solution. 

Note: Stable for several months. 

References: Trump, B.F., et al. 1. Ultrastruct. 5: 343-348; 1961. 

(Modified by c.A. Farley, National Marine Fisheries Service, Oxford, Md.; 1980.) 

76 
w. Von K6ssa (2 hr) 

Specific for calcium. Use a positive control test slide and 

acid-cleaned glassware. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Place slides in 5% silver nitrate' - 30-60 min, exposed 

to direct sunlight 

(4) Rinse in distilled water - 2-3 changes 

(5) 5% sodium thiosulfate2 - 2-3 min 

(6) Wash well in distilled water (2 dips each in 2 changes) 

(7) Counterstain in nuclear fast redJ (Kernechtrot) - 5 

min 

(8) Wash well in distilled water - 6 dips 

(9) Dehydrate 

(10) Clear 

(11) Mount 

Results: 

Calcium salts - black 

Nuclei - red 

Cytoplasm - pink to rose 

Reagents: 

'5% silver nitrate 

Silver nitrate . 

Distilled water 

25% sodium thiosulfate 

Sodium thiosulfate 

Distilled water . . .. . . . . 

'Nuclear fast red (Kernechtrota) 

5% aluminum sulfate (AI2(S04h): 

AI 2(S04h . . 

Distilled water 

0.1 % nuclear fast red: 

Kernechtrot nuclear fast red. 

5% AI2(S04)J . 

Note: Refrigerate; good till loses color. 

5.0gm 

100.0 ml 

5.0gm 

100.0 ml 

25.0 gm 

500.0 ml 

0.5gm 

500.0 ml 

Note: For best results, stain on a sunny day. Hi-intensity lamps 

do not work nearly as well as natural sunlight. 

Reference: Sanders, B.1. Animal Histology Procedures of the 

Pathological Technology Section of the National Cancer 

Institute. Government Printing Office, Washington, D.C.; 

1972. 

77 
L Wilder's Reticulum (WR) (1 hr) 

Specific for reticulum (single slide manipulation). Use acidcleaned glassware. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Oxidize in 10% phosphomolybdic acidl - 1 min 

(4) Rinse well in running water 

(5) Sensitize in 1 % uranium nitrate2 - 1 min 

(6) Rinse in distilled water - 10-20 sec 

(7) Ammoniacal silveil -1 min (change solution frequently) 

(8) Dip quickly in 95% ETOH and go straight to reducing 

solution· - 1 min (change solution frequently) 

(9) Rinse well in distilled water 

(10) Tone in gold chloride~ (until section is lavender) - 1 

min 

(11) Rinse well in distilled water 

(12) 5% sodium thiosulfate6 - 1 min 

(13) Wash well in tap water 

(14) Distilled water - quick dip 

(15) Counterstain as desired (usually use nuclear fast red) 

(16) Nuclearfast red7 (Kernechtrot) - 1-4 min (depending 

on age of stain) 

(17) Dip in distilled water - 6 dips 

(18) Dehydrate 

(19) Clear 

(20) Mount 

Results: 

Reticulum fibers - black 

Collagen - rose 

Other tissue elements - red 

Reagents: 

110% phosphomolybdic acid 

Phosphomolybdic acid 

Distilled water 

21 % uranium nitrate 

Uranium nitrate. 

Distilled water 

J Ammoniacal silver 

10.2% silver nitrate: 

Silver nitrate 

Distilled water 

or 

Silver nitrate 

Distilled water 

3.1 % sodium hydroxide: 

Sodium hydroxide . . .. . . . .. . . 

Distilled water . .. . ... . . 

or 

Sodium hydroxide. 

Distilled water .. .. ... . 

10.0 gm 

100.0 ml 

1.0 gm 

100.0 ml 

1O.2gm 

100.0 ml 

1.02 gm 

.. . . . .. . . . 10.0 ml 

. 3.1 gm 

100.0 ml 

0.31 gm 

10.0 ml 

To 5 ml of 10.2% silver nitrate add 28% ammonium 

hydroxide (concentrated ammonium hydroxide) until precipitate which forms is almost dissolved. Add 5 ml of 3.1 % 

78 

sodium hydroxide and barely dissolve the resulting precipitate with a few drops of ammonium hydroxide. Make 

solution up to 50 ml with distilled water. Use at once. 

4Reducing solution 

Distilled water 

40% neutral formalin 

50.0 ml 

. 0.5ml 

1 % uranium nitrate . 1.5 ml 

Note: Make fresh and just before use add calcium carbonate 

in excess, or add one marble chip. 

51 % gold chloride stock solution 

Gold chloride .. 

Distilled water 

0.2% gold chloride working solution 

1 % gold chloride stock. 

Distilled water 

65% sodium thiosulfate 

Sodium thiosulfate 

Distilled water 

7Nuclear fast red (Kernechtrota) 

5% aluminum sulfate (AI2(S04)3): 

AI2(S04h 

Distilled water .. . . .. . . 

0.1 % nuclear fast red: 

Kernechtrot nuclear fast red . 

5% AI2(S04h . 

Note: Refrigerate; good till loses color. 

15 grains/ glass vial 

100.0 ml 

10.0 ml 

4O.0ml 

5.0gm 

100.0 ml 

25.0 gm 

500.0 ml 

0.5gm 

500.0 ml 

Note: Use acid-cleaned slides and glassware for preparation of 

reagents and for staining. Silver will precipitate on the 

slide very easily, and frequent changes of the solutions are 

recommended. Keep metal away from all solutions, and 

be sure to coat forceps with paraffin before immersing in 

silver solution. 

Reference: Luna, L.G., ed. 1968. Manual of Histologic Staining 

Methods of the Armed Forces Institute of Pathology. 3rd 

ed. McGraw-Hili, New York; 1968. 
y. Ziehl and Harris' Hematoxylin (ZHH) (1 hr 45 

min) 

Specific for acid-fast substances (e.g. detection of acid-fast 

organisms, haplosporidan spores). Use a positive control test 

slide. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Ziehl's fuchsin' - 15 min 

(4) Distilled water - 6 dips 

(5) Destain in 0.1 N sulfuric acid2 - 10-15 min (time 

varies with specific tissue) 

(6) Running tap water - 10 min 

(7) Harris' hematoxylin3 - 2 min 

(8) Distilled water - 6 dips 

(9) Lithium carbonate saturated solution4 - 2 min (not 

necessary if running tap water is slightly alkaline) 

(10) Running tap water - 5 min 

(11) Dehydrate 

(12) Clear 

(13) Mount 

Results: 

Basophilic substances - blue 

Acid-fast substances - bright red, including mature 

haplosporidan sporoplasms, lipofuscins, and acid-fast 

bacteria 

Reagents: 

'Ziehl's fuchsin (Gray, 1954) 

Basic fuchsin (magenta) (Fisher Scientific Cat. 

No. P-38gd) 1.0 gm 

Phenol (liquefied). . . . . . . . . . . . . . . . 5.0 ml 

90% ETOH . 10.0 ml 

Distilled water 100.0 ml 

Grind the fuchsin with phenol in a mortar. When dissolved, 

add the alcohol in 10 successive lots while grinding. Then 

wash stain from mortar with ten 10-ml washings of distilled 

water. Save and filter accumulated washings. 

