NEIL C. HULIM

and JOHN S. GKA'

A Manual

for the

Study of Meiofauna

SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY NUMBER 78



SMITHSONIAN CONTRIBUTIONS TO

ZOOLOGY

NUMBER 78

Neil c. Huiings A Manual

and John S. Gray r ••

tor the

Study of Meiofauna

SMITHSONIAN INSTITUTION PRESS

CITY OF WASHINGTON

1971
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For tale by the Superintendent of Documents, U.S. Government Printing Office

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International Conference on Meiofauna

1-11 July 1969, Tunis, Tunisia

ORGANIZATION

N. C. Hulings, convener

J. S. Gray, co-convener

R. P. Higgins, co-convener

EDITORIAL STAFF FOR MANUAL

N. C. Hulings

J. S. Gray

A. D. Mclntyre

I. M. Newell

W. Sterrer

J. B. J. Wells

PARTICIPANTS

Dr. C. Clausen

Zoological Laboratory

University of Bergen

Bergen, Norway

Dr. M. T. Gomoiu

"Traian SaVelescu" Institute of Biology

Department of Marine Biology

Constantza, Romania

Dr. J. S. Gray

Wellcome Marine Laboratory

Robin Hood's Bay

Yorkshire, England

Dr. R. R. Hessler

Scripps Institution of Oceanography

La Jolla, California 92037

Dr. R. P. Higgins

Office of Oceanography and Limnology

Smithsonian Institution

Washington, D.C. 20560

Dr. W. D. Hope

Department of Invertebrate Zoology

National Museum of Natural History

Washington, D.C. 20560

Dr. N. C. Hulings

Office of Oceanography and Limnology

Smithsonian Institution

Washington, D.C. 20560

Dr. W. D. Hummon

Department of Zoology

Ohio University

Athens, Ohio 45701

Dr. S. Husmann

Limnologische Flusstation

Max-Planck-Instituts fiir Limnologie

Schlitz, West Germany

Dr. B. O. Jansson

Askolaboratoriet

Trosa, Sweden

Dr. C. Jouin

Laboratoire de Zoologie

Faculty des Sciences

7, Quai St. Bernard

Paris, France

Dr. E. Kirsteuer

Department of Living Invertebrates

American Museum of Natural History

New York, New York 10024

Dr. H. Kiihl

Institute fiir Kiisten- und Binnenfischerei

Laboratorium Cuxhaven

Cuxhaven, West Germany

Dr. P. Lasserre

Institut de Biologie Marine

University de Bordeaux

33 Arcachon, France

in
IV SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Mr. A. D. Mclntyre

Marine Laboratory

Aberdeen, Scotland

Dr. F. Monniot

Museum National d'Histoire Naturelle

Laboratoire de Biologie des Invert£bre"s Marins

Paris, France

Dr. I. M. Newell

Department of Life Sciences

University of California

Riverside, California 92502

Dr. W. Noodt

Zoologisches Institut der Universitat

Hegewischstrasse 3

Kiel, West Germany

Dr. O. H. Pilkey

Department of Geology

Duke University

Durham, North Carolina 27708

Dr. L. W. Pollock

Systematics-Ecology Program

Marine Biological Laboratory

Woods Hole, Massachusetts 02543

Dr. J. Renaud-Mornant

Museum National d'Histoire Naturelle

43, rue Cuvier

Paris, France

Dr. F. Riemann

Institut fur Meeresforschung

Bremerhaven, West Germany

Dr. W. Sterrer

Bermuda Biological Station

St. Georges West, Bermuda

Dr. B. Swedmark

Kristineberg Zoological Station

Fiskbackskil, Sweden

Dr. G. Uhlig

Biologisches Anstalt Helgoland

Helgoland, West Germany

Dr. J. B. J. Wells

Department of Zoology

University of Aberdeen

Aberdeen, Scotland

Dr. W. Westheide

II Zoologisches Institut der Universitat

Gottingen, West Germany

Dr. R. Williams

Oceanographic Laboratory (S.M.B.A.)

Edinburgh, Scotland
Foreword

The term meiofauna has been in use for only a relatively brief period. It was

coined by Mare in 1942 to describe metazoans (and Foraminifera) too small to

be retained on the 1-mm sieves used by marine biologists for the extraction of

bottom fauna from samples of sediment when analyzing benthic communities.

Thus, meiofauna refers only to a size class between macrofauna and microfauna.

The term is now used for benthos and phytal fauna, but not for planktonic

organisms, which are treated traditionally as a separate category and by special

methods.

The variation in form and the biological complexity of the meiofauna make

it difficult to determine upper and lower limits in terms of body size which can be

applied to all taxa. For the time being, however, the lack of a definitive and generally accepted definition is less important than standardized methods of work for

the taxonomists and ecologists studying these organisms.

Practically all classes of the Metazoa, including the smaller species of macrofauna, are represented in the meiofauna, while the Protozoa are represented by

their largest forms, e.g., Foraminifera and Ciliata. The smaller protozoans are

generally considered as microfauna. Gnathostomulida, Gastrotricha, Kinorhyncha, and Tardigrada are some groups of metazoans that are known exclusively

from the meiofauna, while Mollusca, Brachiopoda, Echinodennata, and Ascidiacea are examples of groups represented only by a very few species.

Mclntyre (1969) uses the term "permanent members" for species belonging to

the meiofauna during the whole of their life cycle. The meiofauna also includes a

large number of "temporary members," i.e., larvae and juvenile stages of species

that, as adults, belong to the macrofauna.

If we refrain from an exact division according to type of biotope but are satisfied to compare the three main groups—mud, interstitial sand, and phytal meiofauna—we find certain morphological and biological features for each of these

categories, reflecting the adaptation of the organisms to particular environmental

conditions. These adaptations undoubtedly are most marked in the interstitial

fauna of sand. Since Remane's pioneering work in the 1920s the interest of psammonologists has been focused on the study of such adaptations. Within several

systematic groups a greater proportion of elongated forms is found dian in types

other dian the interstitial environment. The development of various types of

adhesive organs is another adaptation found in many groups belonging to the

interstitial fauna. Other adaptations are common for the meiofauna as a whole—

for example, reproductive adaptations, which are related to the relatively small

number of gametes that small animals produce. Copulation, hermaphroditism,

and the formation of egg cocoons, as well as brood protection, improve the prospects of development and ensure the survival of the populations.

The discovery of new groups of animals, particularly in the interstitial sand

fauna—often of high taxonomic rank (Gnathostomulida, Mystacocarida, Actinulida, Macrodasyoidea, Acochlidiacea)—undoubtedly has been one of the major

contributions to systematic zoology during the present century.

Several research workers regard the many aberrant forms, morphological specializations, and appearance of new organs in the meiofauna as having evolved
vi SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

under the influence of the unique environment. Neoteny seems often to have

been one of the reflections of evolution; it has resulted in a simplified organization

compared with the presumed ancestral form and may be manifested in the loss

of organs. In addition, neoteny has led to diminution of body size. Specializations

such as the appearance of new organs, loss of organs, and reduction in body size

have emphasized the systematic isolation of the aberrant species. It is difficult,

for example, to demonstrate unequivocally the relationship between the Gnathostomulida and any definite class of the other Metazoa, or to show the affinity of

the Archiannelida with a definite systematic group within the Polychaeta. The

phylogenetic relationships and the taxonomic problems resulting from regressive

evolution must remain unsolved until we discover new links in the often rather

fragile morphological series. The definition of phylogenetic relationships is today

one of the motives behind the work of many meiobenthologists in the fields of

systematics and morphology. The discovery during recent years of a large number

of new species of Gnathostomulida, Gastrotricha, Nematoda, and Ascidiacea suggests that systematic zoology may still have much to gain from the studies of

meiofauna, especially with the opportunities now available to meiobenthologists

to take part in the exploration of the deep sea.

Regardless of the increased interest in meiofauna today, the systematicsmorphological field is not yet well covered. Most of the studies to date have been

in Europe, thereby leaving much of the rest of the world unstudied. Furthermore,

there must be much more research, not only to meet the general needs of systematics and morphology but for progress in ecology.

In the sphere of ecology, Remane's early work also has been of great importance. But it has been during the past ten or fifteen years that the meiofauna has

attracted widespread attention of ecologists. Important work has been done by

Boaden (1962, 1963a), Gray (1965, 1966a, b), Jansson (1962,1966, 1967a, d, 1968),

Renaud-Debyser (1963), and others concerning, for example, the relation of

vertical and horizontal migration to temperature, salinity, and other ecological

factors. Such studies have been made in the field and experimentally in the

laboratory.

In the field of meiofauna ecologists have interesting and important tasks to

perform. Our knowledge of the dynamics of populations of meiofauna is very

limited at present. The same is true of our knowledge of the part played by the

meiofauna in the organic production within the various zones of the sea. The

studies already made usually have been concerned with single species, as a rule

within the tidal zone. It is also important to increase our knowledge of trophic

relations and the position of the meiofauna in the food chains.

Meiobenthology may be expected to become enriched by several new fields of

research. One such field may be the use of species in the meiofauna as test organisms for the study of the biological effect of components of water pollution. In

ecological biotest laboratories it is important to study the biological effect during

the whole life cycle of the test organism. The rapid development of many of the

species in the meiofauna and the relative facility with which they can be cultivated make meiofauna excellent test organisms. The very small size of meiofauna,

with organs sometimes composed of a very limited number of cells but which

nevertheless are capable of coordinated functions fully comparable to that of

much larger organisms, should interest immunologists, biochemists, and physiologists.
NUMBER 78 vii

The idea of this manual resulted from the International Conference on Meiofauna convened in Tunisia, 1-11 July 1969, by the Mediterranean Marine Sorting

Center at Salammbo and by the Smithsonian Institution. Twenty-eight meiobenthologists were given opportunities to confer and take part in valuable discussions

contributing to this publication. On the basis of their own experiences, the contributors to this manual have recommended certain methods for the study of the

meiofauna. We hope that after being utilized for a few years in different laboratories the manual will be improved and augmented. In any case, we hope it will

be of value to both specialists and students, and will contribute toward a standardization of methods for the study of the meiofauna.

BERTIL SWEDMARK

Kristineberg Zoological Station

Fiskbackskil, Sweden

Contents

Page

GENERAL DESCRIPTION OF HABITAT 3

1.1 Climate 3

1.2 Geography 3

1.3 Geology 3

1.4 Hydrography 3

1.5 Physiography 3

ENVIRONMENTAL ANALYSES 3

2.1 Interstitial Water Samples 4

2.2 Temperature 4

2.2.1 Instantaneous Readings 4

2.2.2 Continuous Recordings 4

2.2.3 Sublittoral Measurements 4

2.3 Salinity 4

2.3.1 Wheatstone Bridge 4

2.3.2 Salinity—Temperature Bridge 5

2.3.3 Other Techniques 5

2.3.4 Sublittoral 5

2.4 pH and Redox 5

2.5 Oxygen 5

2.5.1 Oxygen Concentration 6

2.5.2 Oxygen Availability 6

2.5.3 Sublittoral 6

2.6 Water Saturation 7

2.6.1 Speedy Moisture Tester 7

2.6.2 Displacement Technique 7

2.7 Hydrogen Sulfide 7

2.8 Granulometry 7

2.8.1 Grain Size 7

2.8.2 Roundness 9

2.9 Mineralogy 9

2.9.1 Calcium Carbonate Content 9

2.9.2 Mineralogical Analysis 9

2.10 Porosity 10

2.10.1 Pore Space 10

2.10.2 Spatial Relationship 11

2.10.3 Permeability 13

2.11 Organic Matter 14

2.12 Chlorophyll a 14

2.13 Bacteria 15

2.14 Light 15

2.15 Nutrient Analyses 16

GENERAL COLLECTION TECHNIQUES 16

3.1 Preliminary Survey 16
SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Page

3.2 Littoral Sampling 16

3.2.1 Distribution Sampling 17

3.2.2 Density Sampling 17

3.2.3 Corers 17

3.2.3.1 Simple Tubular Corers 17

3.2.3.2 "Ring-lined" Corers 19

3.2.3.3 Demountable Corers 19

3.2.3.4 "Meiostecher" 20

3.2.4 Sampling the Deep Layers of Beach Sand 20

3.3 Sublittoral Sampling 20

3.3.1 SCUBA 20

3.3.2 Remote Sampling 21

GENERAL EXTRACTION TECHNIQUES 24

4.1 Field Techniques 24

4.1.1 Sand 25

4.1.2 Mud 25

4.1.3 Shipboard Techniques 25

4.2 Laboratory Techniques 26

4.2.1 Sand 26

4.2.1.1 Uhlig Seawater Ice Technique 27

4.2.1.2 Anaesthetization Technique 27

4.2.1.3 Deterioration Technique 27

4.2.1.4 Boisseau Technique 27

4.2.2 Mud 29

4.2.2.1 Quantitative Extraction of Total Fauna 29

4.2.2.2 Swedmark Technique 30

4.2.2.3 Bubbling Technique 30

4.3 Counting and Observation Efficiency 30

GENERAL TAXONOMIC PROCEDURES 30

5.1 Examination of Live Material 30

5.1.1 Techniques 31

5.2 Fixation 31

5.2.1 Fixation of "Soft" Fauna 32

5.2.1.1 Whole Mount Fixation of "Soft" Fauna 32

5.2.1.2 Fixation of "Soft" Fauna for Histology 32

5.2.2 Fixation of "Hard" Fauna 32

5.2.2.1 Fixation of "Hard" Fauna for Histology 32

5.3 Preservation 33

5.3.1 Preservation of "Soft" Fauna 33

5.3.2 Mounting of "Soft" Fauna 33

5.3.2.1 Water Mounting 33

5.3.2.2 Mounting after Dehydration 33

5.3.2.3 Histological Preservation of "Soft" Fauna 33

5.3.3 Preservation of "Hard" Fauna 33

5.4 Curation 33

5.5 Presentation of Data 34

5.5.1 Minimum Requirements 34

5.5.2 Descriptions 34
NUMBER 78 x j

Page

6 COMMENTS ON SELECTED TAXA 35

6.1 Foraminifera 35

6.2 Ciliata 36

6.3 Cnidaria 37

6.4 Turbellaria 37

6.5 Gnathostomulida 38

6.6 Nemertina 39

6.7 Gastrotricha 39

6.8 Kinorhyncha 40

6.9 Priapulida 41

6.10 Netnatoda 41

6.11 Bryozoa 42

6.12 Brachiopoda 43

6.13 Archiannelida 43

6.14 Polychaeta 44

6.15 Oligochaeta 45

6.16 Mollusca 46

6.17 Mystacocarida 47

6.18 Ostracoda 47

6.19 Copepoda 48

6.20 Palpigradida 49

6.21 Halacaridae 49

6.22 Tardigrada 50

6.23 Echinodermata 50

6.24 Tunicata 51

7 CULTURE AND EXPERIMENTAL TECHNIQUES 51

7.1 The Experimental Population 52

7.2 Culture Techniques 52

7.3 Experimental Techniques 53

7.3.1 Field Techniques 53

7.3.2 Laboratory Techniques 53

7.3.2.1 Tolerance Experiments 53

7.3.2.2 Preference Experiments 54

7.3.3 Physiological Techniques 54

7.3.3.1 Osmoregulation and Ion Regulation 54

7.3.3.2 Oxygen Reactions 54

7.3.3.3 Metabolism 55

7.3.3.4 Sense Physiology 55

8 BLOTOPES OTHER THAN SEDIMENT 5 5

9 LITERATURE CITED 56

10 APPENDIX 79
ABSTRACT

Neil C. Hulings and John S. Gray. A Manual for the Study of Meiofauna.

Smithsonian Contributions to Zoology, number 78, 83 pages, 1971.—Procedures

and methods for the study of meiofauna are presented in terms of general description of the habitat, environmental analyses, general collecting and extracting techniques, general taxonomic procedures, comments on selected taxa, and culture and

experimental techniques. Biotopes other than sediment are considered briefly and

sources of equipment and materials for the study of meiofauna are listed. A total

of 361 references dealing with various aspects of meiofauna are given.
Neil C. Hulings

and John S. Gray

A Manual

for the

Study of Meiofauna

An International Conference on Meiofauna was

held in Tunis, Tunisia, 1-11 July 1969. Twentyeight invited participants from seven countries assembled to review the status of the systematics and

ecology of meiofaunal taxa, the ecology of meiobenthic communities, and techniques of faunal and

environmental analyses. A total of thirty papers

covering the above topics was presented during the

first three days of the conference, and these papers

will appear in a volume of proceedings of the conference. One day was devoted to field and laboratory demonstrations of equipment and techniques.

The last week of the conference was devoted to

committee work. Subcommittees were organized to

provide an exchange of ideas and information between ecologists and systematists. Two ecological

subcommittees, Littoral and Sublittoral, and two

systematic subcommittees, "Soft" Fauna and

"Hard" Fauna were established (the terms littoral,

sublittoral, "soft" fauna, and "hard" fauna are

defined below). The memberships of these subcommittees follow. Littoral subcommittee: B. O. Jansson (chairman), J. S. Gray, W. D. Hummon,

S. Husmann, H. Kuhl, L. W. Pollock, R. Williams.

Sublittoral subcommittee: A. D. Mclntyre (chairman), M. T. Gomoiu, R. R. Hessler, O. H. Pilkey,

J. B. J. Wells. "Soft" Fauna subcommittee: W. Sterrer (chairman), C. Clausen, C. Jouin, E. Kirsteuer,

P. Lasserre, B. Swedmark, G. Uhlig, W. Westheide.

"Hard" Fauna subcommittee: W. D. Hope (chairman), R. P. Higgins, N. C. Hulings, F. Monniot,

I. M. Newell, W. Noodt, J. Renaud-Mornant,

F. Riemann. The subcommittees met separately and

Neil C. Hulings, Department of Biology, American University

of Beirut, Beirut, Lebanon. John S. Gray, Wellcome Marine

Laboratory, Robin Hood's Bay, Yorkshire, England.

then they met collectively to integrate the results

of their efforts and incorporate them into this

"Manual for the Study of Meiofauna."

This manual is intended to be multipurpose.

First of all, it is intended as a guide for both the

beginning and the established researcher, whether

ecologist or systematist, in the study of meiofauna.

It is of primary importance that the ecologist be

aware of the problems of the systematist, and vice

versa, if studies of meiofauna are to be meaningful.

The systematist may find the treatment of his particular taxon rather narrow in scope since emphasis

has been placed on techniques of sampling, extracting, fixing, and preserving rather than on detailed

systematics. Such information is important, since

it is not always realized that samples and specimens

must be dealt with in different ways depending

upon the taxon or taxa of greatest concern. The

ecologist, by the same token, must be aware of the

problems of all systematists, especially if he is studying the total meiofauna.

The manual is also intended to provide a standardization for various techniques so that one can

compare data from one geographic area with that

from another and data collected by one investigator

with that by another. At present, one finds great

heterogenity in sampling, reporting of data, etc.,

so that meaningful comparisons are difficult.

Though the manual is brief, the techniques are

described in detail and pertinent references to the

literature are given. Many of the techniques were

developed by the contributors reporting on them.

Most have been extensively tested and all have been

found to be successful.

No attempt has been made to produce a rigid

definition of "meiofauna." The term generally is

1
SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

used to refer to animals, mostly metazoans, that can

pass a 1.0-mm to 0.5-mm screen. The term "littoral" is here defined as intermittently submerged

sedimentary environments (tidal and atidal), including beaches and mud flats; "sublittoral" is

defined as continuously submerged sedimentary environments regardless of depth. Thus, the emphasis

of the manual is on sedimentary meiofauna, whether

it be sand or mud or mixtures thereof regardless of

depth of water. It is hoped that a revision of the

manual, anticipated within two to three years, will

include biotopes other than sediment.

The distinction between "soft" fauna and "hard"

fauna is made subjectively on the basis of the resistance of the integument to mechanical damage.

The "soft" fauna includes taxa with a soft integument and usually a great ability to change shape

and to contract considerably. The taxa grouped

under "soft" include Ciliata, Cnidaria, Turbellaria,

Gnathostomulida, Nemertina, Gastrotricha, Archiannelida, Polychaeta, Oligochaeta, Mollusca, and

Echinodermata. The "hard" fauna consists of taxa

whose representatives contract only slightly, if at

all, and possess a shell or an inelastic cuticle.

The "hard" fauna taxa include Foraminifera,

Kinorhyncha, Priapulida, Nematoda, Bryozoa,

Brachiopoda, Mystacocarida, Ostracoda, Copepoda,

Palpigradida, Halacaridae, Tardigrada, and Tunicata. Though this division may seem artificial, it

fits quite well into methodological groupings. Systematists and ecologists will find that, in general

certain methods can be applied to "soft" fauna and

others to "hard" fauna.

The literature on meiofauna has increased severalfold within the last few years but it is beyond

the scope of this manual to include all the pertinent

references. There are, however, several published

works that are considered basic references on the

ecology and systematics of meiofauna, including Ax

(1966, 1969), Delamare Deboutteville (1960), McIntyre (1969), Remane (1933, 1940, 1952), and

Swedmark (1964). Taken together, these works provide a comprehensive bibliography on meiofauna

and should be consulted as basic sources of information for all phases of the study of meiofauna. An

excellent source of current literature on meiofauna

is Psammonalia, a newsletter of the Association

of Meiobenthologists.1 Past issues of Psammonalia

have included comprehensive bibliographies on

several meiofaunal taxa. The "Proceedings of the

First International Conference on Meiofauna"

(Hulings, 1971a) summarizes the present status of

our knowledge of meiofauna, especially systematics

and ecology. "The Study of Marine Benthos"

(Holme and Mclntyre, in press), International Biological Program Handbook, No. 16, deals with

meiofauna in general terms. A survey of the literature on meiofauna shows that most of our present

knowledge has come from the studies of European

investigators. Thus, there remains much to be done

in both systematics and ecology of meiofauna

throughout the rest of the world.

We are indebted to many persons and organizations for the success of the conference. The Government of Tunisia, including the National Scientific

and Technical Institute for Oceanography and

Fisheries, the Department of Higher Education and

Scientific Research, and the Ministry of Foreign Affairs, endorsed the conference and assisted in many

ways. The American Embassy, Tunis, was most

helpful on numerous occasions. The staff of the

Mediterranean Marine Sorting Center—especially

Dr. H£dia Baccar, Mrs. Genevieve Dargouth, and

Mr. Mohammed Shili—freely provided assistance in

making local arrangements and in the initial preparation of this manual. Funding for the conference

was made possible through the Office of International Activities of the Smithsonian Institution,

Washington, D.C. The management of the Hilton

Hotel, Tunis, assisted in many aspects of the local

arrangements. Many persons in the Departments of

Invertebrate Zoology and Paleobiology of the National Museum of Natural History of the Smithsonian Institution have been extremely helpful in

many ways. Special gratitude is owed to Dr. Ray

Manning and his staff for encouragement and review of the manual and to Mrs. Roberta K. Smith

who prepared the section on Foraminifera. To these

organizations and all of the persons involved, we

express our sincere appreciation.