20.1 N sulfuric acid (H2S04) 

Distilled water 

H2S04 sp. gr. 1.84 

3Harris' hematoxylin 

Harris' hematoxylin (VWR Scientific 

997.2 ml 

.. ... 2.8ml 

Cat. No. AL33050-3ee

) . . . . . . . . . . . . . . . 200.0 ml 

Glacial acetic acid . . . . . . . . . .. 14.0 ml 

4Lithium carbonate saturated solution 

Lithium carbonate. 1.0 gm 

Distilled water . . .. . . .. . . . . . . . . ... . . . . .. 78.0 ml 

Reference: Farley, C.A. 1. Invertebr. Pathol. 7: 144-147; 1965. 

79 
z. Ziehl and Methylene Blue (1 hr 30 min) 

Specific for acid-fast bacteria (AFB). Use a positive control 

test slide. 

Procedure: 

(1) Deparaffinize 

(2) Hydrate to water 

(3) Ziehl's fuchsin! - 15 min 

(4) Distilled water - several dips - (approx. time) 1 min 

(5) Destain in 0.1 N sulfuric acid2 - 10-15 min (time 

varies with specific tissue) 

(6) Running tap water - 10 min 

(7) Distilled water - 3 dips 

(8) Methylene blue3 - dip quickly (stains quickly, depending on tissue) 

(9) Running tap water - 5 min 

(10) Dehydrate 

(11) Clear 

(12) Mount 

Results: 

Background - pale blue 

AFB - brilliant red 

Reagents: 

!Ziehl's fuchsin (Gray, 1954) 

Basic fuchsin (magenta) (Fisher Scientific Cat. 

No. P-389d) 

Phenol (liquefied) , , , , , , , , . , 

90% ETOH, 

Distilled water 

lQ.l N sulfuric acid (H2S04) 

Distilled water 

H2S04 sp. gr. 1.84 ' 

lMethylene blue stock solution (Luna, 1968) 

Methylene blue , 

95% ETOH , 

Methylene blue working solution (Luna, 1968) 

Methylene blue stock 

Tap water ' , , , , , , . , 

1.0gm 

5.0 ml 

10.0 ml 

100.0 ml 

, 997.2 ml 

2.8 ml 

1.4 gm 

100.0 ml 

10.0 ml 

90.0ml 

References: Gray P. The Microtomist's Formulary and Guide. 

The Blakiston Co., New York; 1954. 

Luna, L.G., ed. Manual of Histologic Staining Methods of 

the Armed Forces Institute of Pathology. 3rd ed. McGrawHill, New York; 1968. 

80 
B. PROBLEMS AND RESOLUTIONS 

Table 8 includes some problems or artifacts resulting from 

staining, as well as some means of resolving them. 

C. NOTES 

aRoboz Surgical Instrument Co., Inc., 810 18th Street, N.W., 

Washington , D.C. 

b American Scientific Products, 8855 McGaw Road , Columbia, 

Md. 

cRoboz Surgical Instrument, Co., Inc. , 810 18th Street, N.W. , 

Washington,D.C. 

dFisher Scientific Company, 711 Forbes Avenue, Pittsburgh, 

Pa. 

eVWR Scientific, Inc., 6601 Amberton Drive, Baltimore, Md. 

D. REFERENCES 

Brown , J.H. and L. Brenn. 1931. A method for the differential 

staining of gram-positive and gram-negative bacteria in 

tissue sections. BuH. Johns Hopkins Hosp. 48: 69-73. 

Conn, H.J., M.A. Darrow, and V.M. Emme!. 1962. Staining 

Procedures Used by the Biological Stain Commission. 2nd 

ed. The Williams & Wilkins Co., Baltimore, Md. 

Coolidge, BJ. and R.M. Howard . 1979. Animal Histology 

Procedures of the Pathological Technology Section of the 

National Cancer Institute. 2nd ed. Nat!. Inst. Health Pub!. 

No. 80-275, Bethesda Md. 

Dahl, L.K. and V.P. Dole. 1952. The biphasic nature of renal 

calcification. 1. Exp. Med. 95: 341-346. 

Drury, R.A.B. and S.A. Wallington. 1967. Carleton's Histological 

Technique. 4th ed. Oxford Univ. Press, New York. 

Farley, C.A. 1965. A modification of Noland's stain for 

permanent smears of protozoan flagella, cilia, and spore 

filaments. 1. Parasitol. 51 : 834. 

81 

Farley , C.A. 1969. Probable neoplastic disease of the hematopoietic system in oysters, Crassostrea virginica and 

Crassostrea gigas, pp. 541-555. In Neoplasms and Related 

Disorders of Invertebrate and Lower Vertebrate Animals 

(C.J. Dawe and J.c. Harshbarger, eds.), Natl. Cancer Inst. 

Monogr. 31. 

Feder, N. and T.P. O'Brien. 1968. Plant microtechnique: some 

principles and new methods. Am. 1. Bot. 55: 123-142. 

Gray, P. 1954. The Microtomist's Formulary and Guide. The 

, Blakiston Co., New York. 

Johnson, P.T. 1980. Histology of the Blue Crab, Callinectes 

sapidus. A Model for the Decapoda. Praeger, New York. 

Lillie, R.D. 1954. Histopathologic Technic and Practical 

Histochemistry. McGraw-Hili, New York. 

Lillie, R.D. 1965. Histopathologic Technic and Practical 

Histochemistry. 3rd ed . McGraw-Hill , New York. 

Luna, L.G., ed. 1968. Manual of Histological Staining Methods 

of the Armed Forces Institute of Pathology. 3rd ed. 

McGraw-Hili, New York. 

Mallory, F.B. 1968. Pathological Technique. Hafner Pub!. Co., 

New York. 

Mitchell, H.L. 1966. A simple method of permanent staining of 

intestinal parasites, using dimethyl sulfoxide. Tech. Bull. 

Regist. Med . Techno!. 36: 45-46. 

Pauley, G.B. 1967. A modification of Mallory'S aniline blue 

collagen stain for oyster tissue. 1. Invertebr. Patho!. 9: 

268-269. 

Pearse, A.G.E. 1960. Histochemistry, Theoretical and Applied. 

2nd ed. Little, Brown and Co., Boston. 

Sanders, BJ. 1972. Animal Histology Procedures of the 

Pathological Technology Section of the National Cancer 

Institute. Government Printing Office, Washington, D.C. 

Thompson, S.W. 1966. Selected Histochemical and Histopathological Methods. Charles C. Thomas Pub!., Springfield , 

III. 

Trump, B.F. , E.A. Smucker, and E.P. Benditt. 1961. A method 

for staining epoxy sections for light microscopy. 1. 

UItrastruct. Res. 5: 343. 
TABLE 8. STAINING PROBLEMS AND RESOLUTIONS. 

Problem or artifact Possible causes 

Tissues have greasy, oily appearance. Tissues are not completely dehydrated; water is 

not miscible with xylene ; therefore , the tissues 

will not accept the xylene. Large amounts of water 

cause whitish scum on tissues. 

Tissues appear dull and whitish after using Excess nuclear fast red remains in tissue. 

nuclear fast red. 

Iron reaction does not work on control. Solutions contaminated. 

Silver precipitate all over slides. Dirty slides or contaminated solutions. 

Reactions faded on selected special stains. 

Sections will not take stain, or stain irregularly. 

Alcian blue not distinct. 

Schiffs (Feulgen) reagent nonspecific on 

tissues fixed in 1 % glutaraldehyde/4% 

formaldehyde (I G4F). 

Schiffs (Feulgen) reagent faded or weak 

reaction. 

Hematein in Heidenhain's iron hematoxylin faded, not clear. 

Slides left in water for extended times. 

Paraffin not completely removed before staining. 

Section not uniform in thickness. Tissue "old" 

(long storage in alcohol or, worse, in fixative). 

Fixative not suitable for staining technique. Fixative not fully removed. 

Check expiration date of stain. 

If fixed with IG4F, fixative was inadequately 

washed out before processing tissue. 

Schiffs has become inactive or taken on too much 

moisture. 

Contamination with the mordant. 

Yellow, water-spotted appearance of stain Too much butanol carry-over in xylene. 

on slide after using picromethyl blue. 

Loss of picric acid in picromethyl blue. Too much ethyl alcohol used in dehydration. 

Tissue falls off slide. (I) Moisture left under tissue ; (2) tissue did not dry 

flat; (3) swollen tissue , does not hold together 

well ; (4) tissues difficult to cut (hard, brittle), do 

not hold together well ; (S) effect of stain too 

harsh on tissue , use alternative; (6) alkaline reagents dissolve albumin adhesive, sections start 

to work loose in course of staining or dehydrating; (7) using heat to speed reaction time is usually 

hard on tissue ; or (8) slide dirty or greasy. 

82 

Solution, if any . 

Take tissues back to 100% ETOH (or dehydrant 

used in staining protocol) and dehydrate; use new 

xylenes to clear before trying to coverslip. 

Rinse slides well in distilled water before 

dehydration. 

A void metal racks or forceps. Use acid-cleaned 

glassware. 

Use acid-cleaned slides and glassware (critical) 

and change solutions frequently; avoid metal 

contamination. 

Occasionally check pH of distilled and tap water, 

particularly before special stains. 

Return sections through proper sequence of 

reagents to xylene. Leave until paraffin removed. 

Restain. Change xylene frequently. Return sections through proper sequence of reagents to 

water. Wash overnight and restain. Store tissues 

as embedded paraffin blocks- never store tissues 

in liquids for extended time. Mordant sections in 

recommended fixative . 

Check pH of stain and use test slide before main 

staining run . 

Wash tissues well to remove excess glutaraldehyde 

before processing. May try aldehyde blocking 

agent. 

Make new batch ; color is of teo indicative of the 

stain's potency. Best results are with a peach-straw 

color. Note: After reagent becomes clear, a 

slight purple tint will show stain to be ineffective. 

Always use test slide. 

Take care not to expose hematein directly to mordant. If using solution more than once, color 

change from brown to black-brown will indicate 

contamination. 

Change xylenes frequently if staining a large number of slides. 

Use 1:9 ratio of 100% ETOH to butanol; tissues 

can be destained with acid alcohol and restained 

with picromethyl blue ifcolor not green enough. 

Dry slides adequately before staining. Spread tissue folds if they cannot be avoided, before drying 

tissue. Coat suspected loose tissue slides with celloidin after second 100% ETOH (except with 

alcian blue). For hard or brittle tissues, use extended staining time, e.g. overnight in solution 

instead of 1 hr at 60°C. Be sure slides are clean 

before albuminizing. 
CHAPTER VII. COVERSLIPPING 

A. TOOLS AND TECHNIQUES 

Individual techniques of coverslipping tissue may vary from 

technician to technician. Exactly how to hold the slide or 

coverslip will vary with one's dexterity and comfort. However, 

the basic procedure is simple. 

Materials needed are: (1) fume hood , (2) good light, (3) 

coverslip, (4) probe (straight), (5) paper towels, (6) Kimwipes', 

and (7) mounting medium. 

Once the slides have been stained , dehydrated, and placed 

in xylene, they are ready to be coverslipped for permanence. 

This work should be done under a fume hood. 

Remove slides singly in numerical order. Be sure the etched, 

coded side of the slide is up. With a lint-free tissue (Kimwipes), 

remove excess xylene from the front and back of the slide (Fig. 

72), and place one drop of mounting fluid on the tissue on the 

slide (Fig. 73). Choose the proper size coverslip and gently 

lower the coverslip across the slide gradually (Fig. 74) so that 

the mounting fluid flows evenly from bottom to top. (Do not 

drop coverslip onto slide because this will usually trap air 

bubbles under the coverslip.) 

Once the coverslip is attached, blot the edges of the slide on 

a paper towel (Fig. 75) and examine for occasional air bubbles. 

If present, gently lead the bubble to the edge, out from under 

the coverslip, using a probe. This procedure should be done 

quickly, precisely, and with care to prevent damage to the 

tissue. Place the slide on a flat tray (Fig. 76) along with an 

identifying label with the code, stain, and date of coverslipping. 

The slides are then placed in a drying oven set at approximately 

42°C for at least 2 weeks to allow the mounting medium to 

harden. 

83 

After the mounting medium hardens, the slide can be cleaned, 

labeled, and filed for examination (Fig. 77). Using a singleedged razor blade, gently but firmly remove all excess mounting 

medium from around the coverslip. Be careful not to nick the 

coverslip with the razor blade. Brush off the loosened mounting 

medium, dip the slide in xylene, and gently wipe clean. When 

all the slides have been cleaned, make sure they are in the 

correct series and numerical order. Using a Koh-I-Noor 

rapidograph pen2 containing India ink, write all pertinent codes 

on the etched end of the slide, including staining procedure 

used and date fixed. 

Warning: Coverslip and clean all slides under a fume hood. 

B. PROBLEMS AND RESOLUTIONS 

Table 9 includes some problems or artifacts resulting from 

coverslipping, as wel.l as some means of resolving them. 

C. NOTES 

'Kimberly-Clark Corp., Neenah, Wisc. 

2Koh-I-Noor Rapidograph, Inc., Bloomsbury, N.J. 

D. REFERENCE 

Luna, L.G., ed. 1968. Manual of Histologic Staining Methods 

of the Armed Forces Institute of Pathology. 3rd ed. 

McGraw-Hili, New York. 
TABLE 9. COVERSLIPPING 

Problem or artifact 

Mounting medium will not flow across 

tissue. 

Coverslip tends to slip out of place. 

Air bubbles will not release from under 

slide. 

Black, singed-looking, or parched tissue. 

Sections appear opaque or have highly refractive lines outlining cells and tissues. 

Cloudy slide. 

Black air rings in tissue. 

Tissues dislodged out of place, layered, or 

unrecognizable. 

Possible causes 

Slide has dried out. Mounting medium too thick ; 

part of the solvent may have evaporated. 

Slide too wet with xylene. Mounting medium too 

thin . Too much mounting medium. 

Grit or loose debris under slide. Old coverglasses 

may be warped or dirty. 

Tissue dried out before or during coverslipping or 

large air pockets formed from warped coverslips. 

Clearing agent evaporated before mounting 

medium added. Sections insufficiently cleared or 

clearing agent not miscible with mounting 

medium. 