1

 Inquiries for information on Psammonalia should be

addressed to the editor, currently Dr. John S. Gray, Wellcome Marine Laboratory, Robin Hood's Bay, Yorkshire,

England.
1. General Description of Habitat

A careful definition of the study area in ecological terms should precede every

investigation. For the systematist this means that localities can be found again

without difficulty. Simple collecting of specimens, if accompanied by sufficient

data, can be of great interest to the ecologist. Finally, the ecologist should record

all the data available in order to obtain a complete analysis of the correlation

between fauna and parameters. The following should be noted.

1.1. Climate.—Season, date, illumination and insulation, and water temperature. In coastal areas, weather conditions prior to sampling, noting precipitation,

air temperature, and wind speed and direction.

1.2. Geography.—Latitude and longitude, local name of site, distance and direction to nearest town or physiographic landmark appearing on maps or charts,

means of determining location, and general nature of the physiographic province

(e.g., coastal, continental margin or shelf, abyssal plain, midocean ridge). Littoral

characterizations should include extent of beach and continuity with adjacent

areas.

1.3. Geology.—Substratum, nature of shoreline (deposition or erosion), sediment source and availability, type and structure of rock in adjacent outcrops.

1.4. Hydrography.—Littoral: tidal amplitude, wave height, pattern of daily

tides. Littoral and sublittoral: frequency, intensity and direction of storms, current

direction and rate of flow, effect of each on beach or bottom sediments. Sublittoral:

all available on-station water measurements.

1.5. Physiography.—Littoral: beach width, contour pattern, profile, direction

and exposure, and presence of features such as berms, tide pools, rock ledges, offshore islands, bars and kelp beds. Sublittoral: bottom irregularity, contour patterns,

profile, presence of seamounts, trenches, ripple marks, ridges, canyons, and other

characters.

2. Environmental Analyses

In most cases the same environmental features can be considered in littoral and

sublittoral studies. These include physiography, hydrodynamic regime, temperature, salinity, oxygen availability, granulometry, mineralogy, and sediment porosity. Regional primary productivity, organic matter, bacteria, and other biological

constituents of the environment are important for community analysis. In the

littoral, water saturation and permeability are of prime importance. Amount of

light and benthic chlorophyll should be determined in both environments. In the

sublittoral, depth and amount of particulate organic matter in the water column

may be significant. In the sublittoral suitable for SCUBA, littoral equipment can

be used with suitable modification, thus giving continuity with littoral studies. In

deeper water remote gear must be used.

If the objective of the study is synecological—such as a comparison of polluted

and unpolluted, arctic and tropical areas—faunal and environmental analyses can

be restricted to periods of extremes, i.e., winter and summer or rainy and dry

seasons. If the objective is autecological, detailed seasonal measurements should

be made. Any study employing only a short period of environmental analysis runs

the constant danger of ignoring limitations which occur only during other times
4 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

of the year. In addition, it may not be the absolute value of the environmental

parameter which is important to the organism, but the degree of its stability and

rate of change. Most of the environmental features mentioned are stable in the

deep sea; however, changes in temperature, oxygen, currents, and paniculate organic matter, with depth, in the water column should be measured.

Wherever possible it is best to make measurements directly in the field.

Whether in field or laboratory, the size of the probes used or of the samples taken

must always be related to the size of the animals being investigated in order to determine the environmental conditions that immediately surround the organism.

This is especially important when vertical gradients are studied because of the

variations that often exist.

The general littoral environment should be studied at high tide as well as low

tide. In both littoral and sublittoral, attention should be paid to the characteristics

of the water column. Many features of the sublittoral are difficult to study due to

the lack of suitable instruments that provide the desired data. Gomoiu (1971),

Husmann (1971), Jansson (1971), Mclntyre (1971) and Pollock (1971) have

reviewed various aspects of environmental and faunal analysis in the littoral and

sublittoral.

2.1. Interstitial Water Samples.—When collecting interstitial water samples

care must be taken to prevent contamination from areas other than those to be

sampled. For example, large samplers disturb the sediment, and water from surface layers may run down the side of the sampler and mix with the sample being

taken. Thus, a sampler such as that of Brafield (1964) and Tilzer (1967), or a

microsyringe is most suitable, as mixing of different water levels is avoided. If a

microsyringe is used, the tip should be modified to include a sintered glass screen

which prevents particles from blocking the syringe. Extraction of water samples

must be slow to avoid the entrance of air bubbles and samples should be stored in

clean glass containers or plastic containers (see Section 2.15). The accuracy of

some chemical analyses, however, may be impaired by storage. Samples of the

adjacent seawater (at low tide) or the overlying seawater (at high tide) should

also be taken.

2.2. Temperature.—A number of portable thermistor-type temperature probes

accommodating up to twelve small probes simultaneously are available (see

Appendix, Section 10.1.1). A tubular metal probe with a small sensitive area is

best for this work (e.g., Spembly Technical Products probes). Varying cable

lengths can be obtained. Periodic checks of the probe should be made to test for

age deterioration.

2.2.1. Instantaneous Readings.—These can be made by inserting the probe

vertically into the sediment or horizontally into the side of a hole dug in the

sediment.

2.2.2. Continuous Recordings.—Most telethermistors can be used for continuous recordings in conjunction with a chart recorder (see Appendix, Section

10.1.2). Probes are buried in the sediment, and up to 20 readings can be monitored

simultaneously. Suitable instruments are listed in the Appendix (Section 10.1.1).

2.2.3. Sublittoral Measurements.—It is not necessary to make the measurement in the sediment; readings from the adjacent water column are sufficient.

Probes or reversing thermometers can be employed.

2.3. Salinity.—As the international standard for salinity determinations is

based on conductivity, this method is preferable to others.

2.3.1. Wheatstone Bridge.—This method of determining conductivity is
NUMBER 78 5

cheap and simple and requires only 0.5 to 1 ml of interstitial water. Water samples

should be collected following the methods in Section 2.1.

The smallest type of conductivity cells are equipped with a bulb pipette

through which a water sample can be drawn into the cell. The cell is capped and

immersed in a water bath. As temperature control is of the utmost importance, all

solutions should be kept in a constant-temperature bath. The electrodes should be

washed in distilled water, dried, and immersed in a spare portion of the sample

before being placed in the aliquot to be sampled.

The International Oceanographic Tables, prepared by UNESCO (see Appendix, Section 10.1.3), recommend a conductivity standard for salinity determinations. The best method of making salinity estimations would, therefore, be the

use of these tables. The conductivity of standard seawater (obtained from the

Laboratoire Oc^anographique, Charlottenlund, Denmark) should be determined

first, followed by the solution to be tested. The ratio of these two conductivities is

translated to salinity at the appropriate temperature in the UNESCO tables. A

suitable and relatively cheap commercial instrument is available (see Appendix,

Section 10.1.4.3). A circuit diagram is shown in Figure 12. A list of manufacturers

of components for personal assemblage is given in the Appendix (Section 10.1.4) .

2.3.2. Salinity-Temperature Bridge.—A small portable salinometer (see Appendix, Section 10.1.5) designed primarily for estuarine and coastal hydrographic

use is suitable for determinations on 25 to 50 ml of water. Salinity can be read

from 0 to 38 ppt and temperature from 0° to 35° C. Samples of interstitial water

(obtained by the methods outlined in Section 2.1), are placed in the chamber of

the probe head and values are read from the scale. The major disadvantage of

this technique is the large volume of interstitial water needed for each determination.

2.3.3. Other Techniques.—Chemical methods are cumbersome and may not

be suitable due to insufficient data on the ionic concentration of interstitial water.

Specific gravity methods are less preferable due to the large volume of water

needed for determinations (see Barnes, 1959, for methods). A new but relatively

untested field method, the Goldberg refractometer (see Appendix, Section 10.1.6),

requires only a few drops of interstitial water.

2.3.4 Sublittoral.—As with temperature, it is sufficient in most cases to measure the salinity of the adjacent water column. It should be stressed, however, that

although the salinity of the interstitial water might be equal to that of the overlying water, the relative ionic composition is likely to be different.

2.4. pH and Redox.—The pH has not been studied in sufficient detail to

evaluate its direct effect on meiofauna. The redox potential (Eh), however, is of

biological importance, particularly in sheltered or atidal beaches. On well-sorted

oceanic beaches there is rarely a change in Eh (and pH) with depth. Low Eh

(and possibly low pH) values may indicate the presence of H2S which may be

toxic, and the Eh may indicate limiting biological processes (e.g., sulfur reduction

occurs at an Eh of below +100 mv).

Eh readings should always be accompanied by pH readings. Thus, meters

that measure both simultaneously are preferable. The instruments (see Appendix,

10.1.7) must be modified to use small probes; otherwise, sediment displacement

will give erroneous readings (see Fenchel, 1969, and Whitfield, 1969, for a full

discussion of the significance of pH and Eh).

2.5. Oxygen.—Two types of oxygen measurements are common—total oxygen

concentration and oxygen availability. Total oxygen concentration can be de-
6 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

termined by titrimetric methods, the use of polarographic or exposed electrodes.

If a polarographic electrode is used, determination of oxygen is dependent on

diffusion of oxygen through a teflon membrane; it requires a temperature compensation as well as water-flow rate in excess of 1 cm/sec around the electrode.

Total oxygen also can be measured by a variety of exposed electrodes such as

platinum and thallium, but these have not been evaluated by the participants.

Oxygen availability is measured by a stationary, naked platinum electrode

reducing the oxygen diffusing to it. The latter measurement incorporates ambient

temperatures, water-flow rates, and other parameters affecting oxygen diffusion.

In this way the electrode can be considered to be functioning in a manner comparable to an interstitial animal.

2.5.1. Oxygen Concentration.—Oxygen concentration has been estimated by

chemical titration techniques and by polarographic electrodes. Both macroprocedures and microprocedures for chemical determination of oxygen are described

in detail by Barnes (1959). Membrane-covered polarographic electrodes measure

oxygen content (in ppm) and include temperature compensation. Carey and

Teal (1965) discuss the construction and reliability of these instruments.

2.5.2. Oxygen Availability.—This technique, pioneered by Lemon and Erickson (1952) for soil, has been applied to the marine environment by Jansson

(1966, 1967b) and Fenchel, Jansson, and von Thun (1967). Oxygen diffusion

rates are measured by an exposed platinum electrode at a constant voltage of

0.8 v. Values are measured as ^iA/cm

 2 electrode surface/min. The source, a description of the circuitry used in this instrument, and the procedure for calculation

of oxygen are given in the Appendix (Sections 10.1.8, 10.3 and Figure 13).

Care must be taken when using this technique. The entire electrode surface

must be covered by a water film, otherwise oxygen availability will be underestimated. If multiple recordings at the same depth close to the sediment surface

give highly variable results, it is likely that water content is inadequate to allow

measurements of oxygen availability. "Poisoning" of the electrode may occur

when positively charged particles or colloids have an isoelectric point higher

than the ambient pH precipitate on the electrode surface, thus decreasing its

sensitivity. Detection of this phenomenon may be based on sensitivity tests in

water of high oxygen availability following exposure of the electrode to an area

of potential poisoning. Since electrodes are inexpensive and easily made, it is wise

to include several of them in field supplies. As the diffusion is highly timedependent, readings must be made when equilibrium is attained; e.g., after 3

minutes.

Enckell (1968) and others have shown, however, that above a certain current

velocity the availability of oxygen does not increase with increased velocity. The

sensitivity to current velocity also may be dependent on the initial oxygen concentration. Caution should be exercised, therefore, in interpreting the data derived from the technique described above.

In the littoral, measurements are easily made by digging a hole and inserting

the electrodes to the desired depths in the water. As the effect of length of the

wires or distance between the electrodes is negligible, sand beneath the water table

can be reached by attaching the sensor to a stick. A diver can employ this technique

in the sublittoral by means of an instrument kept on a boat or the shore.

2.5.3. Sublittoral.—Where remote gear is used, particularly in the deep sea,

the main difficulty is obtaining the sample or measurement from the water just

above the bottom or in just the top layer of sediment. For this reason, water-
NUMBER 78 7

sample analysis is unsatisfactory. The oxygen availability electrode offers a

promising solution.

2.6. Water Saturation.—Water saturation varies continuously throughout the

tidal cycle; thus, the percentage saturation should be quoted for the time at which

faunal samples are collected.

2.6.1. Speedy Moisture Tester.—The most accurate field technique is the

Speedy Moisture Tester (see Appendix, Section 10.1.9). This technique, which

requires 1.5 or 3 g of sediment, measures gas pressure produced on shaking the

sample with an absorbent. The gas pressure is proportional to the moisture content.

2.6.2. Displacement Technique.—A less-accurate method is the displacement

technique of Hummon (1969). To determine water saturation of pore space, 5

cm

 3 of water is first placed in a 10 cm 8 graduated cylinder. Sand is then allowed

to settle slowly through the water until the height of the sand column reaches

the 5 cm

 3 level. The cylinder is agitated frequently to increase packing. Five cm 3

of fully saturated sand displaces the water surface 5 cm3 upward to the 10 cm

 3

level, whereas 5 cm3 of dry sand having 60 percent grain and 40 percent pore

volume would only displace the water surface 3 cm

 3 upward to the 8 cm 3 level.

Porosities of field samples can be tested by oven-drying the sand and then determining the minimum displacement value. This minimum final value represents

0 percent water saturation of the pores and the upper value 100 percent saturation.

Intermediate values measure water saturation of the pores proportionately in

percent, and readings to the nearest 2 percent saturation can be obtained.

2.7. Hydrogen Sulfide.—The presence of hydrogen sulfide in sediments indicates areas of high organic decomposition and usually is associated with a lack of

oxygen. Nevertheless, sediments containing hydrogen sulfide may have a characteristic and rich meiofauna which hitherto has been largely ignored. Thus,

measurements of hydrogen sulfide concentration may indicate the extent of this

habitat.

A simple and rapid measure of the hydrogen sulfide content of a sediment

can be made using a Hach H2S test (see Appendix, Section 10.1.10). This test will

detect quantities as little as 0.1 ppm and can be carried out in the field. More

sophisticated techniques are being developed; e.g., the sulfide electrode of Berner

(1963) and the potentiometric titration technique of Fenchel (1969) .

2.8. Granulometry.

2.8.1. Grain Size.—As a general guide, a sieve 3.75 cm in diameter can

process 15 g of sediment, a sieve 7.5 cm in diameter can process 30 g, and a sieve

15 cm in diameter can process 60 g.

Because of rapid horizontal and vertical changes of grain size in the microhabitat, it is desirable to use the same sample for both faunal and grain size

analysis. In order to retain the physical characteristics of the sample and to make

the results comparable with other investigations, the total sample must be studied.

The coarse fraction should not be omitted from consideration.

Initially the sediment sample should be wet-sieved through a 62-/* sieve using

only water of the same salinity as the sample area. This procedure separates the

"fines" (silt and clay sizes) which, if sufficiently abundant, must be analyzed by

procedures different from those used for the sand sizes. Care should be taken during

wet-sieving to prevent crushing of fragile, small calcareous skeletons or the breakdown of fecal pellets. This is a particularly serious problem with deep-sea biogenic

sediments.

The fine sediment fraction (passing the 62-/x sieve) needs special treatment
SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

TABLE 1.—Phi-mm equivalents and suggested sieve series.

U.S. Standard

Sieve Mesh No.

A.S.T.M.

Sieve No. Opening in mm Phi Scale

Size Class

(Wentworth)

5

7

10

14

18

25

35

45

60

80

120

170

230

10

18

35

60

120

230

4.00

2.83

2.00

1.41

1.00

0.71

0.500

0.350

0.250

0.177

0.125

0.083

0.0625

-2. 0

—1.5

-0. 9

-0. 5

0.0

0.5

1.0

IJ5

2.0

2.5

:i.o

S.5

4.0

Granule

Granule

Very coarse sand

Very coarse sand

Very coarse sand

Coarse sand

Coarse sand

Medium sand

Medium sund

Fine sand

Fine sand

Very fine sand

Very fine sand

using the pipette method (see Folk, 1968, or Griffiths, 1967, for methodology). In

many sediments the fine fraction may be important in blocking pore spaces.

Mud samples may include abundant fecal pellets. Such paniculate material

is delicate, and an estimate of its relative quantity is required before dry-sieving.

Dillon (1964) has given a technique for separating fecal pellets and organisms

from sand.

Samples dominated by biogenic materials usually are analyzed in toto on the

assumption that calcareous skeletal material is a granulometric component of the

bottom sediment, just as are quartz grains or fecal pellets. For example, Globigerina tests in Foraminifera ooze are sand-size grains.

The sand fraction remaining in the 62-/i sieve should be washed with many

changes of fresh water to remove the salt and then dried at 105° C. The sample is

sieved through a series of screens at 0.5 phi (r) intervals. Phi-mm equivalents

are shown in Table 1. Ideally, sieves should correspond to the A.S.T.M. series to

avoid confusion. For example, the British Standard sieve 8 is equivalent to

the A.S.T.M. 10. A mechanical shaker should be used where possible for 15

minutes. Longer sieving times may damage the particle structure. It is important

that all samples be sieved for the same length of time. Table 1 shows the suggested sieve series that should be used for sand analysis. If an unusually detailed

analysis is desired, 0.25-phi sieve intervals can be used (Renaud-Debyser and

Salvat, 1963). Cumulative dry-weight percentages should be plotted on probability

paper. The graphic mean, Mz (Folk, 1968), rather than the often-used median

should be plotted for detailed analysis. It is defined as:

Similarly, the best measure of uniformity of sorting is the inclusive graphic

standard deviation, a / (Folk, 1968):

. <f> 84-<h 16

4

<f>95-d>5

6.6

The asymmetry or skewness of the grain-size distribution pattern can be measured

by the inclusive graphic skewness, SKX (Folk, 1968).

SK _<j> 16+^84-2^5 0 4>5-f y 95-2y 50

1

 ~~ 2 (</> 84- £ 16)

 + 2 (^ 95- ^ 5)
HUH

SPHfMCITY

LOW

SPHERICITY

VERY

ANtULAR

tilt- IHi

ANtULAR ANtULAR RtMKt

FIGURE 1.—Scale of roundness for quartz grains. (From Powers, 1953.)

VCLLRtUNtEt

All the points used in the above equations are obtained from the cumulative

frequency curve plotted with phi scale intervals.

2.8.2. Roundness.—This parameter is of importance in pore space determinations and also in determining the distribution of attached organisms and organic

material on grains (Meadows and Anderson, 1966; Webb, 1969).

The scale of grain roundness of Powers (1953) for quartz grains is shown in

Figure 1. To determine the roundness of a sample, fifty or more grains are assigned

to one of the classes (very angular, angular, etc.) by comparison with the photographs. Average roundness of the sample is determined by multiplying the number

of grains in each class by the geometric mean of that class (0.15 for very angular,

0.20 for angular, etc.) and dividing the sum of the products by the total number of

grains analyzed from each sample.

Calcium carbonate particle roundness can be classified according to the scale

of Pilkey, Morton, and Luternauer (1967), shown in Figure 2. Five roundness

classes are recognized. Class 0 (not shown) includes whole shells. Class 1 fragments

exhibit no apparent rounding of edges and corners and appear as though freshly

broken. Classes 2 and 3 are gradations between Class 1, fresh fragments, and Class

4, highly rounded, usually polished and essentially comerless fragments. The

particles are classified by visual comparison (usually under a microscope) with

the figures in the scale.

2.9. Mineralogy.

2.9.1. Calcium Carbonate Content.—The technique of Gomoiu and Gomoiu

(1968) can be used for a precise determination of CaCO3 content. A known

weight of sand can be digested with a measured excess amount of dilute acid.

The reduction in strength of the acid, which is a measure of the quantity of

CaCO3, is then measured by titration with NaOH. A less-accurate technique is

to weigh a sample, add 10 percent HC1, and weigh again after dissolution of the

CaCO8.

2.9.2. Mineralogical Analysis.—The silicate mineral fraction and the calcium

carbonate fraction usually can be distinguished quickly and easily. Sand-size

particles are mineralogically studied by observation of thin sections of the grains
10 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

FIGURE 2.—Scale of roundness for calcium carbonate particles. (From

Pilkey, Morton, and Luternauer, 1967.)

and by staining techniques. In either case, study under the polarizing petrographic

microscope is necessary. In deep-sea muds, coarser components sometimes may

be identified under the microscope, but generally X-ray diffraction techniques

are needed to determine mineralogy. Identification of major deep-sea sediment

types (pelagic oozes, red clay, etc.) is based mainly on microscopic identification

of coarse components and overall sediment color rather than by detailed mineralogic work. The coarse fraction is isolated for study by standard wet-sieving techniques (section 2.8.1).

2.10. Porosity.—The porosity of a sediment is defined as the volume of water

needed to saturate a given weight of dry sand (usually 100 g) at a given temperature. Fraser (1935), Graton and Fraser (1935), and Webb (1958, 1969) should

be consulted for a detailed discussion of porosity.

2.10.1. Pore Space.—Porosity varies slightly with each tidal cycle due to

particle reassortment by wave action. It is, therefore, important to note the tidal

height, wave action, prevailing weather conditions, and grain size distribution.

The range of porosity values due to various packing arrangements can be determined as follows.

Two samples are needed, and each is dried at 105° C. One sample is placed

in a measuring cylinder, producing loose packing. The other sample is treated

similarly but is agitated so that the sediment is compacted, Water is then added
NUMBER 78 11

under a vacuum (which eliminates trapped air bubbles) until the sediment

sample is totally saturated. The volume of water needed to saturate the first

sample indicates maximal porosity, while the second value indicates minimal

porosity. Porosity also can be measured by weight as long as the sediment is

reasonably homogenous and the specific gravity is determined.

2.10.2. Spatial Relationship.—The epoxy resin embedding technique of

Williams (1971) is useful in this context. Grain-size distribution patterns, porosity, and pore space can be estimated by this procedure.