Too much mounting medium or too thick. 

Air trapped during coverslipping. 

Too much pressure on probe to remove air bubbles 

84 

Solution, if any 

Redip slide into xylene and start process again. Use 

new mounting medium or carefully thin down 

medium with solvent. 

Wipe excess xylene from around tissue and from 

back of slide before adding mounting medium. 

Blot edge of slide once coverslip is in place. 

Mounting medium should have the right consistency. Use fresh mounting medium. 

This often occurs with very sandy tissues. Remove 

coverslip by soaking in xylene and start process 

again. If trouble spot can be identified , remove it. 

However, if it lies within the tissue , try mounting 

with thicker mounting medium. Re-coverslip with 

a new coverglass. 

Make sure tissue is covered with xylene before 

coverslipping; tissue should never be allowed to 

dry out after deparaffinizing (Luna , 1968). 

None. Soak off cover. Clear properly. Check quality and nature of clearing agents and mounting 

medium (Luna, 1968). 

Mounting medium should be free-flowing; only 

use amount necessary and blot edge of slide after 

coverslipping to remove excess mounting 

medium. 

Press coverslip down gently and use probe to guide 

bubbles to outer edges of coverslip (Luna , 1968). 

If there is a large air bubble or many air bubbles, 

do not force out. Re-coverslip instead to avoid 

damaging tissue. 
Coverslipping. 

Fig. 72. Wiping excess xylene from back of slide. 

Fig. 73. Dropping mounting medium Over tissue section. 

Fig. 74. Lowering coverglass over tissue. 

Fig. 75. Draining excess mounting medium from slide. 

Fill. 76. Placing a stained slide on a flat tray to dry. 

Fill. 77. Slides cleaned and labeled, ready to file. 

85 

77 
APPENDIX 

A. SPECIAL TECHN.aUES 

1. Culture of Perlcinsus marinus 

Thioglycolate cultures are routinely used on many fresh 

shellfish specimens. The thioglycolate culture technique described below was devised by Mackin and Ray in 1966. It is 

routinely used for detection of the marine fungus, Perkinsus 

marinus (Levine, 1978). This method is apparently specific for 

P. marinus, is effective in discerning both light and heavy 

infections, and does not differ from species to species. 

Procedure for preparing fluid thioglycolate culture mediuml

: 

(I) Heat distilled water to a boil 

(2) Stir in thioglycolate medium (containing dextrose) 

and NaCI (salt) till both are dissolved 

(3) Remove from heat and add chloromycetin2 

(4) Dispense 9.5 ml of the medium into each tube (should 

fill 50 tubes) 

(5) Autoclave 15 min at 15-17 lb pressure 

(6) Store in the dark at room temperature until needed 

(7) Add 0.5 ml of mycostatin3 suspension into each 9.5-ml 

tube immediately before using. Do not shake tubes. 

(8) After placing tissues in culture tube, return to the dark 

for at least 72 hr to culture before examination 

Reagents: 

lFluid thioglycolate culture medium 

Thioglycolate medium ..... ..... . . . 

NaCI 

Distilled water 

2Chloromycetin (chloramphenicol, 

Sigma Cat. No. C'{)376a) 

Chloromycetin ........ . 

Distilled water 

3Mycostatin suspension (nystatin, 

14.6 gm 

10.0 gm 

485.0 ml 

0.25 gm 

1O.0ml 

Sigma Cat. No. N 3503b) 

Mycostatin 

Sterile distilled water . 

500,000 usp units 

125.0 ml 

Shake well before using. Refrigerate unused 

portion for further use. 

Procedure for preparing slides for examination: 

(1) Excise rectal tissue from mollusk during initial necropsy 

(Fig. 78) 

(2) Remove tissue from tube with a thin glass rod with 

hooked end (Fig. 79) and place on the correspondingly 

coded slide with a drop of Lugol's iodine working 

solution4 (Fig. 80) 

(3) Tease tissue apart with probes (Fig. 81) and add another 

drop of Lugol's (Fig. 82 .iodine working solution 

(4) Place a coverslip on the slide (Fig. 83) 

(5) Place finished slide on a tray 

(6) Wait 15 min, then examine the slide 

Note: Clean instruments after handling of each specimen to 

avoid contamination 

86 

Results: 

Examine with the use of a dissecting microscope. A positive infection of "dermo" will show symmetrically round 

blue-black to solid black spheres. See Ray, 1966 and Kern 

et aI., 1973 for additional information. Figure 84 illustrates 

a positive fungus infection of the duck clam, Macoma 

balthica. 

Reagents: 

4Lugol's iodine stock solution 

Potassium iodide 

Iodine 

Distilled water 

Lugol's iodine working solution for thioglycolate 

Distilled water 

LugoJ's stock 

Dispense from 5().ml dropper bottle 

2. Decalcification 

6.0gm 

4.0gm 

100.0 ml 

3D.Oml 

15.0 ml 

Decalcification is not often utilized with molluscan tissues. 

However, in special circumstances, for example when processing unshucked larvae of bivalves, we have found the versenate 

method of decalcification to give excellent results. (Versenate 

is a proprietary term for disodium ethylenediaminetetraacetate 

(EDTA». This chelating method of decalcification works gradually and does not damage the internal animal tissues. 

It will take different time periods for decalcifying larvae, 

depending on species and size. The best results are obtained by 

frequently changing the versenate solution over a period of 

days if necessary. Pearls found in oysters, mussels, and clams 

should be removed prior to embedment unless the investigator 

specifically requires their presence. Also, the sensory eyes of 

scallops, located along the mantle, should be avoided in tissue 

sections unless necessary for specific diagnosis. If either pearls 

or scallop eyes are required by the investigator, they can be 

decalcified by the versenate method. 

Procedure for versenate method of decalCification (Birge and 

Imhoff, 1952): 

(1) Fix tissue in fixative of choice 

(2) Wash well in running water (the time will depend on 

fixative used and size of tissue) 

(3) Place tissue in a saturated solution of versenatel until 

decalcification is complete; check daily 

(4) Wash well in running water - Y2-1 hr (depending on 

size of tissue) 

(5) Place in 70% ETOH until ready to process 

Reagents: 

lVersenate solution 

Versenate . . .. ... . . 

Distilled water (pH 5.5-6.5) ... . ... .. . ... . 

10.0 gm 

100.0 ml 
Note: For the most efficient use of verse nate as a decalcifier, change tissues to a fresh solution every 2-3 days. We 

have had great success using versenate on bones found in 

fish tissue. 

specimens may be left in the versenate solution for 

as long as 14 days without noticeable effect on the staining 

qualities of the cells" (Birge and Imhoff, 1952). 

Warning: Versenate may remove iron and other metals in the 

process of decalcification , according to Carleton:S Histological Technique (Drury and Wallington, 1967). 

3. Transfer of Tissue from Broken Slide 

(Coolidge and Howard, 1979) 

When it is not desirable to mount a broken slide on another 

slide because of excessive thickness (when high resolution is 

needed on the microscope) or if a slide has been broken in too 

many pieces, use the following method : 

(1) In a shallow pan, assemble all pil'!ces of broken slide 

on another slide. 