FIGURE 3.—Impregnation apparatus: a, vessel containing resin; b, dessicator containing anhydrous

CaCl, and the sample (c) for impregnation; d, inlet for air into a and b; e, suction by water

pump via a silica gel vessel inserted into the vacuum line. (From Williams, 1971.)
12 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Undisturbed samples of sediment are collected with the use of a brass

rectangular sampler (10 by 5 by 2.5 cm) with double sliding lids. The sampler

is pressed into the surface of the sediment, and the lower lid is slid into place

after the sediment around the sampler has been cut away. The sampler is then

marked with the orientation, tidal height, and date. The sampler is allowed to

drain, leaving a minimum amount of water, air-dried, and impregnated with

resin. Crystallized salt in the air-dried sample does not hinder impregnation but

tends to maintain the structure of the sediment. The resins used for impregnation

are Crystic 28C and Araldite MY750 (see Appendix, Sections 10.1.11, 10.1.12).

When using Araldite, the inner surface of the sampler should be treated with

Araldite Release Agent QZ12 to aid the release of the hardened impregnated

sample from the sampler. A fluorescent dye, Uvitex SWN, is mixed with the

resin following a technique outlined by Werner (1962). Under ultraviolet light

the dye causes the pore system to fluoresce reddish pink, the quartz fraction red,

and the calcium carbonate blue. The vacuum apparatus for impregnation, a

modification of the method used by Wells (1962), is shown in Figure 3. This

apparatus can easily be constructed using "Quick-fit" glassware. Sufficient vacuum

for complete impregnation of cohesionless sediments can be obtained with the

use of a water pump. Water vapor is prevented from entering the vacuum apparatus by a calcium chloride or silica gel vessel introduced into the vacuum line.

Thin sections are prepared parallel to and at right angles to the surface of

the impregnated sample following the procedure of Dalrymple (1957) and Jongerius and Heintzberger (1963). The impregnated samples are sawed, ground,

polished, and mounted. Sand can be ground to a thickness of about 50 ^ without

cracking the grains. Grinding is finished by hand, using varying grades of carborundum powder. Coverslips are then placed over the thin sections, using

Araldite AY103 as a mountant and taking care to exclude all air bubbles.

For the measurement of the percent volume of pore space in thin sections,

a technique for undisturbed soil developed by Rosiwal and adapted by Swanson

and Peterson (1940) is used. The microscope is focused on the surface of the

section and measurements are taken only in this plane. The slide is traversed

on a mechanical stage parallel to the direction of a graduated scale in the eyepiece, and the distance between the intercepts made by each successive void and

each successive particle is measured. This procedure is repeated in two directions—

parallel to and at right angles to the slide. The total void measurement is then

expressed as percent of the total measurement of void and solid. The method of

measurement is shown in Figure 4. Porosity (P) is determined as follows:

D a+b+c+d

Mr = •

A+a+B + b+C+c+D+d

The measurements of particle and void dimensions are determined systematically by use of a grid system. The grid is formed by traversing the mechanical

stage through a specified number of divisions between each series of measurements. The maximum dimensions in two directions at right angles of each particle,

and including the void, are measured. This is possible because the resin matrix is

transparent and a three dimensional view of the voids and particles can be

obtained. The method of measurement is illustrated in Figure 5. The grain

measurements are given by A^a, B^ , etc, and the void measurements by

aia2, bibs, etc. The procedure is carried out for the sections which are taken

parallel and vertical to the surface of the deposit. Thus obtained are four sets of
NUMBER 78 13

FIGURE 4.—Cross section of a model spherical system showing the method of measuring porosity.

Solid line represents the plane of focus. Measurements are random along the straight grid line.

(From Williams, 1971.)

FIGURE 5.—Cross section of a model spherical system showing the method of measuring voids

and particles. (From Williams, 1971.)

values, representing the maximum dimensions of the voids in three directions

in the deposit, with a duplicate measurement of one of the directions parallel to

the surface. From these measurements the possibility of directional orientation

of the pores or the particles can be checked by analyzing the data statistically.

The measurements of particle and void dimensions can then be plotted against

a Log10 or Log2 scale and the relevant parameters obtained.

2.10.3. Permeability.—Porosity values indicate the volume of pore space

available to the animals, but for meiofauna the permeability (or flow of water

through the sediment) may be of greater significance. For example, fine sands,

in general, hold more water than coarse sands but are less permeable due to the

smaller sizes of the pores.

Permeability can be measured by filling a glass tube (1.5 cm diameter, 70

to 80 cm long) with sand to a depth of 10 cm. Gauze is fitted over the bottom

of the tube. The tube is held vertically and tapped gently to settle the grains.

The time required for a head of 65 cm of water to drain to 15 cm from the
14 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

bottom of the tube (i.e., 50 cm of water through 10 cm of sand) is the permeability (Webb, 1958, 1969). Fraser (1935) used a similar method. Techniques

such as those of Webb and Theodor (1969) for measuring flow rates of interstitial

water in situ may give useful information.

2.11. Organic Matter.—Most current techniques for measuring organic matter are unable to separate the fauna from the analysis, and, thus, are inadequate.

Tietjen (1969), however, by estimating the weight of the meiofauna, made corrections for determinations of the organic content of sediment alone. Whether

what is measured has relevance to the organism remains unknown. In the deep

sea it is likely that most of the organic matter is refractory and unavailable to

the organism, or perhaps even to bacteria.

The Kjeldahl method with ninhydrin finish of Strickland and Parsons (1968)

for determining particulate nitrogen gives reproducible results. The particulate

matter is filtered on a 25-mm Whatman GF/C glass filter. The filtrate (particulate

matter and fauna) is converted to ammonia by the Kjeldahl treatment with

sulphuric acid. The resulting ammonia is determined absorptiometrically using

ninhydrin-hydrindatin reagent without prior distillation. The detailed methodology is outlined in Strickland and Parsons (1968) .

Steele and Baird (1968) determined particulate organic carbon by wet oxidation. They used 0.2 to 1.0 g of sand, removing the particulate matter by filtration

as is done for water samples (Steele and Baird, 1961). Strickland and Parsons

(1968) give methods for the dichromate wet oxidation technique. A similar

method of Maciolek (1962) has been used by Tietjen (1969).

Kvihl (1971) has shown the changes that occur in interstitial water following decomposition of organic matter (including the production of ammonia,

nitrites and nitrates) and related this to sediment characteristics, especially grain

size.

2.12. Chlorophyll a.—Diatoms and other microalgae are important constituents in sediments. Although simple techniques are available for their study,

this phytal component has received little attention. One of the techniques of

Strickland and Parsons (1968) does not estimate phaeophytin; therefore, it is

preferable to other techniques.

The sediment sample is ground in a porcelain mortar for 5 to 10 minutes

and the chlorophyll pigments are extracted by use of 10 ml of 90 percent reagent

grade acetone with the addition of 1 ml MgCO3 suspension (1 g reagent grade

MgCO3/100 ml distilled water). The pigments are extracted for 20 hours in

complete darkness and the extinction values read in a spectrophotometer cell

having a path length of 10 cm but holding 10 ml or less of solution. The concentration of chlorophyll a (mg/liter) is determined by using the formula of

Strickland and Parsons (1968): ll^E^ i - l.SlE^ ^ - O.UEuool (E = extinction at indicated wavelengths after correction for extinction at 75OOA). Care

should be taken to check the extraction efficiency of the acetone. Where chlorophyll values fall below 0.2 mg/liter, the method is less sensitive. Strickland and

Parsons (1968) should be consulted for detailed methodology. Pamatmat (1968)

used the technique described above.

Steele and Baird (1968) estimated chlorophyll a by grinding 1 to 7 g of

vacuum-dried sand in a mortar. The chlorophyll a, extracted with 90 percent

acetone, was estimated by the difference between optical densities at 6300A and

75OOA. The value of 89.31 liter/g cm was used as the coefficient of extinction
NUMBER 78 15

(SCOR-UNESCO, 1964). The SCOR-UNESCO procedure is also given in Strickland and Parsons (1968). Later, Steele and Baird (1968) determined chlorophyll

a by using the Turner fluorometer (Yentsch and Menzel, 1963) and obtained

greater sensitivity for low chlorophyll values. This technique also was used by

Tietjen (1968) and is detailed in Strickland and Parsons (1968) as "Fluorometric

Determination of Chlorophylls."

2.13. Bacteria.—Quantitative assessments of bacteria in sediments using the

standard agar plate techniques underestimate numbers because the medium is

highly selective. Possibly only 10 percent of the bacteria are counted in this

method. Westheide (1968) has studied seasonal variations in bacteria and fungi

by using solid media. Anderson and Meadows (1969) have used a similar technique. Sediment (0.5 to 1.0 g) collected by means of a sterile pipette is added to

10 ml of sterile seawater. Glass beads (Appendix, Section 10.1.20) are added and

the sand is shaken vigorously for 10 minutes. From this suspension 1 ml of solution

is taken and added to 9 ml of sterile seawater. This dilution is repeated three or

four times. From each dilution, 0.05 or 0.1 ml are added to sterile Petri dishes

containing a solid medium (1.2 percent agar, 0.3 percent peptone in aged sterile

seawater or three parts seawater, one part distilled water). The Petri dishes are

incubated at 18° to 22° C for one to three weeks and the number of colonies

forming from each bacterium present on the original sand sample are counted.

The works of Ferguson-Wood (1965), Oppenheimer (1963), and Zobell (1946)

should be consulted for more detailed information.

Respiration rates of sediment samples recently have been shown to give more

reliable estimates of microorganism abundance. Hargrave (1969) used darkened

and transparent cylinders placed on sediment and compared chemically the

changes in oxygen levels in each cylinder to derive a measure of algal respiration.

He used oxygen consumption rate of antibiotic treated and untreated sediment

cores to estimate bacterial respiration. Pamatmat (1968) and Pamatmat and

Banse (1969), using bell jars placed in situ, have recorded sediment respiration

rates with a lead-silver oxygen electrode both intertidally and subtidally.

Meadows and Anderson (1968) used a staining technique to determine the

abundance, distribution, and types of microorganisms (bacteria, blue-green algae,

and diatoms) on sand grains. The samples (1 to 3 ml) are stained within a few

minutes after collection. Each sample is fixed in 2 percent osmic acid for three

minutes, followed by two minutes in Bouin's solution. The sample is then rinsed

in dionized water and stained for six minutes with carbol fuchsin. The samples

can be stored in 10 percent formalin at 0° C. The stained sand grains are then

examined under low-power microscope.

The species of bacteria present may prove to be as significant as their numbers, as shown by Gray and Johnson (1970) in evaluating bacteria as an environmental factor controlling microdistribution of meiofauna.

2.14. light.—Light extinction in sediments is a function of sediment grain

size, and it decreases with a decrease in grain size. Light-extinction rates for

various sediments have been studied by Gomoiu (1967), Gray (1966b), and

Taylor (1964). A simple method is to hold a photocell under a flat-bottomed

glass dish with a strong light source illuminating from above. Sediment is added

slowly, and the change in light intensity is noted with increasing depth of sediment. Care must be taken to ensure that the photocell is sensitive to the wavelengths of light being measured. The wavelength of the light source should be

noted in reporting on this technique.
16 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

The light source should be sufficiently bright to simulate daylight conditions

(not less than 10,000 lux), and readings should be quoted in lux.

2.15. Nutrient Analyses.—Nitrate, phosphate, and silicate are probably the

most important nutrients present in interstitial water. Since all nutrient analyses

need large volumes of water, in excess of 50 ml, they are not particularly suitable

for the analyses of interstitial samples unless auto-analyzers are used. For nitrate,

80 to 90 ml are needed, for phosphate 50 ml, and for silicate 50 ml. The silicate

and phosphate samples cannot be stored in a glass bottle and should be determined immediately. They can, however, be stored in a deep freeze at —20° C.

All the determinations are made using a spectrophotometer. Strickland and

Parsons (1968) give a detailed methodology which should be followed.

Recently, auto-analyzers have been used routinely for nutrient analysis of

seawater (Brewer and Riley, 1965, 1966; Chan and Riley, 1966; Strickland and

Parsons, 1968). These instruments are expensive and require a full-time technician. Many public health and water pollution laboratories have such equipment

and may make these facilities available. This technique has the great advantage

of requiring only small volumes of sample for analysis.

3. General Collection Techniques

Sampling is one of the most difficult problems in meiofaunal investigations. This

is especially true for quantitative sampling, in determining the gear to be used

and the number of samples to be taken. Gray (1971b) and Vitiello (1968) have

approached this problem from the viewpoint of determining the most suitable

core size and the optimal frequency of sampling for meiobenthos. Jones (1961),

using a coring device, collected 30 samples at four stations at six-week intervals

to determine fluctuations in density and dispersion pattern of benthic animals,

and he compared the results of multiple-sampling versus one large sample.

Mclntyre (1971) has evaluated the effects of sampling techniques, especially

coring devices. The results of these investigators and the statistical books of

Cochran (1963), Greig-Smith (1964), and Williams (1964) should be consulted

prior to attempting quantitative sampling.

3.1. Preliminary Survey. —Before any quantitative sampling program is designed it is essential that a preliminary survey be made to define the faunal

components and to establish the dimensions of the ecosystem. In the deeper

sublittoral where remote gear has to be used, and exact replication of samples is

impossible, data can be derived only from previously published information on

the area.

3.2. Littoral Sampling.—Quantitative sampling can be accomplished by

sampling along a transect. A reference line is established perpendicular to the

low-water line along the surface of the sand from a fixed marker at the top of

the beach. The line should extend at least from mean spring low water to mean

spring high water. Samples from two or more stations along the surface are taken

at intervals of 0.5 to 2 m on nontidal beaches and no more than 3 m on tidal

beaches and at various depths from surface down to the level of the low tide

water table. For a given sampling effort a continuum of sampling designs can

be drawn up, ranging from nonsubsampled stations (Section 3.2.1) to multisubsampled grid stations (Section 3.2.2). The former possess broad extension

with little sample reliability, and the latter narrow extension with great sample
NUMBER 78 17

reliability. The two sampling designs given below include these features in

slightly different ways. In any case, sampling should be repeated at different

tidal levels.

3.2.1. Distribution Sampling.—In this case the broadest extension which is

consistent with estimated sample reliability is sought. Placement of the samples

can be made according to a stratified or random procedure, as suits the investigation. The samples are widely spaced along the transect. Such a sample scheme is

well-adapted for determining distribution limits of species for beach surveys and

areas where there is great faunal diversity.

3.2.2. Density Sampling.—In this case extension is less of a consideration

than sample reliability. Six to eight random 1/16 m 2 subsamples are taken from

a 1 /4 m 2 grid. This grid should be used at selected intervals along the transect,

e.g., low-, mid-, and high-tide levels, to obtain maximum contrast. Such a sample

scheme is well-adapted for determining density where a precise estimation of

numbers is more important than the distributional limits of species and where

there is little faunal diversity.

3.2.3. Corers.—For quantitative sampling, coring is the most suitable method.

Almost any type of tubing may be used, but transparent plastic tubing is preferable to metal since it allows visual recognition of core compression so that

precautions can be taken. In sand, samples should be taken from as deep as

possible into the sediment if the total fauna is required (see Section 3.2.4 for

methods of sampling the deeper layers of littoral sand), but on mud most of the

organisms are confined to the top few centimeters and cores of 8 to 10 cm in

length usually contain most of the fauna. On both substrates, however, an initial

examination of a long core will indicate the vertical distribution and suggest

how later sampling should be done. In all cases, if vertical zonation is to be

studied by sectioning cores, the sectioning must be done immediately upon collection, or as soon as possible afterward, to avoid errors due to subsequent redistribution of the fauna.

Ideally, the diameter of the corer should be such that it provides enough

animals so that processing can be done in a reasonable time without recourse

to subsampling, which can introduce errors. The most appropriate core size for

most meiofaunal studies seems to be within the diameter range of 3 to 4 cm.

Narrower cores may not sample the fauna effectively; wider corers may lead to

extraction difficulties. Core and sample size, however, must be related to the

size and relative abundance of the animals to be studied. In narrow cores there

may be the problem of core compression, and where this cannot be avoided the

investigator should note the length of cores that can be taken without this

occurring. In the sublittoral, other factors may be revelant. The presence of a

surface flock or a deposit which may be disturbed and inadequately sampled by

narrow cores may necessitate the use of a corer as wide as 10 cm from which

subsamples can be taken after removal of most of the supernatant water.

3.2.3.1. Simple Tubular Corers.—Simple tubes of plastic or metal are quite

suitable, but the problem of core extraction needs careful thought. Pushing out

the core with a plunger inserted into one end disturbs the surface and will

almost certainly cause core compression. In moist sediments the core will tend

to slide out of the tube under its own weight when it is removed from the substrate, and this tendency may be utilized to achieve an undisturbed sample. One

method is to insert a rubber bung into the top of the tube (when it has been

inserted to the required depth into the substrate) and then remove the corer
18 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

FIGURE 6.—Hope corer.

from the substrate. Gentle pressure on the side of the rubber bung allows air

into the top of the tube, and the core slides out. The admission of air can be

controlled by this simple method, and when a suitable length of core has emerged

it can be sliced off into a container.

A variation of this technique is the use of a corer devised by Hope. This

device (Figure 6) is made from a lucite tube approximately 40 cm long with

an outside diameter of 37 mm and a wall thickness of approximately 1 mm.

Lines are inscribed around what is to be the lower end of the tube to provide a

scale for measuring length or volume of core samples. The opposite or upper

end of the tube is fitted with a rubber bung. An air-intake hole 1 cm in diameter

is drilled through the wall approximately 15 cm below the upper end and then

covered with a rubber sleeve 3 cm wide. A section of bicycle inner tube serves

well for the sleeve.

Sediment samples are taken by pushing the lower end of the coring tube

into sediment, closing the upper end with the bung, and then withdrawing the

coring tube and sample from the substrate. The sample is then removed from

the coring tube by using the thumb of the hand in which the tube is being held

to roll down the rubber sleeve. As the air intake hole is uncovered, air enters

and the core slides out of the lower end of the tube. By using the scale inscribed

on the coring tube, the sample can be lowered from the coring tube by measured

amounts and each portion may be sliced off for vertical distribution studies. This
NUMBER 78 19

device has the disadvantage of having to be used in submerged sediment so that

the space in the corer above the sediment sample is filled with water.

In drier or very course sand or gravel the techniques given above work less

effectively and experimentation may be necessary to find a suitable solution.

3.2.3.2. "Ring-lined" Corers.—Jansson (1967a) has developed a corer which

is lined with a series of rings. The core and rings are pushed out together by

a plunger, and as each ring emerges it can be removed with a segment of core.

This method does not cause core compression as the plunger acts against the

rings and not the sediment.

3.2.3.3. Demountable Corers.—This type of corer can be split open and the

core cut into sections while still within the tube. The Renaud-Debyser (1957)

sampler is of this type and has additional advantages (Section 3.2.4). These

samplers depend on cohesion of the sediment and are difficult to use in drier sand.

FIGURE 7 —Thiel's "Meiostecher." (From Gerlach. 1968.)
20 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

3.2.3.4. "Meiostecher".—The "Meiostecher" (Thiel, 1966) is a modified coring device (see Figure 7 and Appendix, Section 10.1.13). The plexiglass box,

10 by 2.5 cm, is open at the lower end and has beveled cutting edges. It is pressed

into the sediment to the desired depth and is then closed by pushing a thin

copper blade or flexible plastic sheet through the groove (Gerlach, 1968). The

apparatus can be used in flat beaches and also for subsampling (see Section 3.3.2).

3.2.4. Sampling the Deep Layers of Beach Sand.—Although most of the

fauna of beach sands will occur in the top 25 to 30 cm, it is necessary to take

samples down to the level of the permanent water table, which may be as deep

as 2 m or more. Ax (1969) has described and illustrated the various techniques

of sampling above and below the water table.

A tubular corer about 30 cm in length can be inserted into the sand to its

maximum depth. The sand is then dug away to the level of the bottom of the

corer, and the corer is removed. Another corer is then pushed into the sand at

the point reached by the first, and this process is repeated until the water table

is reached.

A long demountable corer of the Renaud-Debyser type (Renaud-Debyser,

1957) can be used. This apparatus can obtain cores of up to 1 m in length.

A pit can be dug into the sand to the depth of the water table and a series

of cores taken horizontally, one below the other, into the wall of the pit (Zinn,

1942, 1968). The disadvantages of this method are the rigidity that it imposes

on the vertical subsampling of the sand column and the unknown effects of the

prolonged presampling disturbance of the sediment on the more motile elements

of the fauna. Hummon (1969) and Pollock (1969) have modified this method

by taking horizontal cores from the base of the pit as progressive depths are

reached. In this way, presampling disturbance is minimized and it is possible

to localize the fauna. Vertical subsampling is resticted, however, since a continuous

core is not obtained.

3.3. Sublittoral Sampling.

3.3.1. SCUBA.—The most satisfactory samples from the sublittoral are without doubt those collected by divers. Many of the littoral sampling techniques

are workable by SCUBA and the diver can select the exact spot from which to

take the sample. The size of the sample can be controlled, sampling irregularities

can be reported, and notes can be made on the extent to which the sample is

representative of the area as a whole. A diver can also sample areas of the shallow

sea inaccessible to remote gear, such as narrow submarine valleys, submarine

caves, small patches of substrate, and the transition zone between two adjacent

sediment types. For qualitative work large volumes of sediment can be collected

easily in bags or containers of any convenient size or shape. For quantitative

work, coring techniques similar to those used for littoral sampling can be used,

perhaps with some modifications. Sections 3.2.3 through 3.2.3.4 should be consulted.

While SCUBA techniques provide the best sublittoral samples, they have

certain obvious limitations, one being depth. The operational limit is 30 to 50 m,

and even towards the extreme of this range complex operations are difficult and

the time available to the diver on the bottom is limited by problems of providing

subsequent decompression facilities. The depth range can be extended only by

the use of complex underwater facilities. In collecting samples the diver must

take care, especially on muddy grounds, that his own activities do not disturb
NUMBER 78 21

the substrate and interfere with further sampling. Finally, it must be emphasized

that SCUBA operations can be dangerous; the necessary precautions and restricttions are adequately covered in the handbooks of national subaqua organizations.

It is imperative that SCUBA be used only by suitably trained personnel.