(2) Slowly pour xylene in pan until slide is covered. 

(3) Soak until all pieces of coverslip can be easily removed. 

Lift slide from container and place on paper towel. 

While broken pieces are still wet with xylene, pour 

DiatexC (plastic) on entire slide. 

(4) Dry overnight at room temperature or in a 60°C oven 

for 2 hr (plastic will be hard). 

(5) Cut around edge of slide with a razor blade. 

(6) Soak in tap water (about 2 hr) until plastic can be easily 

peeled from slide. 

(7) Trim around tissue with scissors. 

(8) Place plastic-coated section on clean albuminized 

slide. Cut a paper towel the size of the slide, dampen 

it, and place it on top of the plastic film. Place a clean 

slide on top. An object should be placed on top of slide 

to hold it in place. Leave at room temperature or place 

in 37°C oven overnight (towel will be dry). 

(9) Remove top slide. Put slide with film and paper towel 

into xylene. The towel will soak loose, and the slide 

should remain in xylene until the plastic is completely 

dissolved. 

(10) Coverslip. 

4. Transfer of Tissue from Paraffin to Plastic 

(Howard and Wade, pers. commun. d) 

The following procedure was devised to use when it is desirable to examine the ultrastructure of a specific area of interest 

on a histological slide. 

A modification of the tissue transfer process from Animal Histology Procedures of the Pathological Technology Section of 

the National Cancer Institute (Coolidge and Howard, 1979) is 

used to re-embed a cut and stained section of tissue from paraffin to resin without loss of orientation: 

87 

(I) Begin by cutting a practice block at 30-50 J-lm of the 

same type of tissue as your specimen at room temperature. The goal here is to cut evenly through the 

tissue without breaking apart the section. 

(2) Align the block of interest in the chuck of the microtome. Cut two 6-J-lm sections before changing the 

cutting gauge to desired thickness (30-50 J-lm). The 

thicker the tissue. can be cut without damage , the 

greater the prospect of finding the desired areas. 

(3) Moisten the block and cut through the whole face of 

the block with an even motion. A thick section will 

generally come off the microtome knife in a thick 

roll. 

(4) Gently and carefully raise the tissue roll with the aid 

of brushes to an albuminized slide moistened with a 

small pool of distilled water. 

(5) Drain excess water and place slide on warming tray 

at 42°C. 

(6) Unroll tissue slowly, carefully manipulating brushes 

(Fig. 85). 

(7) Continue to unroll section (Fig. 86); gently flatten 

section and place slide in oven at 42°C. 

(8) Rechill block which is still in the chuck with a piece 

of ice and cut another 6-J-lm slide section. 

(9) Once the slides are thoroughly dry, deparaffinize and 

stain the 6-J-lm sections in Harris' hematoxylin and 

eosin. 

(10) Check to see if desired area is present. 

(11) Proceed with thick section. Deparaffinize and stain 

with hematoxylin (I min), eosin (30 sec), and mount 

with Diatex synthetic liquid coverglass (Fig. 87). 

(12) Dry overnight at room temperature or in slide oven 

at 42°C until plastic is hard; check under microscope. 

(13) Cover slide with paper towel and soak with tap water 

(about 2 hr) until plastic is pliable and can be gently 

peeled away from slide (spatula may be used for 

thicker sections to keep from damaging tissue section). 

(14) Compare plastic section with 6-pm slides where area 

of interest has been located and marked. 

(15) Cut plastic section as indicated in the marked 6-pm 

sections; use a specific shape (e.g. triangle or truncate 

pyramid) (Fig. 88) to ensure proper orientation of tissue later. 

(16) Place tissue in a porcelain spot plate or small shallow glass dish; add several changes of xylene until 

tissue is free from all Diatex; pipette new xylene on 

and off, trying not to disorient tissue. 

(17) Remove xylene with several changes of 100% ETOH. 

The ETOH mayor may not be replaced with several 

changes of propylene oxide, depending upon the 

choice of embedding medium to be used . 

(18) Leave section in a 1: 1 mix of 100% ETOH and embedding medium for an hour, followed by 1-2 hr in the 

complete embedding medium. 

(19) Embed section of tissue in fresh embedding medium; 

check again for orientation and place mold carefully 

in 70°C oven for 48 hr. 

(20) Before cutting thin sections from the cured blocks, 

cut and stain a I-pm section for positive identification of the tissue desired. 
Procedure for delermining Perkinsus marinus in Ihioglycolale cullured reclallissue of Maeoma balthiea. 

Fig. 78. Pulling fresbly cui reclallissue in Ihioglycolale. Fig.81. Teasing lissue aparl for beller penelralion of iodine. 

Fig. 79. Removing cultured lissue wilh glass hook. Fig. 82. Placing anolber drop of iodine on lissue. 

Fig. SO. Placing lissue inlo drop of Lugol's iodine. Fig. 83. Tissues covered and ready for inspeclion. 

88 
Fig. 84. Positive funj/us infection of duck clam, Macoma 

balthica. 

89 
Tissue transfer from paraffin to plastic. 

Fig. 85. Placing tbick section (appro>:. SO I'mt on slide and rolling open 

Fig. 86. 

Fig. 87. 

Fig. 88. 

by careful manipulation of brushes. 

Section rolling open. 

Covering stained section with Diatex liquid coverglass. 

Cutting plastic section in a specific shape (e.g. truncate pyramid t 

to ensure proper orientation of tissue during embedment. 

90 
5. Record Keeping 

The two data fonns shown on the next two pages cover a wide 

range of infonnation. Each fonn follows the sample through a 

specific step from arrival and code designation through the 

final histological examination. A summary of all information is 

statistically presented on the Molluscan Pathology Report. 

All of the compiled data are filed in a pennanent information 

bank (data fonns courtesy of C.A. Farley). 

6. Filing and Slide Storage 

All samples are kept in permanent files. Once the sample is 

coded, either in the field or after arrival, it is recorded with 

its historical data. Each investigator is responsible for keeping 

his/her own records, but we compile all the infonnation given 

us. It is recorded in our master files and entered into a current 

sample index. In the index, the code number, specimen or identification type, date of arrival, fixative, and embedment, date 

of sectioning and staining, identity of requestor, and source are 

recorded. A work ledger is also kept in which we record daily 

work activities perfonned by the laboratory. Maintaining upto-date records is imperative. 

Once the pennanent slides have been completed and issued 

to the requestor, the tissue blocks are filed consecutively in 

trays and stored. A record book identifies the location of all 

blocks in storage. They can be readily retrieved by code numbers. The block storage area is analogous to a library. Study 

projects with consecutive code numbers are filed together. 

Samples from similar geographic areas are also consolidated 

when possible. A record guide specifies block location by aisle, 

row, and cabinet. If study blocks have been removed, this is 

so indicated, and their current location is recorded. This system 

enables us to keep an orderly collection of blocks which is of 

value for retrieval and future retrospective study. 

91 

B. NOTES 

a,bSigma Chemical Company, P.O. Box 14508, St. Louis, Mo. 

cAmerican Scientific Products, 8855 !y1cGaw Road, 

Columbia, Md. 

dD.H. Howard and IT. Wade, National Marine Fisheries 

Service, Oxford, Md.; 1976. 