3.3.2. Remote Sampling.—For qualitative work, the best sampler is the one

which provides the maximum volume of sediment that can be handled by the

available facilities. For such work a dredge (defined as an instrument which is

towed and which samples horizontally) is probably better than a grab (an

instrument lowered on a vertical warp to take a single bite of sediment), but

where it is suspected that several sediment types may exist side by side the tow

should be of short duration. Several suitable instruments are available; sledges,

which are fitted with skids, skim off the surface layer and are particularly recommended. An example is the Ockelmann (1964) sledge. This instrument was

designed to collect only the larger species, and it requires modification to obtain

the smaller forms.

For quantitative work in shallow water the best remote gear is probably

an open-tube gravity corer. Hopkins (1964) has reviewed the various types of

gravity corers used up to 1964. The instrument devised by Moore and Neill

(1930), or some modification of it, is frequently used. On sand, however, a core

retainer is usually required, and the type described by Mills (1961) is suitable.

More recently corers have been described by Craib (1965) and Willemoes (1964),

the former taking cores of 15 cm in sand and having a built-in retainer. The

box corer of Reineck (1958, 1963) has proved very useful (see Appendix, Section

10.1.14). Drzycimski (1967) described a corer that uses the same principle as a

cutting knife for core retention; such a corer is much smaller and lighter than

Reineck's, and may be very useful in shallow water. A more detailed review of

this type of gear is given by Wells (1971). The same difficulties noted for divercollected cores occur with remote gear (loss of superficial deposit and core compression) , but they are more serious with remote gear since they pass undetected.

Because of this, all remote gear must be tested in shallow water with a diver

observing its mode of action, so that these problems can be eliminated or reduced

to the lowest level possible. Other instruments currently in use for sublittoral

meiobenthos sampling include the "mouse trap" sampler of Muus (1964), which

collects a superficial area of 150 cm

 2 to a depth of 2 to 3 cm, and the sampler of

Corey and Craib (1966), which takes a similar small sample of the surface layer

of the sediment.

Multiple-tube corers that provide a variable number of samples depending on

the instrument have been used successfully on sublittoral mud bottoms. The

Krogh-Sparck multiple corer (Krogh and Sparck, 1936) employs six tubes, each

28 mm in diameter and 50 cm long and having a rubber check valve at the top.

M. L. Jones (Smithsonian Institution, Washington, D. C, personal communication) has developed a "centuple-cell sampler" consisting of 100 square brass

tubes each measuring 2.54 cm (Figure 8). The device is so constructed that the

individual tubes can be disassembled. In the assembled state, the sampler covers

an area of 0.64 m2. Other multiple corers include those of Buzas (1968) and

Willemoes (1964).

Another frequently used method in sublittoral work is the collection of a

core sample from a grab haul. On sandy ground the gear should be as heavy

as is practicable to produce a good sample, but on mud a sampler that is too

heavy will sink into the deposit and disturb the surface layer. As with corers,
22 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

FIGURE 8.—Jones's "centuple-cell sampler.
NUMBER 78 23

the downwash of the instrument may disperse some of the superficial flock, and

if subsampling is to be done from a grab haul an additional problem is the

possible redistribution of the top of the sample during hauling and the possible

loss of all the surface flock.

Deep-sea sampling produces a number of additional problems (Hessler,

1971a). Because of the time involved in lowering and raising the gear and

because of the high proportion of partial or complete failures, careful planning

is required to make the best of each haul. Experience gained from work with

the macrofauna has shown that the standing crop tends to be greatly reduced

in the deep sea, so that quantitative sampling devices, with their comparatively

small volumes of sediment, have been less satisfactory for the systematist than

they have been for the ecologist. Consequently, deep-sea programs tend to include

both large-volume qualitative samples as well as quantitative methods wherever

possible. The generally much higher density of meiofauna may make this unnecessary for some taxa (Thiel, 1966), but until more information is available

it is probably still valid to take samples with as large a volume as possible.

There is no single instrument that can be recommended as a quantitative

tool. Several grabs including the Petersen, Van Veen, Smith-Mclntyre, Campbell,

and Ocean (see Hopkins, 1964 and Holme, 1964, for a review of grab samplers)

have been used with variable success in deep water. Box corers such as the United

States Naval Electronics Laboratory (USNEL) spade corer (Rosfelder and Marshall, 1967) and the Reineck (1958) "Kastengreifer" have only recently been

used at great depths, but they offer great promise because of their high success

ratio and their ability to take large, apparently undisturbed samples. Hessler

(personal communication) has recently succeeded in obtaining a series of undisturbed 0.25-m2

 biological bottom samples from a depth of 5,600 m using the

USNEL spade corer.

At shallower depths, both grabs and box corers may be subsampled to give

replicate quantitative samples of manageable size. Possibly the Hope corer (Section 3.2.3.1) and Thiel's (1966) "Meiostecher" (Section 3.2.3.4) can be used

for subsampling. Clusters of small diameter cores also may be useful. The replicates can be divided horizontally to give information on the vertical distribution

of the fauna.

All these instruments suffer from the disadvantage—already discussed—that

they produce a "bow wave" which will disperse the light superficial sediment and

its associated organisms. This effect can at least be reduced by making every

effort to ventilate the sampler by valves and mesh screens and thereby reduce

the downwash, and by attempting to lower the gear gently to the bottom. The

use of a gauze cover on the grab bucket, as in the Smith-Mclntyre grab, is an

example of such a ventilation method (Smith and Mclntyre, 1954; Wigley and

Mclntyre, 1964). Sou tar (personal communication) has modified this grab so

that the approach to and penetration of the bottom is very gentle, thus solving

the bow-wave problem. Gear should be hauled in at the maximum possible

speed so that winnowing of the sample is reduced, and this speed should be

maintained until the sampler is clear of the water in order to minimize the damage caused by the turbulent surface waters. The sampler should be hauled inboard

and the sample processed with a minimum of delay, thus avoiding final winnowing and damage to surface and near-surface dwellers caused by high-temperature

shock that may result in exposure to air having a much higher temperature

than the seawater.
24 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

One practical problem in deep-sea sampling is in knowing when the gear

has reached the bottom. Wherever possible the use of a pinger attached to the

wire a known distance above the sampler and monitored by a precision depth

recorder is recommended. A recording tensiometer also is useful. The greatest

problem, however, is winnowing during hauling. Few samplers are able to close

completely, and this may not represent a serious problem at shallow stations;

but in the deep-sea, where the gear is being hauled for a considerable time and

through a long column of water, even slight leakage can result in the loss of

most of the sample.

These quantitative devices cover a surface area 0.1 to 0.5 in

 2

, depending on

the type. As already suggested, samples of this size may be too small for the

investigator's requirements in the deep sea and, in particular, they will not

satisfy the systematist. In this case, qualitative samples must be utilized. A suitable qualitative sampler should collect only the top few centimeters of the deposit

over a fairly extensive area and should not be subject to winnowing. Suitable

devices of this type have not yet been designed for deep-sea meiofauna work,

but some of the meiofauna samplers presently in use could be modified appropriately, such as the epibenthic sledges designed at the Woods Hole Oceanographic Institute (Hessler and Sanders, 1967). As used for larger organisms the

epibenthic sledge is fitted with a 0.5-mm mesh bag, but for meiofauna sampling

the mesh size could be reduced. Also, for depths greater than 2,000 m the mouth

should be fitted with a postsampling closing device. In sampling with the sledge

a weight should be attached to the wires to reduce the amount of warp required,

and for quantitative sampling a pinger and tensiometer are extremely valuable.

The latter is particularly useful in that it gives a warning when the gear encounters an obstruction. In trawling, speeds of 1 knot or less are necessary.

Finally, it should be noted that while the use of deep-sea submersibles is

not yet widespread, this facility will permit the extension of well-tested shallowwater techniques to great depths.

4. General Extraction Techniques

This section deals with techniques for the whole or at least the major part of

the fauna in a sample. Reference is made to techniques used by the ecologist

for strictly quantitative work as well as those which treat large volumes of

sediment. The latter would produce specimens which, in some cases at least,

would satisfy the minimum requirements of the systematists.

For living fauna there are two main divisions of the techniques: those which

depend on the natural movements of the animals without the use of anaesthetics,

and these techniques are favored by those requiring material for physiological

experiments; and those which involve removal of the animals from the deposit

by physical disturbance such as sieving or elutriation, when an anaesthetic is

required. For preserved material, staining of the total sample (for example, by

Rose Bengal) usually is required.

4.1. Field Techniques.—These are usually less than quantitative, but are

designed to provide some degree of extraction. Often it is impossible to extract

the meiofauna immediately after collection. Though often it is stated that it is desirable to store the samples at low temperature (around 5° C), J. B. J. Wells

(personal communication) has had good results in the tropics by storing samples
NUMBER 78 25

at or near ambient room or seawater temperature. He found that samples stored

at 10° to 15° C deteriorated within 12 hours with all taxa dead, whereas samples

stored at ambient temperature in tightly sealed containers for 36 to 48 hours,

and in some cases up to four days, were in good condition (the samples did not

fill the containers and there was air above the sample). For those animals having

short generation time, such as Ciliata, this procedure would not be satisfactory

for quantitative studies. Nor would this procedure be satisfactory for highly

predaceous organisms such as certain Nematoda.

4.1.1. Sand.—The following method is recommended. Seawater is filtered

through a fine mesh (62 p) net into a clean bucket, and a relaxant (chloroform,

alcohol, formalin, MgC^, propylene phenoxotol) may or may not be added. The

sand sample is then added to the bucket and left for 10 to 20 minutes to let the

relaxant act. It is then stirred, and after momentary settling the water is decanted

through a 62-/x net; stirring and decanting are repeated until the lighter sediments

are almost entirely removed. The meiofauna is finally washed down into the

cone of the net and the everted net washed into a container. The sample is then

divided into two nearly equal fractions. One of these is preserved in formalin

and the other in alcohol, by graded increments. For formalin preservation, add

4 percent formalin, a few drops at a time, over a period of 40 minutes. For

alcohol preservation add 70 percent alcohol in the same way. At the end of the

process, decant the liquid and replace with fresh 4 percent formalin or 70 percent

alcohol. The well-known method of Chappuis (1942) also is relevant to this

section.

4.1.2. Mud.—The bubbling technique of Higgins (1964) is described in

Figure 9. This technique is suitable, but it may be detrimental to certain taxa

such as Gastrotricha, Gnathostomulida, and Turbellaria, and generally it is ineffective for removing Nematoda.

4.1.3. Shipboard Techniques.—If operating from a ship which is at sea for

only one or two days, it may be possible to maintain the samples in sufficiently

good condition for laboratory extraction by storing them in a refrigerator. But on

cruises lasting more than a few days it usually is necessary to accept the less-thanideal extraction method, which can be done at sea, or to preserve the unprocessed

sample.

If extraction techniques are to be employed on the open sea, it should be remembered that such techniques must be simple, since the ship's motion and

vibration as well as the shortage of space and time on most cruises make complicated techniques impractical. It may be possible to use the Boisseau technique

(Section 4.2.1.4) in good weather, but in general, for small samples of living

material, shaking with MgCl-j and decanting is the most practical procedure.

At the qualitative or semiquantitative level the techniques described for sand

(Section 4.1.1) and mud (Section 4.1.2) can be used on board for collections of

surface deposit obtained with gear such as the epibenthic sledge.

For larger volumes of material from deep-sea samples the elutriation technique used for macrofauna (Sanders, Hessler, and Hampson, 1965) can be applied

to meiofauna work by substituting appropriately smaller sieves. This technique

involves pumping large volumes of seawater at low velocities through the sediment and allowing the supernatant liquid to flow through selected sieves. The

flow of water should be sufficient to prevent even the light but coarse particles,

such as Globigerina tests, from flowing over onto the screens, since these make

later sorting unnecessarily difficult. It is important, however, to accept the fact that
26 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

10

(8-9-10)

FIGURE 9.—The bubbling technique of Higgins (1964)

for extraction of mciofauna. Seawater is filtered

through a net into a clean bucket (1), and if desired

an anaesthetic (2) as described in Section 4.1.1 may be

used. The upper 1 to 2 cm of surface mud from a

minimum area of 1 in- it removed, added to the

bucket (3a,b)—making one part mud to three parts

water—and stirred (U,b). Air is then bubbled through

the mixture (5a,b). and after momentary settling the

water surface is blotted with paper (6b), which is

immediately removed, and the washings are collected

by the net. This process is repeated three to seven

times, after which the fauna is washed down into the

cone of the net (8), which is then inverted and the

collection is washed into a container (9) and preserved

(10). For sand collections, two samples—one preserved in formalin and one in alcohol—should be taken. Finally, the sediment

in the bucket is washed through a 62-p sieve (6a) and accumulated in a separate bucket (7<i). This sediment then is divided

into two parts, which are preserved in alcohol and formalin (11, 12) as described in Section 4.1.1.

the stenotopic deep-sea animals are unlikely to survive manipulation on deck.

Death is almost certain as a result of thermal shock, which accompanies the transfer of the fauna from the cold deep to the warmer surface conditions. The animals

may still be alive on reaching the surface because of the insulating effect of the

large volume of sediment, and if adequate refrigeration facilities are available it

may be possible to keep them alive at least long enough to apply anaesthetics before preservation. If extensive manipulation of the sample is required in the living

state it is best to bring large refrigerated samples ashore for processing, and this

may be practical where a laboratory is close to the deep-sea areas, such as at

Bermuda.

4.2. Laboratory Techniques.

4.2.1. Sand.—A combination of techniques must be used to extract small

motile organisms (e.g., Ciliata) and larger thigmotactic forms (e.g., Nematoda).

The motile forms can be extracted with the Uhlig Seawater Ice Technique (Uhlig,

1964, 1966).

4.2.1.1. Uhlig Seawater Ice Technique.—Seawater from the vicinity of the

sediment sample is frozen. The sediment sample is placed on a nylon gauze

(140-/X pore diameter) at the bottom of a tube. A thin layer of cotton wool is
NUMBER 78 27

placed on top of the sediment, followed by the crushed seawater ice. As the ice

melts, small motile organisms are carried through the gauze and can be collected

in a Petri dish. The technique also has been successfully used with 40° C water

instead of the ice, and this might be preferable in tropical regions.

J. B. J. Wells (personal communication) has found that the Uhlig technique

works as well in the tropics as in cold climates, based on his studies in Ernakulam,

India. Melting time of the ice is, of course, shorter, but he found that this caused

no problems in extraction. He found that cold water (temperatures of 4° to 6° C)

is quite effective, but that it is essential to place a fine-pore filter over the sand to

slow down the filtrate. Very small Ciliata were extracted less well with cold water,

however, than with ice. Wells also found that water at 50° to 60° C extracted more

animals than water at 40° C. The extraction efficiency for Ciliata was, however,

much less, and many animals were killed and washed out passively. He did find

that water of higher temperature had a relaxing effect on many animals.

Fenchel (1967, 1969), Thane-Fenchel (1968), and Uhlig (1964, 1966) should

be consulted for extraction efficiency of the Uhlig technique. Extraction efficiencies of 75 percent have been reported for Ciliata, Flagellata, Turbellaria, Gastrotricha, and Archiannelida. Lower efficiencies are reported for Nematoda,

Rotifera, Harpacticoida, and Tardigrada. Prolonged extraction and the use of

larger extraction tubes appear to increase the extraction efficiency of some taxa.

4.2.1.2. Anaesthetization Technique.—After treatment to remove the motile

organisms the sediment is placed in a beaker, and an equal volume of 6 percent

MgCl2 (i.e., 73.2 g/liter, which is isotonic to 34 ppt seawater) is added. After

allowing 10 minutes for anaesthetization the sediment is stirred thoroughly and

the supernatant poured through a fine mesh sieve or plankton net (62-/i pore

diameter). This is followed by two washes of seawater and a wash of 4 percent

formalin or 10 percent ethyl alcohol. The sieve is inverted in a Petri dish and the

animals are washed off with a jet of filtered seawater.

The sample must be examined periodically after the washing and the extraction efficiency noted. This is necessary when a new locality is sampled. The accuracy of the method must be quoted in any publication.

4.2.1.3. Deterioration Technique.—Although not quantitative, this technique

concentrates the vagile fauna. Large volumes of sediment can be used, but when

mud is used the layer should not be too deep. The sediment is placed in a bucket

and allowed to deteriorate at a temperature slightly above that in the field. The

animals migrate to the sediment surface, provided that deterioration does not

proceed too quickly. Surface layers of sediment are removed and animals are

extracted following the techniques described in Sections 4.2.1.1 or 4.2.1.2. A wide

variety of substrata, including sponges and algae, can be used with this technique.

4.2.1.4. Boisseau Technique.—The apparatus for the Boisseau technique

(Boisseau, 1957) is shown in Figure 10. Sediment is placed in the separation funnel (A) and an equal volume of 6 percent MgCl2 is added. After allowing 10

minutes for anaesthetization, a continuous stream of seawater is introduced

through the tap on the separation funnel. The pressure is adjusted so that a balance is achieved between vigorous agitation of the sand and only a small amount

of sediment is carried over to tube C and the sieve (D). After 15 minutes of elutriation the tap on tube C is opened and the water allowed to drain through the

sieve. The sieve is inverted in a Petri dish and the animals are washed off with a

jet of filtered seawater.
28 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

B

62/J MESH

FIGURE 10.—Boisseau apparatus

for extraction of meiofauna.

(Modified from Boisseau, 1957.)

This technique is not recommended for coarse sands, as the efficiency is reduced. Further, while it can be used as above for live fauna, several groups

(Tardigrada, some Nematoda) are difficult to detach from the sand grains, and

others, such as Turbellaria, quickly recover from the anaesthetic and attach firmly

to the walls of the tube. This difficulty can be overcome by setting up the equipment as a closed system and maintaining an adequate concentration of MgCl2 in

the water pumped through the apparatus. The Boisseau technique, however, is

highly satisfactory with preserved material, at least for the "hard" taxa, and the

apparatus has been modified by Lasserre to operate with one or two liters of sand.

The principal parts of the Lasserre modification are one 5-liter glass flask with

two stirring systems. The sand is agitated by a flow of seawater entering the tubes,

which are divided into four elements, and by a mechanical stirrer used once or

twice during the process. A glass column contains a large mesh net which retains

most of the very coarse particles and fragments of sea grass and algae. The column

is easily drained via the tap on tube C (Figure 10). The animals and fine organic

particles are retained on a tightly fitting nylon gauze, the mesh size of which is
NUMBER 78 29

62 M. From 94 to 95 percent of the fauna contained in one to two liters of sand

can be extracted in 15 minutes of washing.

4.2.2. Mud.

4.2.2.1. Quantitative Extraction of Total Fauna.—In deposits without a good

pore structure the Uhlig technique cannot be used, and when even moderate

quantities of fine material are present the Boisseau or decantation procedures are

not applicable, so that sieving and hand extraction are required.

The usual procedure for living meiobenthos is to divide the sample into two

fractions, using a fine sieve (30 to 60 /i). The coarser fraction is sorted, using a

binocular microscope. The fine fraction, which contains the silt and clay, is stirred

into uniform suspension and the liquid is subsampled. The deposit in the subsamples is allowed to settle out in a thin layer on a Petri dish, and the animals can

be seen and counted by their movements. A similar technique can be used for

preserved material, but the sample is first stained with Rose Bengal (0.1 gm/

SILICONE COATED

LARGE PIPETTE

NYLON MESH

I50J) OR LESS

PLEXIGLASS

NYLON SIEVE

PETRI DISH

FIGURE 11.—Swedmark technique for extraction of meiofauna.
30 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

100-200 ml of 5 percent formalin or 70 percent alcohol), and because of the difficulties of hand-sorting dead specimens from silt and clay material, the finest

fraction is usually discarded. Although laborious, the method used for live material is completely quantitative. With preserved material, it is quantitative within

the limits of sieve used, and it can be made fully quantitative if an occasional live

sample is available to give estimates of what proportion of the fauna passes the

sieve.

4.2.2.2. Swedmark Technique.—This technique (Figure 11) can be used for

fine sand as well as muds. The deposit, stirred to break up lumps, is placed in a

large vessel and covered with 1 or 2 cm of seawater. The mud surface is then

pumped into suspension, using a large silicone-coated pipette (for preparation of

the pipette see Appendix, Sections 10.4 and 10.1.15), and transferred to a nylon

sieve or series of sieves (250 /x, 62 p), the largest having a diameter slightly less than

the normal size of the Petri dish used. Sieving is done by gently rocking the sieve

in seawater either in another vessel or in the original one so that the filtrate is

returned to the original sample. When sieving is complete the sieve is placed in

seawater in a Petri dish so that the fauna can be first examined under a binocular

microscope without the damage that later may be inflicted by transferring from

the sieves. This procedure can continue for as long as the animals remain alive—

for several days if desirable. At this stage, the process does approach a quantitative

method as long as the exact size of the original sample is known.

4.2.2.3. Bubbling Technique.—This technique is discussed in Section 4.1.2,

for field studies. It can be suitably modified for laboratory use.

4.3. Counting and Observation Efficiency.—Although the extraction efficiency may reach 80 to 90 percent, observational efficiency should be determined

periodically by reexamination of previously counted samples. Otherwise this may

be a source of error that is frequently ignored. Provided that extraction and observation efficiencies have been estimated, quantitative comparisons of fauna can

be made. Frequently, however, raw data cannot be used for comparative purposes.

Transformation procedures are necessary for the statistical validity of the data

(see Greig-Smith, 1964; Williams, 1964).

5. General Taxonomic Procedures

This section covers procedures for the examination of live material, for fixation,

preservation, and curation methods, and for data presentation. There are several

publications that provide information on preservation and other techniques for

marine animals: British Museum (Natural History), 1954; Guyer, 1953; Lo

Bianco, 1899; Russell, 1963; Wagstaff and Fidler, 1955. Most of these publications,

however, deal with larger marine invertebrates and they provide little or no information for meiofauna.

5.1. Examination of Live Material.—Examination of live material is highly

desirable in all meiofauna groups, and it is indispensable in the "soft" taxa. A

wide range of biological data, from locomotion to feeding, sexual behavior, and

even life cycles often can be obtained by direct observation, and such data should

be incorporated into systematic descriptions whenever possible. In many "soft"

taxa, furthermore, determination, as well as description, based on preserved material alone is impossible. The lack of live data can only partly be compensated by

time-consuming histological techniques.
NUMBER 78 31

5.1.1. Techniques.—Transmitted light usually is preferable, although incident light should be used for color observations. Single "soft" specimens can be

placed on a slide with a pipette, the end diameter of which should exceed only

slightly the diameter of the specimen. The inside of the pipette should be coated

with silicone to reduce adhesion of animals. Specimens with a cuticle can be

placed on a slide with an Irwin loop (see Appendix, Section 10.1.16). Specimens

should be observed in seawater of the same habitat, using a slide and coverslip.

For "soft" taxa, the coverslip must be supported to prevent damage by weight and

capillary pressure. The edges of the coverslip can be scraped over bee's wax or

plastiline, thus providing the necessary support. Most "hard" taxa do not require

support, although the same procedure may be employed. Observations on behavior, color, body outline, and external structures should be carried out immediately.

Anaesthetics can be added from one side of the coverslip following initial

observations. Observations on body dimensions and position of internal organs

can readily be made. The specimen can be rotated along the longitudinal axis by

applying pressure to the coverslip. Drawings at various magnifications can be

made under these conditions.

Squeezing of the animal can be accomplished by gradually removing all of the

liquid and applying pressure to the coverslip. The animal becomes more transparent, permitting a detailed analysis of the internal organization. Should it be

desirable to keep the specimen for histological purposes, squeezing should be kept

to a minimum. If histological studies are not required, further squeezing often

will reveal more delicate details, such as cuticular structures, gametes, etc.

Although the squeezing method does not present major problems, it can be

much simplified by using a "Roto-Compressor," described by Heunert and Uhlig

(1966). Use of such an instrument is especially advisable where the animal should

be kept in a constant situation over a longer period of time (e.g., for time-lapse

filming).

Excellent optical equipment is mandatory for critical observation. Phase contrast is absolutely indispensable for most meiofauna research, and the new interference contrast (Nachet, Reichert, Zeiss) already is adding a new dimension to

the analysis of small organisms. Whereas the use of electron and scanning microscopes is not necessarily required for taxonomical investigation, good microphotographic equipment and scale drawing apparatus (Wild, Zeiss, Leitz) should be

part of the standard optical equipment. Where a drawing apparatus is lacking,

at least photographs should be made to complement the drawings. If both are

lacking, drawings should be made with the help of a measuring eyepiece, and

whole mounts of specimens should be kept for a check of meristic data until better

equipment is available.

In order to obtain maximum information from every specimen examined, a

"count-down" of characters should be set up for every taxon. This means a complete listing of all characters to be observed, arranged according to their optimal

visibility and/or logical progression during the different stages of examination.

5.2. Fixation.—Meiofauna may be fixed while still in the sediment, as a

mixed aggregate of organisms after extraction or as individual taxa or individual

specimens after extraction and sorting. Fixation of unsorted meiofauna in sediment is recommended only as a last resort, since there is no single method for

optimally fixing both "hard" and "soft" fauna, nor of simultaneously fixing all

forms of "hard" fauna. The preferred method is to separate the "hard" fauna;
32 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

single specimen fixation is preferable since it allows appropriate techniques for

both whole mounts and further histological (light and electron microscopy)

studies. It is emphasized that in the case of "soft" fauna any type of fixation without live examination is acceptable only under special conditions.

Unfortunately, it is often impractical or impossible to extract and sort meiofauna before fixation because of the time involved and lack of help or facilities.

Under these circumstances the most satisfactory compromise is to anaesthetize

(Section 4.2.1.2) the organisms, whether extracted or not, and then add sufficient

full-strength buffered formalin to the sample to bring the final concentration to

approximately 5 percent for at least 24 hours.

5.2.1. Fixation of "Soft" Fauna.—For unsorted "soft" fauna samples, the use

of warm Bouin's fixative for two to four hours following anaesthetization is recommended. Individual fixation can be for whole mounts or for histological purposes. Whole mounts are preferable where hard structures are important for

determination and for reference collections and type specimens.

5.2.1.1. Whole Mount Fixation of "Soft" Fauna.—Often it is necessary to

fix representatives of "soft" fauna while they are compressed, especially when

particular characters appear under this circumstance. A drop of 10 percent formalin is added on one side of the coverslip, and the original medium is removed with

filter paper on the other side. A mixture of glycerol and formalin (3:1) is a satisfactory combined medium for fixation and preservation.

5.2.1.2. Fixation of "Soft" Fauna for Histology.—For general histological

purpose, Bouin's fixative, preferably at 60° C, is recommended. Following anaesthetization the animals are pipetted from the seawater and then rapidly placed

into the warm fixative. The animals should be left in the Bouin's solution for at

least two hours, but preservation in the fluid for days or weeks often has no effect

on the quality of the preparations.

For electron microscopy, primary fixation in glutaraldehyde in seawater seems

to be widely used. Methods using this technique, however, are still in a stage of

experimentation.

5.2.2. Fixation of "Hard" Fauna.—There appears to be no single fixative

that is optimal for all taxa of "hard" fauna. Formalin makes the internal structure of mites resistant to pepsin digestion (see Section 6.21), and unbuffered

formalin erodes calcareous structures which often are important taxonomic features. Alcohol fixation, on the other hand, causes Nematoda to collapse, and

internal organs, which are of taxonomic importance, may be badly distorted. The

best fixation for most kinds of "hard" fauna, however, is 5 percent buffered formalin in seawater for at least 24 hours.

5.2.2.1. Fixation of "Hard" Fauna for Histology.—Usually it is not necessary

to make histological preparations for taxonomic studies of most forms of "hard"

fauna since external body features are used extensively for this purpose. However,

in instances where anatomical studies necessitate such preparation, especially in

the case of Nematoda, it is desirable to avoid alcohol fixatives which extract water

faster than alcohol can pass inward through the body wall, thus causing the organism to collapse. In addition, tissues may tend to shrink, causing the tissue

components of the body wall to pull away from the extracellular component, or

cuticle. The need to preserve water-soluble inclusions of cells or nuclear structures

may override the disadvantages of alcohol fixation in certain studies.

Formalin (5 percent) fixation is recommended for histological study of Nematoda, and Bouin's fixative for most other hard fauna.
NUMBER 78 33

5.3. Preservation.—The preservative selected must be compatible with the

fixatives used, as well as with the nature of the preparations that must be employed

in subsequent studies of the specimen.

5.3.1. Preservation of "Soft" Fauna.—As in the case of fixation, it is preferred

that as much sorting as possible be accomplished before the specimens are preserved. If sorting cannot precede preservation, the specimens should be preserved

in 70 percent alcohol, and it is further recommended that this concentration of

alcohol be reached gradually, i.e., over a period of one hour or two hours by using

graded concentrations of alcohol.

5.3.2. Mounting of "Soft" Fauna.—Two types of whole-mount media can be

distinguished: those which mix with water and fixatives and, therefore, can be

applied directly to the fixed specimen; and those where preservation has to be

preceded by dehydration techniques.

It is important that whole mounts be permanent and made with great care,

especially where type material is involved.

5.3.2.1. Water Mounting.—Glycerol is generally accepted as a good medium.

It is drawn under the coverslip where a squeezed and fixed animal can be maintained in a particular position, or added to the animal on the slide before the

coverslip is put in place. The coverslip is then ringed carefully, avoiding air bubbles, with a commercial varnish such as Eukitt or Murrayite (see Appendix,

Sections 10.1.17, 10.1.18). Other aqueous media are more applicable to different

taxa and are dealt with accordingly.

5.3.2.2. Mounting after Dehydration.—Fixed animals are washed in 70 percent ethyl alcohol for 10 minutes; then placed in 95 percent ethyl alcohol (three

changes, 5 to 15 minutes each); and then placed in absolute ethyl alcohol or butyl

alcohol (three changes, 15 minutes each). After three subsequent 5-minute washings in xylol the animals can be mounted in Canada balsam.

5.3.2.3. Histological Preservation of "Soft" Fauna.—After fixation in Bouin's

solution, specimens are rinsed in increasing concentrations of ethyl alcohol. The

optimum is 10 minutes with 40 percent ethyl alcohol followed by several hours

in 70 percent ethyl alcohol. The specimens then are transferred into 80 percent

ethyl alcohol.

5.3.3. Preservation of "Hard" Fauna.—Formalin often is recommended as

both a fixative and preservative, but when used as a preservative it should be buffered and the pH checked periodically. Preferred preservation of Nematoda and

Harpacticoida is in buffered formalin or anhydrous glycerin. Ostracoda and other

fauna that may be degraded in inadequately buffered formalin should be preserved in alcohol.

5.4. Curation.—Frequently, large collections of specimens must be held in

bulk storage before sorting and indentification. Since this may involve a considerable lapse of time, it is imperative that specimens be stored in suitable containers

with durable labels bearing collection data. Records with detailed collection notes

and ecological data, when available, also should be retained in permanent files.

If only selected taxa are sorted from general collections and studied, measures

should be taken to bring the remaining organisms to the attention of other qualified systematists, especially when the collections originate from remote localities

and/or are accompanied by considerable ecological data.

Finally, it is most important that after specimens are set aside as type material

they should be deposited in museums or other institutions where facilities and
34 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

personnel exist to assure their proper preservation. Catalog numbers or identifying numbers must be included in descriptions, thus making the specimens readily

available to other qualified systematists.

5.5. Presentation of Data.—Recently, Mayr (1969) comprehensively summarized our present conception of taxonomic descriptions. Since all of his statements apply to meiofauna, the following subsection deals only with problems that

are more relevant to or are most often neglected in studies of meiofauna. Most of

the following statements refer to original species descriptions, but they also are

true for complementary descriptions and identification. In the past, descriptions

often have been kept to that minimum necessary to distinguish them from species

described previously. A modern description, however, should be aimed to the

future and include all characters available.

5.5.1. Minimum Requirements.—Whereas it is impossible to recommend a

minimum number of specimens on which a description should be based, the series

should be as abundant as possible or manageable. Complete and undamaged

adult specimens of both sexes are required; descriptions based on juveniles can be

accepted only under exceptional circumstances. Observation of the living animals

is absolutely indispensable in "soft" fauna except in the few cases where valuable

material is available only in preserved state; optimally preserved specimens are

required in the "hard" fauna. Descriptions without illustrations must be rejected,

and it is recommended that illustrations be made with a drawing apparatus, or

checked with photographs, or otherwise carefully constructed. Collection data

must be given as completely as possible; and types must be deposited in competent

institutions. If the species is not known, it can be designated as "sp." or by placing

"cf." (confer, compare with) before the species name wherever there is the slightest doubt about species identity or in case the determination was made by a person

who does not consider himself fully competent in the particular taxon. The danger

of using "cf." is that this designation is often omitted in subsequent publications,

thus leading to erroneous reporting of the species. It is important to note any

differences from the original description. The inclusion of all observable morphological characters will provide more useful comparative data.

5.5.2. Descriptions.—It is imperative to distinguish between documentation

(presentation of data) and speculation (discussion of systematic relationships,

phylogeny, etc.). A good description should yield the following data, preferably

in a consistently uniform arrangement.

(a) A detailed statement should be given on number of specimens, develop

mental stage (juvenile, adult), and state (live, whole mount, or sectioned) of

the material examined.

(b) Data on locality, depth, substratum, associated fauna, and as many ecological parameters as possible should be included.

(c) Names of new genera and new species should be etymologically explained.

(d) Numbers, state of preservation, and location of type material should

be given.

(e) The description (sensu stricto) should follow the same order within a

given taxon (e.g., external features anterior to posterior, then internal organs).

It also is desirable to number the specimens available during examination and

to do the same in the description ("one specimen was exceptionally long" would

then read "Specimen 17 has the exceptional length of 870 p," and a caption such

as "anterior end" would be better as "anterior end of specimen 10"). The concept
NUMBER 78 35

of the specimen as the lowest taxonomical entity facilitates revisions (as the

material for a description later on sometimes turns out to comprise several species)

and makes the material more available for statistical studies.

For relative body dimensions and position of organs, it is advisable to take

the body length as 100 Units (U 0 to U 100), e.g., "Ovary from U 60-U 72"; or

"maximal width 33 jn = 7 U at U 65"), or as percents.

(f) Drawings should be given of all taxonomically important features and

of features that are remotely suspected to be important. Photographs are complementary in view of the original optical impression and should not be used without proper line drawings. All illustrations should be accompanied by scales.

(g) Meristic data, from as many specimens as possible, are ideally presented

in the form of tables.

(h) As many biological observations, including behavior, feeding, reproduction, etc., as are available should be included.

(i) Every new description must be accompanied by a diagnosis. Where new

facts have to be incorporated in an earlier diagnosis, the modified diagnosis has to

be given. For new genera, the type-species must be designated.

(j) A discussion should contain the reason (s) for placing the species in a

particular genus, further considerations of the taxonomic value of characters and

of systematic relationships and, if necessary, explanations for biological or ecological features observed.

6. Comments on Selected Taxa

This section deals with problems concerning individual taxa as compiled by the

specialists. It is concerned with optimal sample size, extraction, examination,

fixation, and preservation procedures.

6.1. Foraminifera.—Foraminifera are abundant in most types of sediment

and at all depths. Any dredge, corer, or grab sampler that will retain fine sediment is adequate for qualitative sampling. For more accurate relative and absolute abundance data, specific volumes, which are preferable (Walton, 1955), or

weights often suffice. For very careful numerical analyses, short cores (or tops of

long cores) should be used, and mixing in cores should be avoided. Core diameter should be 3.5 cm (Phleger, 1960); Foraminifera tend to be dense enough that

the top centimeter of this diameter usually yields hundreds of specimens. If not,

the top 1 to 4 cm can be analyzed, centimeter by centimeter. The number of

samples depends on the particular problem. Quantitative studies require several

samples or replicates (Walton, 1955; Brooks, 1967; Buzas, 1969).

Five percent buffered formalin is a suitable preservative for whole sediment

samples, but the samples should be washed over a 62-/A sieve and stored in 70 percent alcohol as soon as possible. Organic material may become acidic, reducing the

buffered formalin which destroys calcareous tests and cements. With samples kept

in formalin for any time, the pH should be checked frequently and more buffer

(hexamine or Na2CO3) added as necessary.

To determine living specimens, the sample can be stained with Rose Bengal,

a protoplasmic stain, following a procedure modified below from Walton (1952).

The Rose Bengal solution is prepared by the addition of 0.1 g/100-200 ml of

5 percent formalin or 70 percent alcohol. It is important to avoid a precipitate

caused by using an excess of powdered Rose Bengal. The stain can be added at
36 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

collecting time, or, better, a few hours before study. The sample should be mixed

well to insure stain distribution; and this should be done a few hours before study

because, with samples fixed in formalin, the pH cannot be checked with pH paper,

and, less conveniently, a pH meter must be used. In addition the stain fades with

time, although fading can be retarded by storing samples in the dark or in amber

glass jars. Stained preserved specimens should be sorted from water-covered

samples.

If extraction of specimens from preserved samples is necessary, samples should

be washed with water on a 62-/i sieve. This is followed by drying and separation,

using heavy liquids. Liquids used include carbon tetrachloride, bromoform, tetrabaromethane, tetracholorethane, Clerici solution, and others (Gibson and Walker, 1967). Bromoform seems preferable. Some undesirable organic debris can be

dissolved with NaOCl or H2O2, and, even better, by subsequent ignition in a muffle furnace (Smith, 1967). These techniques should not be used if one wishes to

recognize live specimens and if arenaceous specimens are present or are of interest.

After separation or concentration, the foraminiferal concentrates are examined microscopically. If numbers of specimens are very high, a suitable aliquot of

about 300 specimens can be obtained by using a microsplitter, with a dry sample.

Usually, specimens are picked out of examination dishes or picking trays (preferably black with a grid) with a wet 00 or 000 red sable brush and placed in 18-to-48-

ply assemblage slides. Slides should be precoated lightly with water-soluble gum

tragacanth so that specimens will adhere.

If live specimens are desired, they may or may not be extracted upon collection. Arn61d (1954) gives a method for extracting very shallow-water specimens

associated with plants. Extraction is generally by microscopic picking of unwashed

sediment or sediment water-washed over a 62-/x sieve. Otherwise, the entire sample

may be placed in aquaria or other suitable receptacles. Natural seawater or

distilled water, to which a compound such as R1LA marine mix is added to the

desired salinity, may be used and aerated. Arnold (1964) and Lee, Pierce,

Tentchoff, and McLaughlin (1961) should be consulted for culturing and study

of living and preserved specimens and for histology of Foraminifera.

There are several keys and studies of systematics, including Cushman (1948),

Galloway (1933), Glaessner (1945), Le Calvez (1953), Orlov (1959), and Loeblich and Tappan (1964). Cushman has been the most widely used in North

America. Of the above references, those previous to Loeblich and Tappan

(1964) reflect the earlier concepts of the systematics of Foraminifera, and considerable latitude is allowed the individual in taxonomic placement.

6.2. Ciliata.—Ciliata are found in almost all sediments. Their abundance,

however, shows great fluctuations. Because of the small size and usually great

abundance of Ciliata, small-diameter core samplers can be used. The Reineck

(1958) box corer has been used successfully in the sublittoral (Section 3.3.2).

Most species are fragile and are easily damaged. Indeed, some will be destroyed immediately upon coming into contact with the water surface. Nearly all

Ciliata must be collected fresh, studied alive, and specially prepared for identification. Only a few can be identified under low magnification. Bright field phase

contrast microscopy usually is necessary for identification.

Qualitatively, Ciliata can be extracted from the sediment by the deterioration

technique (Section 4.2.1.3). For quantitative studies the Uhlig Seawater Ice Technique is considered the most efficient (Section 4.2.1.1) since over 90 percent extraction can be achieved in 30 minutes. Isolated Ciliata are best preserved after silver
NUMBER 78 37

impregnation. As in many "soft" taxa, there is no suitable technique available

for preserving Ciliata within the sediment. Fenchel (1967, 1968a,b, 1969) should

be consulted for recent methodology.

No general keys are available, but the extensive reviews of Dragesco (1960)

and Kahl (1930-1935) should be consulted.

6.3. Cnidaria.—The classes Hydrozoa, Scyphozoa, and Anthozoa are all rep

resented in the meiofauna. In all, 15 genera are known, and most of these comprise only one or two species; however, some species are described from only a

polyp or medusa and this constitutes an important taxonomic problem.

Most species may be restricted to localities of only a few meters in extent.

Usually the animals are rare, although in some areas they may be locally common.

Most Cnidaria are robust and taxonomic characters are not likely to be easily

damaged.

Between 5 and 40 liters of sediment, obtained with a grab or dredge, is considered a suitable sample for Cnidaria. Quantitative samples can be obtained by

using corers or subsampling with cores from a grab. Live animals can be obtained

by using the Uhlig Seawater Ice Technique (Section 4.2.1.1) or by MgCl2 anaesthetization (Section 4.2.1.2). For mud samples, the Swedmark technique (Section

4.2.2.2) is successful. Whole sediment samples can be preserved in 5 percent buffered formalin and animals extracted later by the decantation technique (Section

4.1.1).

Live animals are best studied after anaesthetization with MgCl^ under phase

contrast optics. Neutral red should be added to aid identification of the nematocysts. Astomocnides retain their color after discharge, whereas stomocnides lose

theirs. The addition of acetic acid accelerates discharge.

For histological purposes 5 percent formalin or Bouin's fixative is recommended. Whole mounts can be stained with carmine and mounted in Canada

balsam. Buffered gluteraldehyde and epoxy resin embedding is recommended for

thin sections.

Alcohol preserves larger forms best, whereas small forms are best preserved as

whole mounts. For publication purposes nematocysts should be drawn in both

discharged and undischarged states with measurements given for the different

parts.

No keys are as yet available. Clausen (1971), Kramp (1961), Salvini Plawen

(1966), and Swedmark (1964) provide the most useful review articles.

6.4. Turbellaria.—This fairly large group (about 6,000 species to date) is

represented in all marine and freshwater biotopes, often as one of the most abundant taxa. Except for many polyclads (which are common on deep mud bottoms) ,

marine Turbellaria can be grouped as a whole in the meiofauna.

Determination requires live material, although in some groups, such as the

Acoela, sectioning is necessary. For comparison with earlier descriptions, which

were based mostly on preserved specimens, sectioning also is necessary. For original descriptions, examination of live and sectioned material should be interrelated.

For qualitative studies, samples of sediment to the order of 10 liters should

be used. Quantitatively, 100 ml to 1 liter samples are recommended. Core samplers

of most common types can be used effectively (Section 3.2.3.1). Preserved sediment will not yield identifiable specimens.

Animals can be extracted by the anaesthetization technique (Section 4.2.1.2)
38 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

or Uhlig Seawater Ice Technique (Section 4.2.1.1); both give quantitative data.

Deep-sea Turbellaria should not be anaesthetized. For mud samples the Swedmark

technique is effective (Section 4.2.2.2). Most Turbellaria can be extracted using

the deterioration technique (Section 4.2.1.3). Examination of live material after

anaesthetization with MgCl^ (except for deep-sea forms) is recommended.

Fixation can be preceded by anaesthetization, but for many forms contraction

during fixation is an advantage as many minute organs (e.g., canals) become more

compacted and easy to see. Warm Bouin's fixative or a warm corrosive sublimate

solution (acidulated with acetic acid after killing) is excellent. For electron

microscopy (or l-fi to 5-/t sections) phosphate-buffered glutaraldehyde is recommended. For fixation of live material, especially for type collections, the addition

of a drop of glycerin:formalin (1:1) followed by a slide sealer such as Murrayite

(see Appendix, Section 10.1.18) is recommended.

No modern comprehensive monograph is available; for smaller groups see Ax

(1955, Otoplanidae), Dorjes (1968, Acoela) and Luther (1955, Dalyelliidae).

Excellent starting aids are Karling's (1963) and Luther's (1960, 1961, 1962, 1963)

investigations of the Fennoscandian fauna.

6.5. Gnathostomulida.—The Gnathostomulida, although only recently described, have been shown to be cosmopolitan and very abundant. They occur in

all types of sand with a high organic content from high-tide level to 100 m deep.

About 18 genera with 80 species are known so far.

Sampling procedures must take into account the fact that the Gnathostomulida prefer deeper sand layers that require sampling with a spade or corer in the

littoral (Section 3.2.4) and with a deep-scraping dredge in the sublittoral (Section

3.3.2). Samples of 5 to 10 liters are usually sufficient for qualitative studies; and

samples of 100 ml are suitable for quantitative studies.

The deterioration technique (Section 4.2.1.3) is best for extraction of animals

from qualitative samples. It is important, however, to let the sample deteriorate

for a considerable time, usually one week, since the Gnathostomulida are very

slow in coming to the surface of the sediment, probably due to their anaerobic

metabolism. Superficial sand layers, 250 to 500 ml, are then placed in an Erlenmeyer

flask (2,000 ml) and left for 10 minutes with isotonic MgCl2 solution (Section

4.2.1.2). Two washes with MgClg and one wash with seawater are filtered through

a 62-fi mesh diameter nylon net mounted on a plastic ring (height 2 cm, diameter

9 cm; see Section 4.2.2.2). The filter and deposit are then placed in a dish with

seawater, whereupon the animals tend to move through the net into the dish and

can be collected free from the sediment.