C. REFERENCES 

Birge , E.A. and C.E. Imhoff. 1952. Versenate as a decalcifying 

agent for bone. Am. 1. Clin. Pathol. 22: 192-193. 

Coolidge, B.1. and R.M. Howard. 1979. Animal Histology Procedures of the Pathological Technology Section of the 

National Cancer Institute. 2nd ed. Natl. lnst. Health Publ. 

No. 80-275, Bethesda, Md. 

Drury, R.A.B. and E.A. Wallington. 1967. Carleton's Histological Technique. 4th ed. Oxford Univ. Press, New York. 

Kern, F.G., L.c. Sullivan, and M. Takata. 1973. Labyrinthomyxa-Iike organisms associated with mass mortalities of 

oysters. Proc. Natl. Shellfish. Assoc. 63: 43-46. 

Levine, N.D. 1978. Perkinsus gen. n. and other new taxa in the 

protozoan phylum Apicomplexa. 1 Parasitol. 64: 549. 

Mackin,J.G. and S.M. Ray. 1966. The taxonomic relationships 

of Dermocystidium marinum Mackin, Owen , and Collier. 

1 Invertebr. Pathol. 8: 544-545. 

Mackin , lG., H.M. Owen , and A. Collier. 1950. Preliminary 

note on the occurrence of a new protistan parasite , Dermocystidium marinum n. sp. in Crassostrea virginica (Gmelin). 

Science 111 : 328-329. 

Ray, S.M. 1966. A review of the culture method for detecting 

Dermocystidium marinum, with suggested modifications 

and precautions. Proc. Natl. Shellfish. Assoc. 54: 55-69. 
FIELD DATA ON OYSTER SAMPLES 

USNMFS, Oxford, Maryland 

Comparative Invertebrate Pathology 

Date sample collected 

Date sample received 

------------------

--------------------- Source: State No. oysters Code designation --------------- ---------- --------------------

Estuary Bar Depth water ft. ----------------------- ------------------- ----------------------

Bottom water: temp. °C ----------- salinity 0/00 ------------

Nature of bottom: hard, medium, soft, sandy, mud, shell, 

other ------------------------------

Additional comments on field collection and condition of oysters: 

GROSS EXAMINATION OF INDIVIDUAL OYSTERS in Laboratory (over), by --------------------

Date Time oysters examined and fixed; --------------- ---------------

Fixative ---------------------------------

92 
MOLLUSCAN PATHOLOGY REPORT 

Date sample received Purpose Date sent - ---- -- ----------------------- --------------- Requested by Phone 

Send report to _____________________________________________ _ 

Environmental data: 

Temperature °C Salinity 0/00 pH ----- - ---- ---- Oxygen ppm Other ----~ ---------

Species Stock ------------------------------------------ Origin ____________________________________________ _ 

History --------------------------------------------

Location sample came from ------------------------------------ Number in sample Date collected Date fixed Code ----- ------ ----- --------

Fixative Other histologic data - --- ---- --------------------------------------------

GROSS PATHOLOGY 

Condition Mantl e recession Pale digestive gland Cliona Polydora Other 

Fat Medium Watery 

% % % % % % % % - ----------- ------------------ ---

MICROSCOPIC PATHOLOGY 

Pathologic lesions Parasites 

Category Type of lesion Prevalence Degree Group Species Prevalence Degree 

(0-9) (0-9) 

Inflammation % Vi ruses % ------- ------------- - ---- ------ % % ------------------------------------ ------

% Chlamydia % ---------------------------------

% % --------------------------- --------- ------

% Bacteria % ----------------------------------

% % --------- -------

% Fungi % ---------------------- ------- % % ----~---------- ---------- -------

% Protozoa % ------------------- ------- ------

% % -------------------- - - ---- ------

% % --------------------------- --------- ------

% Metazoa % ------------------ ------

% % --------------

SUMMARY --------------------------------------------------------------------------

93 
GENERAL REFERENCES 

Bloom, W. and D. Fawcett. 1968. A Textbook of Histology. 

9th ed. W.B. Saunders Co., Philadelphia, Pa. 

Coolidge, B.I. and R.M. Howard. 1979. Animal Histology Procedures of the Pathological Technology Section of the 

National Cancer Institute. 2nd ed. Natl. Inst. Health Publ. 

No. 80-275, Bethesda, Md. 

Drury, R.A.B. and E.A. Wallington. 1967. Carleton's Histological Technique. 4th ed. Oxford Univ. Press, New York. 

Galtsoff, P.S. 1964. The American Oyster, Crassostrea virginica 

Gmelin. U.S. Bur. Com mer. Fish., Fish. Bull. 64,480 pp. 

Gray, P. 1954. The Microtomist's Formulary and Guide. The 

Blakiston Co., New York. 

Grizzle, 1M. and W.A. Robers. 1976. Anatomy and Histology 

of the Channel Catfish. Auburn Univ. Agricultural Experiment Station, Auburn, AJa. Auburn Printing, Inc., Aubuni, 

Ala. 

Haas, E. 1980. SO Diagnostic Special Stains for Surgical Pathology. All-Type Editorial, Los Angeles, Calif. 

Ham, A.W. 1969. Histology. 6th ed. lB. Lippincott Co., Philadelphia, Pa. 

Humason, G.L. 1967. Animal Tissue Techniques. 2nd ed. W.H. 

Freeman and Co., San Francisco, Calif. 

Hyman, L.H.1940. The Invertebrates: Protozoa Through Ctenophora. Vol. 1. McGraw-Hill, New York. 

Hyman, L.H. 1951. The Invertebrates: Platyhelminthes and 

Rhynchocoela. Vol. II. McGraw-Hill, New York. 

Hyman, L.H. 1951. The Invertebrates: Acanthocephala, Aschelminthes, and Entoprocta. Vol. III. McGraw-Hill, New 

York. 

Hyman, L.H. 1955. The Invertebrates: Echinodermata. Vol. IV. 

McGraw-Hill, New York. 

Hyman, L.H. 1959. The Invertebrates: Smaller Coelom ate 

Groups. Vol. V. McGraw-Hill, New York. 

94 

Hyman, L.H. 1967. The Invertebrates. Vol. VI. Mollusca I. 

McGraw-Hill, New York. 

Johnson, P.T. 1980. Histology of the Blue Crab, Ca/linecles 

sapidus. A Model for the Decapoda. Praeger, New York. 

Klontz, G.W. 1973. Syllabus of Fish Health Management. Texas 

A&M Univ., College Station, Tex. 

Lillie, R.D. 1965. Histopathologic Technic and Practical Histochemistry. 3rd ed. McGraw-Hill New York. 

Lucky, Z. 1977. Methods for the Diagnosis of Fish Diseases. 

Amerind Publ. Co., New Delhi. 

Luna, L.G., ed. 1968. Manual of Histologic Staining Methods of 

the Armed Forces Institute of Pathology. 3rd ed. McGrawHill, New York. 

Pearse, A.G.E. 1960. Histochemistry, Theoretical and Applied. 

2nd ed. Little, Brown and Co., Boston. 

Roberts, R.I., ed. 1978. Fish Pathology. Bailliere Tindall, 

London. 