For quantitative work the same filter is used, but upside down. The deposit

is then washed into a dish. The Uhlig Seawater Ice Technique (Section 4.2.1.1)

also has been found to be effective. Since only live animals can be identified, whole

sediment preservation is unsatisfactory.

In general, examination, fixation, and preservation are the same as for "soft"

fauna. For the purpose of determination by a specialist, drawings (and photographs) of the following characters should be made on the living specimen: dimensions and proportions of body and head; number and exact location of sensory

bristles; position and dimensions of pharynx, ovary, bursa, testes, and male organs;

jaws and basal plate; and sperm. Furthermore, two whole mounts per species

should be made, one only slightly squeezed and the other maximally.

No keys are available as yet. For the systematics see Sterrer (in press, b), and

for reviews see Riedl (1969) and Sterrer (1969, 1971, and in press, b).
NUMBER 78 39

6.6. Nemertina.—Although an abundant, mainly marine phylum, the Nemertina (about 850 species, ranging in length from 1 mm to 30 mm) are not a major

group in the meiobenthos. Comparatively large volumes of sediment are necessary

to obtain an adequate number of specimens of the small forms.

From sandy sediments, samples of 5 to 7 liters are taken, and for extraction

the deterioration technique (Section 4.2.1.3) is used, allowing the sand to stand

3 to 5 days before the upper sediment layer is removed and treated with the

anaesthetization technique (Section 4.2.1.2). Mud samples should comprise 30

to 40 liters of sediment, and for extraction the Swedmark technique (Section

4.2.2.2) is recommended.

The systematics of the Nemertina is based on internal anatomy, since earlier

workers studied preserved material for which external characteristics were not

recorded. Thus, identification of live Nemertina often is exceedingly difficult. It

is essential to combine the anatomical account with a detailed description of the

external features of the animal.

The following details should be noted from the living animal: shape, arrangement of eyes, cephalic slits or grooves, main color and color patterns, behavior,

size of relaxed animal. Under phase contrast the proboscis armature (present in

Hoplonemertina) can be drawn. Probosics eversion can be achieved by adding a

few drops of 70 percent alcohol to the seawater containing the animal. The

proboscis can be cut off and squeezed under a coverslip. Statocysts and the number

of statoliths, if present, should be observed by squeezing preparations of whole

animals.

No satisfactory technique is available for the preservation of Nemertina

within the sediment samples. Before fixation all Nemertina must be relaxed slowly

(urethane, chloral hydrate, or MgCl2 are suitable). This may take from ten minutes to two hours. Animals are best fixed in Bouin's solution (5 hours or more)

and preserved in 70 percent alcohol, which should be renewed when it turns

yellow. Collecting procedures and treatment prior to preservation of benthic

Nemertina in general were outlined by Kirsteuer (1967a).

Microsections (5 to 7 fi) are necessary to study the internal organization. All

descriptions of Nemertina should include a graphic reconstruction of at least the

anterior body region (see Adam and Czihak, 1964; Kirsteuer, 1967b). Ecological

data are very important to new species descriptions and should be included also in

redescriptions. Useful keys can be found in Burger (1895), Coe (1905, 1943, 1954),

Correa (1961), Friedrich (1936, 1950, 1955, 1961), Hylbom (1957), Iwata (1954),

and Miiller (1968). Kirsteuer (1971) has reviewed the systematics and ecology of

interstitial Nemertina.

6.7. Gastrotricha.—Some 160 species of Gastrotricha from 34 genera are

known from marine and brackish water habitats. Densities as high as 400/cm3

have been recorded, but those of 2/cm

 3 to 10/cm 3 are common. The Gastrotricha

range from strap-shaped animals possessing lateral and posterior adhesive tubes to

bottle-shaped animals with a prominent caudal furca. Species with thin cuticles

are easily damaged, and even MgCl2 can cause cuticle blistering in some species.

Sediment samples of up to 10 liters of sand can be easily handled for qualitative studies, although for quantitative work 10 cm

 3 has been found to be preferable.

Animals can be extracted quantitatively from the sediment by using the Uhlig

Seawater Ice Technique (Section 4.2.1.1) or the anaesthetization technique (Section 4.2.1.2). With the Uhlig technique, the extraction efficiency varies from 60
40 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

to 90 percent, depending on the species. Extraction efficiency of 95 percent can be

achieved by the anaesthetization technique but observational efficiency is reduced.

Thus, both methods probably give equivalent qualitative data. As MgCL is detrimental to some species, the Uhlig technique is preferred. Whole-sediment

preservation in 5 percent formalin must be preceded by relaxation, using MgCl2

or propylene phenoxytol. Staining the sample with Rose Bengal is the best technique for subsequent extraction.

Whole mounts are best prepared by adding increasing mounts of formalin to

relaxed specimens. The animal can then be transferred by means of an Irwin loop

(see Appendix, Section 10.1.16) to a solution of one part glycerin and one part

formalin (10 percent). The water is then evaporated, leaving pure glycerin. The

specimen can then be mounted in glycerin, using a Cobb aluminum frame (see

Appendix, Section 10.1.19), allowing examination of both dorsal and ventral

surfaces.

For taxonomic purposes the following should be especially noted: size of

relaxed animals, caudal appendages, other appendages, adhesive tubules, scale

pattern, ciliation pattern, digestive tract, reproductive system, and protonephridia.

Keys published since 1950 include those by Boaden (1963b,c), Brunson

(1959), Clausen (1965a,b), d'Hondt (1965, 1966), Kaplan (1958), Pennak (1953),

Voigt (1958), and Wieser (1957a). Hummon (1971) has reviewed the status of

the systematics of Gastrotricha.

6.8. Kinorhyncha.—The Kinorhyncha are small (less than 1 mm) and,

although abundant in certain areas, have been little studied quantitatively.

Typically they are mud dwellers and are least abundant in sandy biotopes. Sublittoral deposits yield much fewer specimens than littoral ones. Most Kinorhynca

are restricted to the upper 1 to 2 cm of sediment, although Catena Gerlach (a

sand beach inhabitat) has been found to a depth of 90 cm.

Qualitatively, Kinorhyncha are best obtained by using an epibenthic sledge

(Section 3.3.2) in which 10 to 20 liters of interface sediment are collected. Quantitative samples can be made by using corers or subsampling with cores from a

grab (Section 3.3.2).

Whole mud samples can be preserved with 7 to 10 percent formalin or, less

satisfactorily, with 70 percent ethyl alcohol. It is best to sieve the sediment through

a 62-/X screen and preserve the retained material.

The most efficient extraction method for mud samples is the bubbling technique (Section 4.2.2.3), which can be 90 percent efficient. If less than two liters of

sediment are to be extracted, however, the method is less efficient and sieving

through a 62-/i sieve may be more satisfactory. Sorting of the fauna is greatly

facilitated by the use of an Irwin loop (see Appendix, Section 10.1.16). From

preserved sediment, Rose Bengal stained material is easiest to extract by the

sieving method, but not all animals stain.

Relaxation, preferably in propylene phenoxytol, is recommended (prior to

fixation) in 4 to 7 percent formalin or 70 percent ethyl or isopropyl alcohol

(fresh water contact for 1 to 2 minutes aids extrusion of the head). For histological

purposes hot Bouin's or Gilson's fixative are satisfactory, but fixation is difficult.

Preservation should be in 70 percent ethyl or isopropyl alcohol to which

glycerin has been added. Storage in anhydrous glycerin is excellent. Glycerin is not

so useful as a mounting medium because of difficulties in clearing, and Hoyer's

medium colored with Lugol's iodine is recommended. A double coverslip mount
NUMBER 78 41

using Cobb aluminum frames (see Appendix, Section 10.1.19) is essential. Murrayite (see Appendix, Section 10.1.18) is an excellent ringing compound.

Most specimens should be dorsoventrally oriented. If sufficient specimens are

available a few should be laterally oriented. Hoyer's medium eventually clears

eggs and sperm beyond recognition, and these should be noted immediately after

the animal is mounted as external sexual dimorphic characters are often lacking.

Spines and hairs are critical diagnostic characters for species identification

and great care should be taken in noting their presence or absence. Spines may

break off at the base; hence, a spine scar may be the only remaining evidence.

This is a difficult character and requires the utmost caution. In general the species

characters outlined by Higgins (1967, 1968, 1969a,b) should be followed.

No satisfactory keys have been developed. The monograph of Zelinka (1928)

and works by Higgins (1967, 1968, 1969a,b, 1971) should be consulted.

6.9. Priapulida.—Priapulida have only recently been found in the meiobenthos. They are known from the Red Sea and from Bermuda and Curasao, and

live in shallow water. Since no quantitative data are yet available, as much surface sediment (mud and muddy shell gravel) as possible should be collected.

The specimens obtained were extracted by sieving on a 62-fi sieve and sorting

the debris. The fixation and preservation techniques are identical with those

described for the Kinorhyncha (Section 6.8).

It is important that both larval and adult characteristics are noted. Extension

of the proboscis is essential, as the critical taxonomic characters are based on the

teeth. Only three references are available, those of Coull (1968), Remane (1963),

and van der Land (1968).

6.10. Nematoda.—The Nematoda usually are the most abundant metazoans

found in marine sediments. They generally occur in greatest numbers in the top

5 cm of sediment, so care must be taken not to disturb the surface layers while

sampling. Qualitative sampling of exposed sediments may be accomplished simply

by skimming off the upper layers with shovel, trawl, or similar device. The Hope

corer (Section 3.2.3.1) works well when collecting by diving in relatively shallow

waters, and the various grabs, corers, and dredges may be used there as well as in

deeper waters. Direct coring or subsampling grab samples (if the sediment surface

has not been disturbed) with a corer are recommended for quantitative samples.

Several extraction techniques have been developed by soil nematologists (see

Goodey, 1963), few of which serve well in the extraction of marine forms. The

modified Baermann funnel technique, used alone or in combination with screening, is not effective because marine Nematoda will not pass through the tissue

paper barrier used in that method.

The Seinhorst elutriator has been demonstrated to be one of the most effective

means of extracting Nematoda from soil. It has not been widely used by nematologists, however, because it takes nearly 30 minutes to process one sample if

operated as recommended, and the equipment is fragile, impractical to use in the

field, and difficult to clean.

A sugar flotation technique for soil Nematoda has been modified for extraction of marine Nematoda and other small invertebrates (Teal, 1960). While this

technique is effective in separating Nematoda from small particles having a substantially higher density, it is much less satisfactory for samples containing lowdensity organic detritus.

Perhaps the technique with fewest objections is the following sieving technique: Sediment (usually 100 ml or less) is thoroughly stirred in a container of
42 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

approximately one liter of seawater; after allowing the heavier particles to settle

for about 15 seconds, supernatant is poured through a sieve having approximately

50-/x openings; Nematoda and other material are rinsed in a second container by

pouring clean seawater through the inverted screen. Final separation of Nematoda

from detritus residues may necessitate hand sorting. Staining with Rose Bengal

may help in hand sorting, but not all Nematoda will stain and some may be

overlooked. Four or five sievings of each sample may yield better than 90 percent

extraction.

Many marine Nematoda attach themselves by caudal adhesive glands to solid

surfaces they contact, thus reducing the efficiency of extraction. To overcome

this problem Hopper (personal communication) has used a mixture of seawater

and fresh water while extracting Nematoda, thus temporarily disabling them by

osmotic stress. Some of the methods of relaxing might also be used for this purpose.

Nematoda should be relaxed at the time of fixation, as they otherwise bend

or coil. This may be accomplished with osmotic stress, or heat (50° C for 10

minutes), but these methods may impair specimen quality and often are impractical in the field. J. B. J. Wells (personal communication) found that in the

tropics hot water at a temperature of 50° to 55° C was ineffective in killing and

preventing coiling unless the treatment was prolonged up to one hour. He found,

instead, that heating rapidly to 60° to 70° C was very effective. Chemical relaxants

may be most effective and practical, but these materials and methods for their use

need further study.

Five percent formalin in seawater for at least 24 hours is recommended for

fixation. Alcohol should never be used unless necessary for histochemical purposes,

as this fixative may cause Nematoda to collapse and possibly shrink. Five percent

formalin buffered with CaCO3 is recommended for preservation. Anhydrous

glycerin also is an excellent preservative, and it is routinely used as a mounting

medium for the preparation of permanent microscope slide mounts.

Cobb aluminum frames (see Appendix, Section 10.1.19) should be used when

preparing microscope slide mounts, and coverslips should be supported with glass

beads (see Appendix, Section 10.1.20) corresponding in size to the diameter of

the Nematoda.

Identification of marine Nematoda is based on numerous anatomical features,

some of the more important being the cuticle, amphid, cephalic and somatic setae,

stoma, esophagus, tail, and reproductive organs (especially the male copulatory

apparatus). Most keys are outdated. The most extensive keys are those of de

Coninck (1965) and Wieser (1953, 1954, 1956), although they have inadequacies.

Hope (1971) has reviewed the current status of the systematics of marine

Nematoda.

6.11. Bryozoa.—Only two groups of meiobenthic Bryozoa are known. One,

the genus Monobryozoon Remane, is considered a typical member of meiobenthos.

The other group, the lunulitiform Bryozoa, are questionable components because

as adults most are usually larger than the upper size limit of meiobenthos and

they live on sand surfaces.

The genus Monobryozoon has only two species, one of which lives in mud

(Monobryozoon limicola Franzen) and one of which lives in "Amphioxus sand"

{Monobryozoon ambulans Remane). Monobryozoon limicola can be collected

with an Ockelmann sledge, whereas Monobryozoon ambulans is sampled with a

dredge or grab. The lunulitiform Bryozoa can be sampled with any gear. The

meiobenthic Bryozoa are not abundant and so far only qualitative sampling has
NUMBER 78 43

been used. Monobryozoon limicola has been found in fair quantities in the

Gullmar Fjord, Sweden (Franzen, 1960), and at Roscoff, France (F. Monniot,

unpublished observation) . Monobryozoon ambulans is exceedingly rare and is

known only from about 20 specimens obtained from Helgoland, Germany (Remane, 1936, 1938), and Robin Hood's Bay, England (Gray, in press, a). The

lunulitiform Bryozoa have been reported from eastern Africa and the northern

Indian Ocean (Cook, 1966), Gulf of Mexico (Lagaaij, 1963b), and Brazil (Marcus

and Marcus, 1962).

Animals can be extracted from mud by the Swedmark technique (Section

4.2.2.2), from "Amphioxus sand" by the anaesthetization technique (Section

4.2.1.2), and from sand in general by the deterioration technique (Section 4.2.1.3).

Extreme care must be taken to thoroughly examine all sample washes as Monobryozoon is the same color and of smaller size than the substratum it inhabits.

Examination under low magnification is sufficient for the identification process.

Animals can be relaxed in MgCL or propylene phenoxytol. Preservation in 6

percent formalin or 1.5 percent propylene phenoxytol and fixation in Bouin's

fluid or 6 percent formalin are recommended.

As so few species are known, keys are unnecessary. For Monobryozoon, see

Franzen (1960), Gray (1971b) and Remane (1936, 1938); for lunulitiform

Bryozoa, see Cook (1966), Lagaaij (1963a,b), and Marcus and Marcus (1962).

Gray (1971b) has reviewed meiobenthic Bryozoa.

6.12. Brachiopoda.—Up to the present time only one species of Brachiopoda,

Gwynia capsula (Jeffreys), is known to occur in meiobenthos. It is about 1 mm in

size and possesses primitive anatomical characters. The species is known from the

shallow sublittoral in sediments containing a high proportion of shell debris. The

animals live inside fragments of serpulid tubes or in crevices on shell fragments of

Gastropoda, barnacles, and other Brachiopoda.

Dredging (Section 3.3.2) is the most satisfactory method of sampling. Samples

of 1 liter yield sufficient specimens, and 75 to 80 percent alcohol is a good preservative. Since Gwynia Jeffreys is sedentary, no techniques of extraction can be employed. For fixation 75 percent alcohol or neutral formalin can be used. Bouin's

fixative is recommended. Whole mounts can be made in Canada balsam for histological studies for which the shell is not to be preserved.

Live specimens should be used for observations on behavior and study of

external characters. Animals can be stored for long periods in embryological cups

in a small volume of seawater which should be changed twice a week. Under such

conditions, the animals probably feed on bacteria or on dissolved organic matter

(McCammon, 1969) and it is not necessary to add food. For literature, see

Beauchamp (1960) and Swedmark (1967, 1971).

6.13. Archiannelida.—The Archiannelida include a heterogeneous group of

five families ranging in size from 250 y, to over 2 cm long. Most Archiannelida live

in sand or mud and are interstitial forms. Troglochaetus Delachaux is the only

freshwater genus. The Archiannelida can be easily damaged by most extraction

techniques and, for accurate identification, living animals in good condition are

essential. Since the Protodrilidae and Saccocirridae have to be sexually mature

before they can be identified as to species, samples have to be taken at different

seasons.

Most Archiannelida occur below mean tide level. For qualitative samples it is

necessary to bring 1-liter samples of sediment to the laboratory for extraction.
44 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Quantitative samples can be taken with almost any core sampler. Subtidally,

dredges, cores, or the Ockelmann sledge can be used effectively.

For live animals, the optimum extraction method is the anaesthetization technique (Section 4.2.1.2). The Uhlig Seawater Ice Technique (Section 4.2.1.1) or

a simple seawater wash also can be used. For mud samples, the Boisseau technique

(Section 4.2.1.4) has been found to be excellent. Whole sediment samples can

be fixed with 10 percent formalin but should be stored in 70 percent alcohol.

From such preserved samples, the Boisseau technique (Section 4.2.1.4) has been

found to be effective for extraction.

Animals, after relaxation, should be fixed with Bouin's, Helly's or Zenker's

fixative. Carm-alum (5 percent) and light green have been found to be useful for

the study of internal organs, especially gonads and salivary glands in the Protodrilidae. After clearing in butyl alcohol, mounting should be made in Canada

balsam.

The principal characters used in identification are behavior, color, size and

number of segments, characteristics of epidermis (ciliation, sensory hairs, glands),

reproductive and associated organs, and setae when present.

Mature specimens often are white, and on live examination they can be

separated easily. Internally, the salivary glands, number of fertile segments, lateral

organs and spermaducts (for Protodrilidae), position and shape of spermducts

and sacs and of oviducts and spermatheca (for Saccocirridae), and size of ova and

their number per segment should all be noted. The shape of the head of the

spermatozoa is a good character even on preserved specimens, especially for the

Polygoridae and Nerillidae (Jouin, in press).

Outdated keys appear in works by Fauvel (1927), Remane (1932), and

Hempelman (1931). Keys for the Saccocirridae have been published by Alikunhi

(1948), Jouin (1971), and Wu and Yang (1962) ; for the Polygordidae by Marcus

(1948) ; and for the Nerillidae by Jouin (1971). Other important references include Aiyar and Alikunhi (1944), Ax (1967), Boaden (1961), Fraipont (1887),

Gray (1969), Jagersten (1944, 1952), Jouin (1966), Jouin and Swedmark (1965),

Kirsteuer (1966, 1967c), Pierantoni (1908), Swedmark (1959a,b), Tenerelli

(1967), Westheide (1967b), and Wieser (1957b). Jouin (1971) has reviewed the

status of the systematics and ecology of Archiannelida.

6.14. Polychaeta.—Approximately 70 families including 10,000 species of

Polychaeta are known. The group is heterogeneous, with species varying in size

from only a few millimeters to more than 3 meters. Representatives of many

families are found in the meiofauna, and the literature on the meiofaunal species

is diffuse. The characters needed for identification differ greatly within the families.

Qualitative samples using the technique described in Section 4.1.1 are

recommended. For quantitative littoral sampling, subsampling with a core (3 to

5 cm long) into the sides of a pit (Section 3.2.4) is satisfactory. In deeper layers

with subsoil water, an iron corer can be inserted vertically into the sediment. For

sublittoral sampling, the techniques given in Section 3.3 are satisfactory. Extraction can be effected by using the Uhlig Seawater Ice Technique (Section 4.2.1.1)

for sandy sediments or the deterioration technique (Section 4.2.1.3) for mud.

The total sediment sample can best be preserved in 10 percent formalin.

There are no special preservation methods needed for the Polychaeta, except that

alcohol should not be used for fixation. After fixation, it is well to transfer the

specimens to 70 percent alcohol. The general provisions described in Section 5

should be observed.
NUMBER 78 45

Histological sections usually are not necessary for species determinations.

Differences in specific characters of smaller animals, however, are slight. Some

delicate features such as color, shape of body appendages, glands, etc., can be

seen only in the living animals. Thus, a combination of living and fixed material

is optimal. Anaesthetization with isotonic MgCl2 or MgSO4 is often necessary for

observation of living animals as well as for fixation. An addition of a small amount

of acetic acid often causes the evagination of the proboscis, which is important for

identification in some families such as the Phyllodocidae. Clearing in glycerin or

oil of cloves is profitable for recognition of specific internal features in fixed whole

animals. For a complete examination of setae it often is necessary to mount in

lactophenol (one part lactic acid and one part phenol; Gee, 1964). The slides are

warmed gently before applying a coverslip. Lactophenol partially dissolves the

body tissues, leaving cleared chitinous setae.

The general characters used in identification include shape of anterior part

of body; number and shape of tentacles and cirri; sensory organs; number, arrangement, and exact shape of setae in parapodia; shape of pygidium; shape and

position of proboscis; color in living and fixed specimens; and measurements of

length. Special characters of each genus have to be added.

General works on the Polychaeta that cover keys to the families, genera, and

species include those of Berkeley and Berkeley (1948, 1952), Day (1967), Fauvel

(1923, 1927, 1953), Hartman (1968), Pettibone (1963), and Uschakov (1955,

1965). A nearly complete list of meiofaunal Polychaeta exists only for interstitial

species of sandy biotopes (Laubier, 1967). Publications dealing with some meiofaunal genera include the following: for Pisione Grube (Pisionidae), Aiyar and

Alikunhi (1940), Alikunhi (1947, 1951), Hartman (1939), Laubier (1967),

Siewing (1955a), Stecher (1968), Tenerelli (1965); for Eteonides HartmannSchrdder, Hesionura Hartmann-Schrdder, Mystides The"el, and Protomystides

Czerniavsky (Phyllodocidae), Hartmann-Schroder (1963), Laubier (1967); for

Hesionides Friedrich and Microphthalmus Metschnikow (Hesionidae), Ax

(1966), Laubier (1968), Westheide (1967a), Westheide and Ax (1965); for

Petitia Siewing (Syllidae), Siewing (1955b); for Syllidae in general, Cognetti

(1957) ; for Psammodrilus Swedmark and Psammodriloides Swedmark (Psammodrilidae), Swedmark (1955, 1959a); for Stygocapitella Knollner and Parergodrilus Reisinger (Stygocapitellidae), Karling (1958). The "Catalogue of the

Polychaetous Annelids of the World," by Hartman (1959, 1965), should be consulted for questions of nomenclature. Westheide (1971) has reviewed the interstitial Polychaeta.

6.15. Oligochaeta.—Although the Oligochaeta are predominantly freshwater

and terrestrial, some families are marine. The coarse and medium sand of the

supralittoral zone is a highly favorable habitat, but "Amphiouxus sand" and

muddy sands also are inhabited. Taxonomic characters are not likely to be easily

damaged.

Samples are taken with coring devices, the Ockelmann sledge, or grabs. The

sample size varies from 100 ml to 0.1 m 3 in poor sediments. Whole sediments samples can be preserved in 5 to 10 percent formalin, but 80 percent ethyl alcohol is

best for long storage.

Animals can be extracted qualitatively by simply washing and filtering on a

nylon screen. Quantitative samples can be extracted by the Uhlig Seawater Ice

Technique (Section 4.2.1.1) or the Boisseau technique (Section 4.2.1.4) for
46 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

preserved sediment. With mud samples, the Swedmark technique (Section 4.2.2.2)

is preferred.

For living animals, 5 percent alcohol or chloroform vapor is a good anaesthetic. J. B. J. Wells (personal communication) found that tropical Oligochaeta can

be relaxed and killed by heating water rapidly to 60° to 70° C, a technique that

was almost 100 percent effective. Fixation can be effected in 5 percent formalin,

but for storage 70 percent alcohol should be used. Bouin's fixative should be used

for histological purposes. For mounting in water, miscible media such as polyvinyl

lactophenol is recommended. Permanent mounts should be stained in chlorophydric carmine or acetic haematoxylin. Canada balsam is recommended for permanent mounts.

The haploid chromosome number is a useful aid for species determination.

Acetic orcein (orcein, 1 g; glacial acetic acid, 45 ml; distilled water, 55 ml) has

been found to be a successful chromosome stain. The acetic orcein is usable for

only one month. With live animals, considerable practice is necessary. The examination of immature specimens generally serves no useful purpose in the process

of identification.

The principal criteria used in species identification are coloration, size,

number of segments, setae, brain morphology, shape of gut and diverticula, septal

glands, nephridia, blood vessels, and shape of male reproductive organs. Useful

keys to the marine Oligochaeta can be found in Brinkhurst (1963, 1965, 1966a,b),

Brinkhurst and Cook (1966),Bunke (1967), Cook (1969), Lasserre (1964,1966,

1967a,b), Nielsen and Christensen (1959, 1963a,b), and Sperber (1950). Lasserre

(1971) has reviewed the taxonomy and ecology of marine Oligochaeta.

6.16. Mollusca.—The Mollusca are represented in the meiofauna by Solengasters, a few Prosobranchia, Scaphopoda, and Protobranchia, and by Opisthobranchia. The latter includes species belonging to the orders Acochlidiacea (12

species in 1969) and Philinoglossacea (2 species in 1969) and the aeolid family

Pseudovermidae (8 species in 1969). As yet, the Solenogasters, which are known

from the European Mediterranean and Atlantic coasts and the northwest Pacific

coast of America (Friday Harbor), are incompletely studied. Scaphopoda

(Siphonodentalium M. Sars and Cadulus Philippi) and some Protobranchia are

represented in the meiobenthos in subtidal sediments to depths of 3,000 m.

For qualitative sampling in sand and mud, epibenthic sledges such as the

Ockelmann sledge (Section 3.3.2) are most satisfactory. Other sledges and grab

samplers have been used with success. For quantitative sampling in mud, corers

(Section 3.2.3) or the Petersen or Smith-Mclntyre grab (Section 3.3.2) covering

an area of 0.1 m 2 are used.

Eighty percent alcohol is a suitable preservative. Extraction techniques include the Uhlig Seawater Ice Technique (Section 4.2.1.1) and the Swedmark

technique (Section 4.2.2.2). The specimens should be anaesthetized (Section

4.2.1.2) carefully for live examination and before fixation. Bouin's fixative is

recommended as a standard, but any fixative for histological purposes may be

used. If shells (Scaphopoda and Protobranchia), spicules (Acochlidiacea), or

scales (Solenogasters) must be preserved, acid fixatives must be avoided and

buffered formalin used. Whole mounts can be made in Canada balsam.

Spicules, scales, epidermal glands, radula, statocysts, and extension of cilia

should be studied on living material. Most of the anatomy can be studied on

squeezed specimens. The radula should be analyzed in detail, using the formula

n (2.1.2) where n = number of radula series, 2 = one median plate plus one
NUMBER 78 47

lateral plate, and 1 = central tooth. Keys for the identification of species have been

published for the Acochlidiacea (Odhner, 1952; Marcus, 1953; Swedmark, 1971),

for the Pseudovermidae (Salvini Plawen and Sterrer, 1968), and for the Scaphopoda (Muus, 1959). Other useful references include Marcus and Marcus (1954)

and Marion and Kowalevsky (1886).

6.17. Mystacocarida.—The Mystacocarida are morphologically and behaviorally distinctive; thus, they are easy to sample. Where present they are

abundant, and since they do not cling to the substratum they are easy to extract.

Sediment samples from holes 10 cm in diameter and 40 cm deep are extracted

in the field following the technique described in Section 4.1.1. The animals are

durable and may easily be brought back to the laboratory alive, or they may be

preserved in the field. Four percent formalin is the most convenient field preservative despite its disadvantage of killing the animals in a variety of contorted positions, thereby making uniform measuring techniques impossible. Alcohol tends

to distort animals less than formalin, but it is disadvantagous for field use because

it forms a precipitate with seawater.

Where subsequent measurement is desired, it is best to bring the live filtrate

back to the laboratory and pick out the desired individuals and kill them in lactic

acid which has been stained lightly with methyl (not methylene) blue. This kills

the animals in a uniform, relaxed position and stains them perfectly for microscopic investigation of external anatomy, which should be carried out in unstained

lactic acid. The animals may be left in lactic acid for a maximum of several weeks.

Dilute lactic acid (1:1) can be used to kill the entire filtrate in the field, but later,

during sorting, it is difficult to see the animals in this medium. Formalin is best

for long-term preservation. Whole sediment samples may be preserved in formalin

and extracted by an abbreviated decantation technique (Section 4.1.1).

All present evidence indicates that Mystacocarida are extraordinarily conservative; the species are differentiated on the basis of just a few slight details.

Therefore, it is important to note even subtle differences between populations.

Ten to twenty individuals from each population should be checked for individual

variation.

Delamare Deboutteville (1960) and Hessler (1971b) give introductions to

the literature on Mystacocarida. The works of Dahl (1952), Delamare Deboutteville (1953), Hessler and Sanders (1966), Noodt (1954), and Hessler (1969) are

especially useful in systematic studies.

6.18. Ostracoda.—Benthic marine Ostracoda range from 0.4 to 1.5 mm in

size. They occur on sand, mud, and algae at all depths. Some species are truly

interstitial while others are epibenthic. Most species occur in the upper few

centimeters of sediment but are not generally abundant. Thus, large quantities

of sediment must be collected and sorted. One hundred milliliter samples of

sediment may give from 10 to 300 animals. Probably a more suitable sample size

for littoral work is 0.1 m2. Coring devices usually do not provide enough living

specimens, so dredges and grabs are preferred for sublittoral work. Deep-sea samples of at least 5 liters of surface sediment usually are necessary to obtain enough

specimens.

Animals can be extracted from the sediment by sieving. Most species are

retained on 140-/* sieves. Care should be taken that highly ornamented shells are

not damaged. The Boisseau technique (Section 4.2.1.4) or the bubbling technique

(Section 4.2.2.3) can be used but the sediment must be reexamined after using

these techniques as separated valves and highly calcified species often remain.
48 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Buffered 5 percent formalin or propylene phenoxytol can be used to preserve

the whole sediment. Though little histological work has been done on Ostracoda,

Bouin's fixative is recommended. Animals can be preserved in 5 percent buffered

formalin or 70 percent alcohol. Buffering is essential for whole sediment samples

to prevent decalcification of the valves. Calcium carbonate, hexamine, and borax

can be used as buffering agents.

Ostracoda are best dissected in glycerin and the dissected appendages left in

glycerin or mounted permanently in glycerin jelly. Glycerin mounts, if sealed, are

satisfactory. Polyvinyl lactophenol is a useful semipermanent medium for mounting appendages; however, it cannot be used for valves because it causes decalcification. CMC-S (see Appendix, 10.1.21) is a water-soluble staining and mounting

medium, appendages can be arranged in position prior to the coverslip addition.

It is preferable that the valves be mounted on micropaleontological slides.

The specimens should be glued to the slides with a very slight amount of gum

tragacanth. A few drops of formalin, phenol, or oil of cinnamon can be added to

the tragacanth to prevent the development of molds. As the glue is water-soluble,

the valves can be removed from the slides.

Particular attention should be given to a description of both right and left

valves, appendages, and copulatory organs, and distinction should be made between molts and adults. Moore (1961) and Morkhoven (1962) give detailed

techniques for the study of ostracod valves and some information on appendage

morphology. Morkhoven (1963) gives generic descriptions and references to

studies of appendage morphology of all post-Paleozoic Ostracoda. Hartmann

(1964) has integrated the neontological and paleontological classification schemes.

Hulings (1971b) has reviewed the status of the taxonomy and ecology of marine

benthic Ostracoda.

6.19. Copepoda.—Next to the Nematoda, the Copepoda are the most

abundant group in the meiobenthos. Probably less than 50 percent of the existing

species have been described. Copepoda inhabit not only sand and mud areas but

are associated with algae and other biotopes. Sediments may be sampled fairly

easily; however, sampling of other biotopes is more difficult, and this is especially

true for the Copepoda, most of which swim and jump actively and are not, therefore, sampled by traditional techniques.

Qualitative sampling is easy for most biotopes, and samples of sand, mud,

algae, etc., can be easily obtained. No satisfactory quantitative techniques are

available for biotopes other than sediment. Core samplers (5 cm in diameter)

have been used effectively for sampling littoral and sublittoral sediments down

to SCUBA limits. Subsampling from grabs by cores or the Thiel "Meiostecher"

(Section 3.2.3.4) can be used in the sublittoral.

Whole sediment samples are best preserved in 5 percent formalin or 75 to 80

percent alcohol. Preserved sediment should be extracted as soon as possible since

some specimens become fragile in preserved conditions.

Copepoda can be extracted from the sediment by the anaesthetization (Section 4.2.1.2) or Boisseau (Section 4.2.1.4) techniques; however, washing and

decantation may give greater extraction efficiency. For preserved sediment the

Boisseau technique is highly effective, and the addition of Rose Bengal aids

sorting, although all the animals may not be stained.

Copepoda may be extracted from algae by shining a light from one side of a

dish and collecting animals that migrate to the light. Washing and sieving may

be more effective.
NUMBER 78 49

Preservation in 70 percent alcohol with 5 percent glycerin has been found

successful, and the addition of 1 percent phenol prevents growth of fungus in this

preservative. Animals can be dissected easily in lactic acid or glycerin, and stains

such as methyl blue (Section 6.15) may be useful. Mounts made in glycerin or

Zeiss W15 (see Appendix, Section 10.1.22) mounting medium ringed with Murray ite or Glyceel (see Appendix, Sections 10.1.18, 10.1.23) are preferred. Poly vinyl

lactophenol or CMC (Appendix, Section 10.1.21) also can be used for mounting.

Most taxonomic work has been on preserved material but some information

on species biology is available. The basic literature includes the monographs of

Lang (1948, 1965). Species and references subsequent to Lang's (1948) monograph are summarized by Bodin (1967). Noodt (1971) has reviewed the ecology

of the Harpacticoida.

6.20. Palpigradida.—The Palpigradida are very primitive Arachnida. They

were originally discovered in the interstitial environment by Cond£ (1965) and

later by Monniot (1966). Two species are known, Leptokoenenia gerlachi Cond^

from beaches of the Red Sea and Leptokoenenia scurra Monniot from high-tide

level on the beach at Pointe Noire, Republic of Congo. The Palpigradida are very

fragile and the techniques of collecting, fixing, and preserving them are the same

as those used for the Mystacocarida. For the systematics of this group, see Cond£

(1965) and Monniot (1966).

6.21. Halacaridae.—The Halacaridae generally are less than 600 fx in size and

thus may pass through the screens of many types of gear. While some species are

discontinuously distributed, others are continuously distributed. Thus, only

qualitative sampling techniques are usually employed. For sand and gravel forms,

four to five liters of sediment are sufficient. Animals can be extracted by using the

anaesthetization technique (Section 4.2.1.2) but substituting chloroform for

MgCl2. The animals can be preserved in 65 percent ethyl alcohol. The alcohol

concentration should be increased gradually to this concentration. Algal living

species can be extracted in a similar manner.

Whole sediment samples can be preserved in 65 percent ethyl alcohol. Screens

of 1 mm and 62 fi can be used. Nearly all mites are retained by a 62-/i sieve. Rose

Bengal staining aids sorting.

Halacaridae are best preserved in Hyrax, which has a high refractive index

(about 1.70) and is suited for resolving fine details. Because of the high refractive

index, it is essential to remove the soft parts as fully as possible, otherwise the

specimens are nearly opaque and cannot be studied with the microscope. The

most suitable method for removing the soft parts is hydrolysis with pepsin. KOH

and NaOH cannot be employed because they affect the cuticle adversely and

damage delicate setae. Even lactic acid and chloral hydrate weaken the cuticle

perceptibly and produce distortion in the final mounts.

The use of pepsin as a hydrolyzing agent imposes two additional requirements. First, the specimens must be pierced with fine needles to permit the

entrance of pepsin molecules; second, the specimens must not at any time be

placed in formaldehyde, which makes the soft parts insoluble in pepsin.

Halacaridae should not be placed in formalin, but some considerations make

it desirable or even necessary to have determinations made on such material. This

is possible by the use of alternative but less-desirable methods.

Material which has been preserved in formalin can be handled by mounting

it in Hoyer's medium, but this does not yield permanent mounts, and the specimens are less suitable for descriptive purposes because of the refractive index.
50 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Hoyer's mounted specimens can be made permanent by soaking off the coverslip

in water and transferring the specimens to anhydrous glycerin by the evaporation

technique. Permanent mounts can then be made by one of several available

techniques.

If formalin-fixed material is being sent for determination, it is very important

to point out that the specimens have been in formalin at one time; otherwise,

much time may be wasted in trying to mount the material in Hyrax.

No completely satisfactory keys are available. The most comprehensive keys

are those of Viets (1939, 1940) for the European fauna and of Newell (1947)

for the North American fauna. Newell (1971) has reviewed the problems of

studying subtidal Halacaridae.

6.22. Tardigrada.—The marine meiobenthic Tardigrada have been found

from the littoral to depths of 5,000 m. Only 25 marine species are known, 9 of

which are in one genus. Densities of 90/cm3 of sand have been recorded. The

animals are small (300 n) and generally are attached to sand grains or algae.

In general, 50-ml samples of sediment are adequate for taxonomic and ecological purposes. Tardigrada can survive long periods in dry sand and are not

damaged by such treatment, reviving immediately upon addition of seawater.

Whole sediment samples can be preserved in 10 percent formalin; however,

nearly dry sand samples can be used to keep Tardigrada indefinitely.

For extraction of live animals from sediment the anaesthetization technique

(Section 4.2.1.2) is preferable to the Uhlig Seawater Ice Technique (Section

4.2.1.1). The latter is only 20 percent efficient for Tardigrada. For quantitative

studies the Boisseau technique (Section 4.2.1.4) has been used effectively.

For preserved sediment, a simple washing technique is effective. Tardigrada

should be relaxed before being fixed in 70 percent alcohol or 10 percent buffered

formalin. Preserved specimens should be mounted in anhydrous glycerin. For

histological sections, alcohol Bouin's solution seems to be the best fixative. Preembedding in a drop of agar jelly prior to paraffin embedding is recommended.

The agar jelly can be stained with fast green to aid viewing. Sections are stained

in haematoxylin eosin.

Particular attention should be given to study of the head and cirri, mouth

and stylets, cuticular ornaments, genital openings, legs and toes, and claws and

secondary spurs. The definitive work of Ramazzotti (1962) includes a key to the

genera and species described. Renaud-Mornant and Pollock (1971) have reviewed

the systematics and ecology of marine Tardigrada and have included a key to the

genera.

6.23. Echinodermata.—Among the Echinodermata, only a few Holothuroidea

belonging to the order Apodida are known as permanent members of the meiofauna. Their body lengths range from 2 to 8 mm; all are more or less vermiform.

Along the European coasts, two species, Rhabdomolgus ruber (Keferstein) and

Leptosynapta minuta (Becher), are frequent in "Amphioxus sand." A third

species, Labidoplax buski M'Intosh, is common in subtidal mud. Rao (1968)

reported a new interstitial genus and species (Psammothuria ganapatii) from

beach sands in India.

Qualitative samples of subtidal sand can be taken with any type of dredge or

grab, and quantitative samples with a suitable corer. Qualitative samples of subtidal mud meiofauna can be taken with an Ockelmann sledge or other epibenthic

sledges (Section 3.3.2), and quantitative samples with a corer (Section 3.2.3).

For preservation, 75 to 80 percent alcohol is satisfactory. The extraction
NUMBER 78 51

technique commonly employed is that of Swedmark (Section 4.2.2.2). Anaesthetization (Section 4.2.1.2) using MgCL or MgSO4 should be carried out. For

fixation, Bouin's fluid is recommended, but it should be replaced by neutralized

formalin or 75 percent alcohol if the calcareous spicules are to be preserved. But

neutralized formalin, as a long-term preservative, seems to affect calcareous structures adversely. Whole mounts can be made in Canada balsam.

There are no special keys for the identification of species of Echinodermata

belonging to the meiofauna. For European species, Mortensen and Lieberkind

(1928) can be used. Swedmark (1971) has reviewed meiobenthic Echinodermata.

6.24. Tunicata.—Meiobenthic Tunicata live only in coarse sand at a depth

greater than 5 m. As they are rare, sediment samples should be as large as possible

(20 liters). Sampling with a dredge or anchor dredge (Section 3.3.2) is the best

method. The animals penetrate the sand and have a fine network of rhizoids to

hold themselves in place. Samples cannot be held longer than 48 hours if live

specimens are wanted. Preservation of whole sediment samples may be pointless,

as the animals are so rare.

The only successful extraction method found is that of simply washing the

sediment with a strong current of water on a fine screen. No quantitative methods

have been employed.

Animals can be preserved for 48 hours in buffered 5 percent formalin or in

75 percent alcohol. For fixation purposes, 10 percent formalin is effective. Specimens should be carefully dissected and the mantle, attached organs, and branchial

sac stained in Masson's hemalum for 1 to 3 minutes for the former two and for

5 to 10 minutes for the latter. Following a 10-minute wash in fresh water and dehydration in 70 percent and 95 percent alcohol and 10 minutes in butyl alcohol, the

mantle with attached organs and the branchial sac can be mounted in Canada

balsam on separate slides.

For taxonomic purposes the following should be noted: size (in three dimensions) , appearance and thickness of tunic, form of siphons and atrial tentacles,

branchial sac, neural gland, mantle, gut, gonads, kidney, incubatory pouch, larva,

and environmental data. No keys are available. Monniot (1965) should be consulted for subtidal species. For deep-sea ascidians see Millar (1959, 1960, 1964),

Monniot (in press), and Monniot and Monniot (1968, 1970). Monniot (1971)

has reviewed meiobenthic Tunicata.

7. Culture and Experimental Techniques

As the experimental treatment of the meiobenthos is still at an early stage, the aim

of this short survey is not to give an exhaustive description of various types of

experiments but rather to stress the great need for experimental work in the

hope of encouraging further studies.

The great number of continuously varying parameters in the field makes it

impossible to ascertain fully the correlation between a morphological character

and sediment type (long bristles/coarse sediment) or an environmental parameter

and a faunal distribution in a beach. The experimental approach, both in taxonomical and ecological studies, must, therefore, be highly recommended.

The taxonomist may be able to explain morphological modifications as

adaptations to different media by means of culture experiments or preference

experiments. Life cycle studies, already stressed as most important in taxonomic
52 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

descriptions, should be more standardized and controlled in order to deliver

maximal ecological information about the animals.

The ecologist has great need of tolerance and preference experiments for

explanation of field distributions (for a discussion see Jansson, 1967a-d, 1971)

and of behavioral studies to recognize the types of reactions of the organisms

(Boaden, 1963a, 1968; Gray, 1966a). In productivity studies there is a great need

for more information on community metabolism, including a preferred way of

measuring laboratory studies of the most important pathways of the energy flow

(Odum and Hoskin, 1958; Pamatmat, 1968).

The physiologist has an important role in studying the physiological background of the tolerance and preference reactions of the organisms and in explaining field distributions (Lasserre, 1969, 1970, in press) when simple ecological

experiments are not adequate. The very small size of meiofauna, however, precludes most of the common methods of experimental physiology (e.g., Warburg

apparatus, microrespirometers, inorganic estimation in coelomic fluid, etc.) There

difficulties merely present a challenge either to adapt the classical methods or

develop new ones. Some recent and refined microtechniques and ultramicrotechniques used in cytochemistry (see Glick, 1962) may be very useful in physiological experiments with meiofauna.

7.1. The Experimental Population.—The experimental population should

be described as clearly as possible. The following points should be recorded:

(a) name of species (with possible morphological aberrations), age, sex, physiological condition (eggs and/or sperm, grade of activity); (b) the habitat where

the animals were collected and the microclimatic conditions before and at sampling. Pretreatment in the laboratory under controlled conditions can in many

cases be recommended. The environmental conditions of the animals prior to the

experiment should be carefully noted.