Sheehan, D.C. and B.B. Hrapchak. 1973. Theory and Practice 

of Histotechnology. C.V. Mosby Co., St. Louis. 

Sheehan, D.C. and B.B. Hrapchak. 1980. Theory and Practice 

of Histotechnology. 2nd ed. C.V. Mosby Co., St. Louis. 

Sindermann, C.I. 1970. Principal Diseases of Marine Fish and 

Shellfish. Academic Press, New York . 

Sparks, A.K. 1972. Invertebrate Pathology. Noncommunicable 

Diseases. Academic Press, New York. 

Thompson, S. W. 1966. Selected Histochemical and Histopathological Methods. Charles C. Thomas Pub!., Springfield, 

Ill. 

Yevich, P. and C.A. Barszcz. 1979. Preparation of Aquatic 

Animals for Histopathological Examination. Mimeograph, 

U.S. Environmental Protection Agency, Narragansett, R.1. 
INDEX 

Acid clean glassware, . . . . . . . .. . ....... 54 Fats, . . . . . . . . . . . . . . . . . . . . . . . . . .. .. 50,51 ,71 ,75 

Acid-fast, . . . . 51 ,79 ,80 stains for, oil red 0, .. • 71 

stains for, Ziehl and Harris' hematoxylin, .. . 51 ,79 Sudan black B, . . . . . . . . . . . . . . . . . . .. . .15 

Ziehl and methylene blue, . .. . . . . . . . . . . . 51 ,80 Feulgen gram stain, . . . . . . . . . . . . . . . 50,51,60 

Acid fuchsin, ...... 68 Feulgen picromethyl blue stain , ... ... 50,51 ,61 

Aldehyde blocking technique, . . . .. . 54,82 Filing and storage , . . . . . . . . . . . . . . . .... . . . .. 91 

Alcian blue stain , . . .. 54,55,70,82 Fixation, . . . . . . . . . . . .. 5,35 

Alizarin red S stain, . . ... 50,51,56 furmu~ ,.. .... ~ 

Aniline blue, . . . . . . . . . . . . . ...... 68 sea water, special note on, .... . .. . . .. .. .. 37 

Arctica sp., processing of, . ... .. ... . 20 problems and resolutions, . . . . . .. . . .. 37,38 

Azure-eosin/Giemsa stain, . . . . . . . . . . ... 50,51,57,58 fu~~, .. . . ..... ~ 

Bacteria, . . . . . . . . . . . . . . . .. .. 50 Bouin's, . . . .. . 35,37,59,65 

stains for, azure-eosin, ..... 57,58 Davidson's, . . . . . . . . . . . . . . . 35,37,59,65 

Feulgen gram, .... 50,51,60 formalin , . . . . . . . . . . ..... 35,59,65 

gram, . . . . . . . . . . . . . . 62 glutaraldehyde, . . . . . . . . . . . . . . . . . .... .. . . . . 35 

Ziehl and methylene blue , . . . .. 51 ,80 1% glutaraldehyde/ 4% formalin ,. . 36,37 

Bivalve larvae, processing of, . . . . . . . . . . . . 21 ,86 Helly's, . . .. 36,37,53,59 ,64 

Blood, . . . . . . . . . . . . . . . . 50,51 ,57,58,76 Zenker's, ... 36,37,53,59;64 

stains for, azure eosin/Giemsa, ..... . 57,58 Formalin fixative, . . . . . . . . . 35,37,59,65 

toluidine blue 0, ... 51 ,76 Frozen tissue techniques, ..... 8,33,34 

Bouin's fixative, . . . . . . . . . . . ... .. 35,59 description of, . . . 33 

Calcium, . . . . . . . . . . . .. 50,51,56,77 Fungi, stains for, . . . . . . . . .50,51 ,63,72,73 

stains for, alizarin red S, . . 56 culture techniques, . . . . . . . . . . . . . . . . .. 86 

Von K6ssa, . . . . . . . . . . . . ...... . . . .77 Grocott's methenamine silver, ..... 63 

Celloidin, coating for slides, . . ... 54 periodic acid Schiff, . . . .. 72,73 

Clams, ....... 20 General tissue, stains for, . ..... . .. ... . 50,52 

cross-section, . . . . . . . . .... ... . . . .... . 20,26 Harris' hematoxylin, . . . .. 64,65 

gross anatomy, . . .. ... . .. .. ... .. . .. .... .. 25 Glutaraldehyde fixative, . .35 

processing of, duck clams, . . . 20,23,88 1 % glutaraldehyde/ 4% formaldehyde fixative , ... .. 36,37,82 

hard clams, . . . . . . . . . . ... .. 20,24 Glycogen, . . ..... .... 50,52,69,73 

soft-shell clams, . . . . . . . . . . . . 20,22 stains for, malt periodic acid Schiff, . . .. .. . .. ... 69 

surf clams, . . ...... 20 periodic acid Schiff, . . . . . . . . . . . .. 36,72,73 

Collagen, . . . . . . . . . . . .50,51,68,73 Gram stain ,. . . . . . . . . . . . . . . . . . . . . . . . . . 50,51,60,62 

Connective tissue, . . ....... . .. . 50,51,61,68,78 Grocott's methenamine silver stain , . . . . . . . . ... 50,51 ,63 

stains for, Feulgen picromethyl blue, ..... 50,51,61 Hand honing, . . . . . . . . . . . . . . . . . . . . . .. . . . .. ... .. 47 

Mallory's trichrome , . . . . . . . . . . . . 68 Handling live animal samples, . . ....... .. .. 5 

Wilder's reticulum, . . .78 Harris' hematoxylin stain, . 50,52,65,69,79 

Copper,. . . . . . . . . . . . .. . . 50,51 ,67 progressive,. . . . . . . . . . . . . . . . .. .... . 64 

stain for, Mallory's hematein, .. . 50,51,67 regressive, . . . . . . . . . . . .. ... .. 65 

Coverslipping, .. . .. 53,83,84,85 Heidenhain's iron hematoxylin stain , . . . 50,52,66,73,82 

problems and resolutions, . . .... ... 83 HeUy's fixative , . . . . . . . . . . 36,37,53,59,64 

tools and techniques, . .. ........ ... 83 Hematein, . . . . . . . . . .... 50,51,66,67,82 

Crystal violet, .. ...... . . . . . . ...... 60,62 Hematoxylin , . . . . . . . . . . .. 50,52,64,65,66,87 

Crystalline style, . . ... 8,20,22,44 Hemocytes, . . . . . . . . . . ....... 5,6,51,76 

Davidson's fixative, . . ... 35,59,64 Iron reaction , . . . . . . . . . . . . . . . . . . . . ....... 82 

Decalcification, .... 86 

technique for, . . .. 86 

Dermocystidi.um, . . . . . . . . . . . . 8 

DNA, stain for, . . . . . . . . . . . .. 50,51 ,61 

stain for, Perls' Prussian blue, ... 50,52,74 

Larvae, . .. . . . .. . . 21,86 

bivalve, processing of, .. 21,86 

Light green , . . . . . . . . . 56 see Feulgen picromethyl blue, . . . . . . . . . .. .... . 61 Macoma sp., .... 20,23,88,89,90 