7.2. Culture Techniques.—Although a vast number of meiobenthic organisms are known to species level, few life history studies have been completed. This

emphasizes the great need for culture experiments.

An adequate culture of organisms presupposes a good knowledge of the

species and of the environmental conditions in the normal habitat. The usual

approach to culture techniques is to imitate the natural habitat; this may be not

only the best method but also may be necessary in order to secure normal specimens. General principles and successful culture methods for different phyla are

given in Costello, Davidson, Eggers, Fox, and Henley (1957), Needham (1959),

and Kinne and Bulnheim (1970).

For the culture of meiobenthic animals a substratum usually is necessary.

Glass beads (Appendix, Section 10.1.20), which can be obtained lead-free, have

been found useful for culture work. Animals can be seen through the beads, and

the chemical nature of the beads is infinitely easier to control than the natural

substrata. This method has been used successfully with the archiannelid Dinophilus gyrociliatus Schmidt and the harpacticoid Amphiascus tenuiremis (Brady),

changing the seawater monthly. On the other hand, the interstitial harpacticoids

Paraleptastacus spinicauda Klie and Paramesochra constricta Nicholls have been

reared only in the natural substratum, using darkened vessels and changing the

seawater weekly. The gastrotrich Turbanella hyalina Schultze has been cultured

using glass beads but it reproduced only when bacteria were kept at a low level in

the culture vessels. Predatory species such as the kalyptorhynchid Turbellaria are
NUMBER 78 53

more difficult to culture. This is probably due to lack of knowledge of their food

requirements (Gray, unpublished). Other methods may be found in the taxonomic

literature of different groups, e.g., Nematoda (von Thun, 1966), Enchytraeida

(Christensen, 1956).

7.3. Experimental Techniques.

7.3.1. Field Techniques.—The most common type of field experiment is the

colonization experiment (Boaden, 1962; Renaud-Debyser, 1959, 1963). Sterile substratum is enclosed in permeable gauze bags or tubes open at both ends and buried

in the sediment. After the experiment the animals are sorted out and counted. In

such a way the attractiveness of graded sand (Boaden, 1962) or migration due to

tidal movements of the interstitial water (Renaud-Debyser, 1959, 1963) have

been studied. As the environmental parameters except for the one studied are

allowed to fluctuate in a normal way, these variations must be considered when

evaluating the results.

Other experiments such as measuring wave exposure and meiofaunal distribution or the tolerance of introduced meiofaunal populations in polluted areas

are possible but have not been attempted.

7.3.2. Laboratory Techniques.—In all laboratory experiments adequate control tests must be set up. Without suitable controls the results of the experiments

are open to criticism.

7.3.2.1. Tolerance Experiments.—A simple but much-criticized type of experiment is that for tolerance or resistance. The organisms are kept in graded

concentrations of a variable factor (e.g., salinities from 5 to 35 ppt), while the

other environmental factors are kept constant, and the survival of the organisms

is noted. While most other parameters are rather easy to keep optimal, the food

factor may cause some difficulties. If "natural" seawater is used for the experiment

a moderate bacterial growth usually develops, enough to keep the animals alive

even for months as noted by Jansson (1968) for Turbellaria, Oligochaeta, and

Harpacticoida. Sometimes a vigorous bacterial growth may spoil some of the

results. It may, therefore, be necessary to feed the animals regularly, intermittently

changing the water to new, sterile solutions. The time of exposure to air must be

correlated with the normal exposure endured by the organism.

For comparison with other investigations, the results should be expressed

as LD-50 values (lethal dose 50), which means the time or dose at which 50

percent of the experimental population died. Corrections should be made for

eventual deaths in the control population according to Abbott (for a description

of this type of experiment see Andrewartha, 1961). This method has been

criticized, and rightly so, for its crudeness. But it gives a rough estimation of the

end-points of the physiological ability of the organism and, in a simple way,

makes possible comparisons between quite different types of organisms. A careful

observation of the organisms during the experiment, noting whether they are

sluggish, in coma, etc., will provide information for further studies and give the

data a greater biological significance.

A better way of studying the tolerance is to find some way to measure the

"efficiency" of the organism. Good examples are growth, respiration, and locomotory rates of species.

It is better to observe the grade of recovery of the experimental population

after transfer to optimal conditions than simply to note LD-50 values. Quantification poses difficulties, however.
54 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

7.3.2.2. Preference Experiments.—In a preference experiment the organism

is allowed to choose the most favorable of two or more alternatives or the most

attractive area in a gradient. The divergence from the theoretical random distribution is taken as a measure of the intensity of the preference.

The number of animals should be as large as possible to represent the reactions of the whole population from which the organisms were sampled and, thus,

to represent the morphological and physiological variation within the species.

This emphasizes the need of working with several populations, isolated in time

and geographical distribution. The results should be treated statistically.

It is convenient to divide the preference experiments into alternative experiments and gradient experiments. In alternative experiments, the animals are

offered two or more alternative properties of the parameter studied, e.g., types of

food, grades of sand, salinities (Gray, 1966a,b, in press.fe; Jansson, 1962, 1968).

In gradient experiments, the organisms are transferred to a gradient of the

factor studied, e.g., temperature (Gray, 1965; Jansson, 1966). Compared with the

alternative experiment, not so many runs have to be made in order to establish

the preference area. Alternative experiments, however, usually are easier to carry

out and to evaluate.

The following important points have to be considered when making preference experiments: (a) parameters other than those studied must be kept constant;

(b) the alternatives or gradients must be stable during the experiment, and new

equipment must be tested; (c) the behavior of the organisms during experimental

conditions should be observed in order to make sure that the prerequisites for a

proper choice are fulfilled, i.e., that the animals really migrate in the gradient,

that no gregarious activity occurs, etc.; (d) time of experiment must be sufficiently

long for the animals to try every alternative or part of the gradient many times

(this requires introductory studies of the locomotory ability of the organism).

7.3.3. Physiological Techniques.—Where the ecologist ends his task, the

physiologist sometimes can carry it further to explain the metabolic processes

which determine the reactions of the animals. When pure ecological techniques fail

to explain the distribution of the organisms, physiological proofs may be shown.

The following important subjects, closely related to ecology, deserve serious

attention.

7.3.3.1. Osmoregulation and Ion Regulation.—Previous studies on osmoregulation and ion regulation are summarized by Kinne (1964), Potts and Parry

(1964), and Remane and Schlieper (1958). The diverging composition occurring

in sediments and in polluted areas stress the need of further research in connection

with field studies.

7.3.3.2. Oxygen Reactions.—The often low oxygen tensions in sediments indicate a great stress for the organisms (for a recent summary see Theede, Ponat,

Hiroki, and Schlieper, 1969). Tolerance to oxygen deficiency and hydrogen sulfide

should be studied. Wieser and Kanwisher (1961) and Theede, Ponat, Hiroki, and

Schlieper (1969) give techniques for studying such tolerances.

Oxygen consumption often has been taken as a measure of the metabolic

activity of the organism. Studies of the oxygen consumption in waters of different

salinities and ion composition similar to those of Lasserre (1969, 1970, in press)

give valuable information for the ecologist in explaining field distributions.

The most accurate method of measuring the oxygen consumption of small

organisms is the Cartesian diver technique (Glick, 1962; Holter and Linderstr0mLang, 1943). The minute volume capacity of the divers allows measurements on
NUMBER 78 55

single plank tonic animals (Zeuthen, 1947) and interstitial animals (Lasserre,

1969, 1970, in press). All measurements require statistical analysis (see Snedecor

and Cochran, 1967) ; thus, they should be replicated (Zeuthen, 1953).

7.3.3.3. Metabolism.—The food and feeding processes of meiofauna are incompletely known. The large group of "detritus consumers" certainly consists of

animals of great specialization. For example, one species may eat certain bacteria

present in detritus, with the rest of the material being returned as fecal pellets to

the detritus pool of the sediment and then perhaps consumed by another animal

species. Thus, another part of the detritus is utilized and competition is avoided.

There is an almost complete lack of information in this area, and much work is

needed.

7.3.3.4. Sense Physiology.—The organisms are guided in the field by their

sense organs. Little is known about the function and even morphology of the

various types of sense organs of the meiofauna. This, again, is an important field

for future work.

8. Biotopes other than Sediment

The meiobenthos of nonsedimentary habitats usually is different from that of the

sediment habitat, with only a few taxa being common to both. Few of the faunal

elements are directly associated with the macrobenthos (e.g., Cnidaria, Bryozoa).

The most important nonsediment biotopes are sessile and floating algae

(littoral, sublittoral and Sargassum), sessile macrofauna, vagile macrofauna, rock

surfaces, and dead organic materials.

Quantitative sampling of the nonsediment meiobenthos is difficult. Dahl

(1948), Hagerman (1966), A.-M. Jansson (1966), Ott (1967), Segerstrale (1944),

and Wieser (1952, 1959) have attempted such sampling in a variety of ways. The

extraction procedures outlined in Section 4 are generally applicable to the nonsediment meiobenthos.

The following data should be noted, in addition to those mentioned in Section 1: name of macrobiotic organism, if any; special characteristics (e.g.,

branched algae, bryozoan encrustations, mobility of organism); microclimate

associated with macrobiotic organism (e.g., light, currents, salinity changes,

oxygen, organic material) ; and microflora (e.g., diatom film on rock or algal

surface).

This section is necessarily brief because of the lack of expertise within the

group that prepared this manual. Knowledge of these important biotopes is poor

and further detailed studies on lines parallel to the sediment meiobenthos are

needed.
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1966. Erfahrungen mit einer neuen Kammer zur Lebendbeobachtung beweglicher Mikroorganismen. Research Film, 5: 642-649.

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1964. A Method for Meiobenthic Invertebrate Collection. American Zoologist, 4: 291.

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1964. Methods of Sampling the Benthos. Advances in Marine Biology,

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1965. Coup d'oeil sur les Gastrotriches Macrodasyoides du bassin d'Arcachon.

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1971. The Current Status of the Systematics of Marine Nematodes. In N. C.

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Thane-Fenchel, A.

1968. Distribution and Ecology of Non-Planktonic Brackish-Water Rotifers

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1969. Studies on the Resistance of Marine Bottom Invertebrates to OxygenDeficiency and Hydrogen Sulphide. Marine Biology, 2: 325-337.

Thiel, H.

1966. Quantitative Untersuchungen iiber die Meiofauna von Tiefseebdden.

Veroffentlichung des Instituts fur Meeresforschung in Bremerhaven,

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Thun, von W.

1966. Eine Methode zur Kultivierung der Mikrofauna. Veroffentlichung des

Instituts fiir Meeresforschung in Bremerhaven, sonderband 2: 277-280.

Tietjen, J. H.

1968. Chlorophyll and Pheo-Pigments in Estuarine Sediments. Limnology

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1969. The Ecology of Shallow Water Meiofauna in Two New England

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Tilzer, von M.

1967. Ein neues Gerat zur chemischen Probenentnahme aus interstitiellaquatischen Lebensraumen. Gewasser und Abwdsser, 44/45: 117-120.

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1964. Eine einfache Methode zur Extraktion der vagilen, mesopsammalen

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76 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

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1968. A New Aschelminth, Probably Related to the Priapulida. Zoologische

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1939. Meeresmilben aus der Adria. (Halacaridae und Hydrachnellae, Acari.)

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1968. Variations de la density du microbenthos sur une aire restreinte.

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1958. Gastrotricha. Tierwelt Mitteleuropas, 1 (4a): 1-74.

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1955. The Preservation of Natural History Specimens. Volume 1. Invertebrates. 205 pages. London: H. F. and G. Witherby.

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1958. The Ecology of Lagos Lagoon. V. Some Physical Properties of Lagoon

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1969. Irrigation of Submerged Marine Sands through Wave Action. Nature,

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1962. Resin Impregnation of Soil Samples. Nature, 193: 804.

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1967a. Monographic der Gattungen Hesionides Friedrich und Microphthalmus

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NUMBER 78 77

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1964. Some Quantitative Comparisons of Offshore Meiobenthos and Macrobenthos South of Martha's Vineyard. Limnology and Oceanography,

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1964. A Ball-Stoppered Quantitative Sampler for the Microbenthos. Ophelia,

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1964. Patterns in the Balance of Nature and Related Problems in Quantitative

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78 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Williams, R.

1971. A Technique for Measuring the Interstitial Voids of a Sediment Based

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Wu, B. L., and D. W. Yang

1962. The Taxonomy and Distribution of the Genus Saccocirrus (Archiannelida). Oceanologic and Limnologie Sinica, 4 (3—4) : 164—179.

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1928. Monographic der Echinodera. 396 pages. Leipzig: Wilhelm Engelmann.

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1947. Body Size and Metabolic Rate in the Animal Kingdom with Special

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Yentsch, C. S., and D. W. Menzel

1963. A Method for the Determination of Phytoplankton Chlorophyll and

Phaeophytin by Fluorescence. Deep-Sea Research, 10: 221-231.

Zinn, D.J.

1942. An Ecological Study of the Interstitial Microfauna of some Marine

Sandy Beaches with Special Reference to the Copepoda. Unpublished

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1968. A Brief Consideration of the Current Terminology and Sampling Procedures Used by Investigators of Marine Interstitial Fauna. Transactions of the American Microscopical Society, 87: 219-225.

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1946. Marine Microbiology. 240 pages.. Waltham, Massachusetts: Chronica

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10. Appendix

10.1. Equipment, materials, and manufacturers.

10.1.1. Telethermistors.

10.1.1.1. Rustrak Instrument Company, Municipal Airport, Manchester,

New Hampshire 03103, U.S.A.

10.1.1.2. Shandon Scientific Company Ltd., 65 Pound Lane, Willesden,

London N.W. 10, England. (Prices from $400.)

10.1.1.3. Spembly Technical Products, Trinity Trading Estate, Sittingbourne, Kent, England. (Price, $380.)

10.1.1.4. Veco Victory Engineering Corporation, Union, New Jersey 08611,

U.S.A.

10.1.1.5. West Instruments, The Hyde, Brighton 7, Sussex, England.

(Prices from $200.)

10.1.1.6 Yellow Springs Instrument Company, P.O. Box 279, Yellow Springs,

Ohio 45387, U.S.A.

10.1.2 Chart Records.

10.1.2.1 Amprobe Recorder Service Division, 630 Merrick Road, Lynbrook,

New York 11563, U.S.A.

10.1.2.2 Rustrak Instrument Company, Municipal Airport, Manchester,

New Hampshire 03103, U.S.A.

10.1.3. UNESCO Oceanographic Tables.

10.1.3.1. Libraririe de l'UNESCO, Place de Fontenoy, 75 Paris 7e, France.

10.1.3.2. M. M. Stationery Office, P.O. Box 569, London S.E. 1, England.

10.1.4. Conductivity Cells and Wheats tone Bridge.

10.1.4.1. Mullard Equipment, Ltd., Aboyne Works, Aboyne Road, Tooting,

London S.W. 17, England.

10.1.4.2 Pye Unicam Ltd., York Street, Cambridge, England. (Conductivity cell: Philips PR 9513 microcell, 0.5 ml; price, $35.)

10.1.4.3. Electronic Switchgear Ltd., 58 Wilbury Way, Hitchin, Herts,

England. (Conductivity meter Type MCI Mkv; price, $180. Conductivity cell

(3 ml), specify CJ1B; price, $30.)

10.1.5. Salinity-Temperature Bridge.—Electronic Switchgear, Ltd., 58

Wilbury Way, Hitchin, Herts, England. (Price, $500.)

10.1.6. Goldberg Refractometer.—American Optical Corporation, Scientific

Instruments Division, Buffalo, New York 14245, U.S.A. (Price, $210.)

10.1.7. pH-Redox Meters.—Analytical Measurements, Ltd., Feltham,

Middlesex, England. (Price, $200.)

10.1.8. Oxygen-Availability Meter.—Direct inquiries to: Dr. B. O. Jansson,

Askolaboratoriet, Trosa, Sweden. (Price, $170.)

10.1.9. Speedy Moisture Tester.

10.1.9.1. Griffin and George, Ltd., Ealing Road, Alperton, Wembley, Middlesex, England. (Price, $105.)

10.1.9.2. Thomas Ashworth and Company, Ltd., Sycamore Avenue, Burnley,

Lanes, England. (Price, $100.)

10.1.9.3. Townson and Mercer, Beddington Lane, Croydon, Surrey, England.

(Price, $95.)

79
80 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

10.1.10. Hydrogen Sulfide.—Hach Chemical Company, P.O. Box 907, 713

South Duff, Ames, Iowa 50011, U.S.A.

10.1.11. Crystic 28C—Scott Bader and Company, Ltd., Wollaston, Wellingborough, Northamptonshire, England.

10.1.12. Araldite.—C.I.B.A. (A.R.L.), Ltd., Duxford, Cambridge, England.

10.1.13. "Meiostecher."—Hydro-Bios Apparatebau GmbH, Am Jagersberg

7, 23 Kiel, Holtenau, West Germany. (Price, $5.)

10.1.14. Reineck Box Corer.

10.1.14.1. Hermann Siefken, Mozartstrasse 16, Wilhelmshaven, West Germany. (Price, §800, including shipping.)

10.1.14.2. Friedrich Leutert, D-3141, Erbstorf, West Germany. (Price,

$2,200, including shipping.)

10.1.14.3. Tetra Tech, 742 Herschel Avenue, La Jolla 92037, California,

U.S.A. (Price, $5,570.)

10.1.15. Siliclad.—Clay-Adams, Inc., New York, New York 10012, U.S.A.

10.1.16. Irwin Loops.—Welch Scientific Company, 7300 Linder Avenue,

Skokie, Illinois 60076, U.S.A. (Price, $3 per set of three.)

10.1.17. Eukitt.—O. Kindler, Deckglaszuschneiderei, Freiburg/Breisgau,

West Germany.

10.1.18. Murrayite.—Edward Gurr, Ltd., London S.W. 14, England.

10.1.19. Cobb Aluminum Frames.

10.1.19.1. Metaalwarenfabrik "Amstel," Gijsbrecht van Aermstelstraat

16-18, Amsterdam O, Netherlands.

10.1.19.2. Sartory Instruments, Ltd., 7 Steele Road, Chiswick, London W 4,

England.

10.1.20. Glass Beads.

10.1.20.1. Catathote Corporation, Microbeads Division, Jackson, Mississippi

39205, U.S.A.

10.1.20.2. Ballotini Manufacturing Company, Ltd., Barnsley, Yorkshire,

England.

10.1.21. CMC, CMC-5.—General Biological Supply Company, 8200 South

Hoyne Avenue, Chicago, Illinois 60620, U.S.A. (Prices $2 for 4 ounces; $7.25 for

1 pound.)

10.1.22. Zeiss W15.—Carl Zeiss, Inc. (Price, $1 per 30 cm3.)

10.1.23. Glyceel.—Edward Gurr, Ltd., London S.W. 14, England.

10.2. Wheatstone Bridge.—The Wheatstone bridge incorporates a magiceye detector unit in conjunction with a dip-type conductivity cell. Two of the

ratio arms of a conventional Wheatstone bridge are combined in a single

potentiometer so arranged that for maximum rotation of the dial the ratio change

(i.e., R3/R4) is exactly 100:1. The scale is calibrated from 0.1 to 10; hence, when

the bridge is balanced, the dial reading when multiplied by the resistance inserted

at R2 allows the resistance of the conductivity cell to be calculated, at balance, as

When the bridge is balanced the magic eye indicator shows the minimum

fluorescence. Figure 12 shows the circuit diagram for the Wheatstone bridge.

10.3. Oxygen Availability.—The oxygen availability can be expressed by the

formula

f

 _ it

1
NUMBER 78 81

R2

RESISTANCE BOX

9

I DETECTOR

MULLARD UNIVERSAL

BRIDGE

OSCILLATOR

CONDUCTIVITY CELL

FIGURE 12.—Circuit diagram of the Wheatstone bridge.

where / = flux of oxygen (96,500 coul), i, = amp, n = number of electrons involved in oxygen reduction at Pt surface (=4), A = electrode area (cm2

). For

example, if ^4 =0.11 cm2, then

* =

(4) (96,500) (0.11)

moles cm—2sec—*.

To convert moles cm~2sec~1 to g 02 cm~2 min- 1, multiply by 32 (mol wt 02) and

by 60 (sec/min). Then,
82 SMITHSONIAN CONTRIBUTIONS TO ZOOLOGY

Readings of it are made in microamps; thus, the formula becomes:

/

 =-o9TT (10"e

 S °2 cm"2 min-1).

The reference electrode can be made either by using a commercial pH reference

electrode (e.g., Beckman), or a self-manufactured one. A filter crucible of porosity

15 to 40 fi is filled with 0.1 N KC1. A wire is led through a tight-fitting rubber

stopper and is attached to a bundle of silver wool (available from any laboratory

apparatus supplier). The wire lead then can be attached to the oxygen availability

meter. In operation, the reference electrode must make a good contact with wet

sand, and care should be taken to avoid trapped air from remaining under the

crucible.

The naked platinum electrode is made by first attaching the platinum to a wire

lead, preferably of single strand copper. Soldering should be avoided as it interferes with the weak electrode current. The platinum is made to protrude from

the end of a glass tube and is sealed in. The surface area of the exposed platinum

should be calculated accurately in cm2 by using a micrometer screw.

For the circuit diagram (Figure 13), the units are as follows: B = 1.35v, Mallory

ZM-9; Pt = 1000 ohm; P2 = 150 ohm; i?x = 100 kohm; R2 = 1000 ohm, 1%;

R3 = 1000 ohm, 1%; R4 = 2000 ohm, 1%; R6 = 4000 ohm; Rt = 400 ohm;

R7 = 3900 ohm; /?„ = 35 kohm.

10.4. Preparation of Silicone-Coated Pipette.—A water-soluble silicone such

as Siliclad (see Section 10.1.15) can be used. This concentrated solution is diluted

into 100 parts of distilled water. The pipette should be washed thoroughly and

ELECTRODES ON

FICURE 13.—Circuit diagram of the Jansson-Oden oxygen diffusion meter.
NUMBER 78 83

rinsed in distilled water. The inside of the pipette is coated by forcing the silicone

solution through it. After complete immersion, it is removed immediately and

rinsed thoroughly with clear water to remove any excess silicone from the surface.

It then is air dried for 24 hours at room temperature. The pipette may be heated

at 100° C for 10 minutes to speed the curing process. The Siliclad solution gives

a coating that is physiologically inert and under normal conditions is unaffected

by heat, moisture, and seawater. Rinsing the pipette is all that is necessary to keep

it clean.
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84

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