Embedding, . . . . . . . . . . . . . . ... . .. 39,41 cross-section, .. .... . 20,23 

general procedure , . . . . . . . . . . . ... 39 processing of, . . . . . . . . . . . . . . . . . 20,23 

problems and resolutions, . . . . . . . . . . . . .. .. . 39,40 Mallory's hematein stain, . . . . . . . . . . . . . . . . . . . 50,51,67 

schedules, . . . ...... .. 40 Mallory's trichrome stain, . . . . . . . . . . . . 50,51,68 

~hr, . . .. 40 Malt periodic acid Schiff stain, . . . . . . . . . . ... 50,52,69 

19-hr,. . . . . . . . . ... 40 Malt periodic acid Schiff-alcian blue stain , . . ... 50,52,70 

2-hr, short,. . . . . . . . . . . . . . . . . 40 McIlvaine-Lillie buffer table, . . ... .. .... 59 

7-hr, short,. . . . . . . . . . 40 Mercenaria sp., . . 20,26 

Eosin bluish , .. 64,66 cross-section, . . . .. . .. . . . ... .. .. 20,26 

Eosiny, . .... . . . . .... . 65 processing of, . . . . . . . . . . . . . . . .. 20,24 

95 
Minchiniasis, 6 

Mucopolysaccharides, . . ... 50,52 

stains for, alcian blue,. . . . . . . . . . . . . ..... 55 

malt periodic acid Schiff, .................. 36,69,73 

malt periodic acid Schiff-alcian blue, ....... 70 

periodic acid Schiff, . . .72,73 

Mussels, .... 20,21 

cross-section, . 21,27,29 

gross anatomy, ...... 28 

processing of, .. 20,21,27 

Mya arenaria, ........... . . . .... 20,22 

cross-section, ... 20 

processing of, . . . . . . . . . ... 20,22 

Mytilus spp., ..... . ............. . ........... 20,21,27 

cross-section, . . ... 21,27,29 

processing of, 20,21,27 

Necropsy, . .5 

gross abnormalities, . . .. 5,7 

abscesses and tumors, .................... 5,7 

pearls, ..... . . . . . . . . . . . . . ..... 5,7 

watery cysts, ... 5,7,15 

parasites, 

Bucephalus sp., ....... . ...... 5,7 

Mytilicola sp.,. . ... 5,6,13 

Pinnotheres sp., .. 5,6,13 

Sphenophrya sp., . . . . . . . . . . ... 5,6,13 

physiological conditions, ..... 5,7,14,15 

color, . . . . . . . . . . . . . . . . . . . . .... 5,7 

digestive diverticulum, .5,7,15 

swollen tissue/edema, ..... 5,7,15 

processing steps, . . ........... . ............... 5 

shell abnormalities, .. 5,9,10,11,12 

calcareous malformations, . . 6,11 

Cliona,. . .. 5,6,9 

drill cases, .. 6,12 

"maladie du pied", ......... . .......... 6,11 

mantle recession,. . .... 5,6,9 

Polydora sp., . . . . . . . . . . . . . . . . . . . . . . . .5,6,10 

recovery, . . . . . . . . . . ... 5,6,9 

shell blisters, . . . . . . . . . . . . .5,6,11 

shell pustules, ... 5,6,11 

Neutral red, . . . . . . . . . . . . ... 74,75 

Nuclear detail, ................................ 52,66 

stain for, Heidenhain's iron hematoxylin 

and eosin, 

Nuclear fast red, 

Oil red 0 stain, . 

Oysters, 

cross-section, . 

gross abnormalities, . 

gross anatomy, ........ . . . 

50,52,66 

. 55,77,78 

. ... 50,51,71 

...... 7 

. . . . . . 8,19 

............. 7 

18 

parasites, ................................... 5,13 

physiological conditions, .. 14,15 

processing of, 7,16,17 

shell abnormalities, . 5 

Parasites, 6,13 

Bucephalus sp., ....... 5,7 

Mytilicola sp., .... . .............. . .......... 5,6,13 

Pinnotheres sp., ... 5,6,13 

Sphenophrya sp., ........ 5,6,13 

Periodic acid Schiff stain, . ... 50,52,72,73 

Perkinsus marinus. culture of, . . ... 86,88 

96 

Perls' Prussian blue stain, . 

Phospholipids, . 

Picromethyl blue, 

Processing, ............. .. . . 

bivalve larvae, 

clams, . 

mussels, . 

oysters, ....... . . 

scallops, . 

Record keeping, . . ..... . . 

field data on oyster samples,. 

gross examination data, 

molluscan pathology report, 

Reticulum, . 

Safety, 

Safranin 0, . 

Scallops, 

cross-section, . 

gross anatomy, . 

processing of, . 

Sensory eyes, 

Schiffs Feulgen reaction, 

Sea water, ........... . .. . 

Sectioning, . 

preparation of, . 

problems and resolutions, 

techniques, . 

serial, . 

Seed size bivalves, . 

Shipping, 

shellfish, 

Slides, filing and storage, 

Special techniques . 

culture of Perkinsus marinus .. 

decalcification, . 

filing and slide storage, . 

record keeping,. 

transfer of tissue, 

from broken slide,. 

from paraffin to plastic, . 

Spisula solidissima .. 

processing of, ........ . 

Staining, 

acid-clean glassware, 

aldehyde blocking, ..... . . 

celloidin coating, ...... . 

clearing, . 

dehydrating, . 

deparaffinizing, . 

hydrating to water, . 

mercury removal of, . 

methods and use, ...... . . 

mounting, . 

problems and resolutions,. 

stains synoptic list of, 

Sudan black B stain. 

Thioglycolate culture,. 

Transfer of tissue, 

from broken slide, 

from paraffin to plastic, . 

Toluidine blue 0 stain, 

Versenate decalcification method, 

..... 50,52,74 

. . .... .. . . ..... 51,75 

.. 60,61,82 

...... 7 

.. 21,87 

. ..... 20 

20,21,27 

7,16,17 

.21,30 

91 

.. 92 

.... 92 

..... .. ... 7,93 

... 51,78 

.. 3 

... 62 

....... 21,30 

.21,30,32 

31 

21,30 

21,44,86 

.36,60,61,69,70,72,82 

37 

... 42,46 

.42 

· .. 43,44,45 

42 

.... 43,48,49 

· .21,43 

... 5 

.. 5 

91 

....... 86 

.... 86,88 

..... 86 

· .. 91 

.. 91 

87 

· ... 87 

87,90 

. . . . . . . . .. 20 

.20 

54 

.54 

· .. 54 

· ... 53 

· ... 53 

· ... 53 

.53 

· .. 53 

50 

· ...... 53 

81 

.50,51,52 

51,75 

.8,86,88 

.87 

· ...... 87 

...... 87,90 

........ 51,76 

21,74,86,87 
Von K6ssa stain, . 

Weigert's hematoxylin ,. 

Wilder's reticulum stain, . .. . . .. . 

Zenker's fixative , .. 

Ziehl and Harris' hematoxylin stain, . 

Ziehl and methylene blue stain, ...... . . . . 

Ziehl's fuchsin, .... ... ... . . 

. .. 51 ,77 

. .72 

. .. .. 51 ,78 

. 36,37,59,65 

. ...... 51 ,79 

. . 51 ,80 

. 79,80 

97