Molluscan Research: Techniques for collecting, handling, preparing

Transcription

Molluscan Research 27(1): 1–50
http://www.mapress.com/mr/
ISSN 1323-5818
Magnolia Press
Techniques for collecting, handling, preparing, storing and examining small
molluscan specimens
DANIEL L. GEIGER1, BRUCE A. MARSHALL2, WINSTON F. PONDER3, TAKENORI SASAKI4 & ANDERS WARÉN5
1
Santa Barbara Museum of Natural History, 2559 Puesta del Sol Road, Santa Barbara, CA 93105, USA. Email: geiger@vetigastropoda.com.
2
Museum of New Zealand Te Papa Tongarewa, P.O. Box 467, 169 Tory Street, Wellington, New Zealand. Email: brucem@tepapa.govt.nz.
3
Australian Museum Sydney, 6 College Street, Sydney NSW 2010, Australia. Email: winston.ponder@austmus.gov.au.
4
The University Museum, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan. Email: sasaki@um.u-tokyo.ac.jp.
5Department of Invertebrate Zoology, Swedish Museum of Natural History, Box 50007, SE-10405 Stockholm, Sweden.
Email: anders.waren@nrm.se.
Abstract
Micromolluscs are small-sized molluscs (< 5 mm), and include the great majority of undescribed molluscan taxa. Such species
require special collecting, sorting and handling techniques and different storage requirements to those routinely used for larger
specimens. Similarly, the preparation of shells, opercula, radulae and animals poses some challenges for scanning electron
microscopy (SEM). An overview of experiences with various techniques is presented, both positive and negative. Issues discussed include those relating to storage of dry specimens and interaction of specimens with glass, gelatine and paper products,
handling techniques and storage in various fluids. Techniques for cleaning shells for SEM are described and compared, as well
as those for radular extraction. The interactions of chemicals used for the dissolution of tissue with calcareous micromolluscs
are described. Methods for handling and mounting small radulae for SEM are detailed and brief guides to SEM and light photography are given. An appendix listing details of frequently-used chemicals is provided.
Key words: Review, methodology, collection, preservation, storage, museology, SEM, radula, shell, Byne's disease
Table of Contents
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
Institutional Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . 2
Other abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
The workspace . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
Equipment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5
Sieves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
Microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Pipettes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Forceps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Microscissors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Scalpels. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Pin and needle holders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Pins and needles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Hairs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Brushes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Pliers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Drills. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Tool sharpening . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Bowls . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Collecting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Hand collecting methods . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Other methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
Narcotisation and relaxation. . . . . . . . . . . . . . . . . . . . . . . . 10
Wet micromolluscs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11
Sorting from bulk samples . . . . . . . . . . . . . . . . . . . . . . . . . 11
Fixation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11
Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12
Switching storage media . . . . . . . . . . . . . . . . . . . . . . . . . . 13
Boiling method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13
Dry micromollusc shells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13
Initial drying . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
Handling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
Specimen containers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15
Labels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15
Chemical deterioration . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16
Byne’s ‘disease’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16
COPYRIGHT © 2007 MALACOLOGICAL SOCIETY OF AUSTRALASIA
Glass ‘disease’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .16
Preparation of micromollusc shells for SEM . . . . . . . . . . . . . .16
Cleaning. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .16
Mounting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .17
Preventing charging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .18
SEM parameters. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .18
Specimen removal from stubs . . . . . . . . . . . . . . . . . . . . . . .21
Separating the valves of minute bivalves . . . . . . . . . . . . . .20
SEM imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .21
SEM preparation of animals . . . . . . . . . . . . . . . . . . . . . . . . . . .22
Preliminary inspection . . . . . . . . . . . . . . . . . . . . . . . . . . . . .22
Limpets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .22
Coiled gastropods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .22
Opercula. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .24
Removing the shell the fast way . . . . . . . . . . . . . . . . . . . . .24
Tissue preparation for SEM . . . . . . . . . . . . . . . . . . . . . . . . .24
Extraction of radulae from micromolluscs . . . . . . . . . . . . . . . .26
Standard method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .26
Maceration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27
Cleaning the radula . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .28
SEM mounting of micromollusc radulae . . . . . . . . . . . . . . . . .29
Orientation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .29
Special techniques for small radulae . . . . . . . . . . . . . . . . . .29
Very small specimens . . . . . . . . . . . . . . . . . . . . . . . . . . . . .30
Manipulation of radula . . . . . . . . . . . . . . . . . . . . . . . . . . . .30
Radula, histology and X-ray computer tomography . . . . . .30
Optical photography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .32
SLR camera (film or digital) . . . . . . . . . . . . . . . . . . . . . . . .32
Stereo-microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .34
Lighting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .34
Depth of field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
Positioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
Chemicals. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
Literature Cited. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35
Appendix 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .39
1
2
Introduction
The majority of biodiversity to be discovered and described
is of small to minute size (e.g., Bouchet et al. 2002). For
molluscs, that number is in the range of at least a hundred
thousand (Steitz and Stengel 1984; Brusca and Brusca 2003).
Bouchet et al. (2002) found that the modal size of molluscs
from New Caledonia is only 3 mm in the most diverse size
class of 1.9–4.1 mm, which contains a quarter of all
specimens sampled. As investigators working on small sized
molluscs, we have developed and assessed various
collecting, sorting and handling techniques that facilitate
their study. To our knowledge, there is no previous detailed
and comprehensive account of working methods for
micromolluscs, other than a few very general discussions
(e.g., Robertson 1961; McLean 1984; Kurtz 2005).
We give here a summary of our joint experiences, while
acknowledging that further refinement will inevitably be
needed. The notes given here arise from trial and error
experimentation by the authors over more than a century of
professional working years. While the observations reported
do not stem from controlled experiments, they provide
important observational data and a starting point for future
experimentation and improvements. Thus the methods
presented are neither exhaustive nor foolproof. In part, our
intention is also to provide guidelines that should assist
others to find the most efficient methods for them and to
avoid known problems. For that reason, we include
discussions of failed methods and remarks on some of the
problems encountered. There rarely is a single ‘best’
technique for any given procedure and the techniques used
by any given practitioner reflect personal preference and
individual modification to some degree. Although most of
the techniques described have been applied by us in marine
or freshwater settings with shelled molluscs, many if not
most of the techniques described here could also be applied
to terrestrial shelled molluscs. However, different techniques
to those given here may be necessary with shell-less species,
specifically those relating to collection and narcotisation. We
do not deal with methods relating to histology or
transmission electron microscopy as these are well covered
elsewhere. While we describe suitable equipment that can be
used for dissection of micromolluscs, we do not elaborate on
dissection methods and techniques.
A mollusc is here considered small if the largest
dimension of the animal or last whorl of the shell (if a
gastropod – even if tall spired) is less than 5 mm in size. The
smallest molluscs reach around 0.6 mm in adult size, but
many larval or juvenile stages are smaller. While we use the
term ‘micromolluscs’ for species that are less than 5 mm in
maximum dimension as adults, this is clearly arbitrary.
Figures 1–3 show some of the diversity of micromolluscs.
Institutional Abbreviations
AMS—Australian Museum Sydney, New South Wales,
Australia
BMNH—The Natural History Museum, London, Great
GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27
Britain
GNM—Natural History Museum, Gotenburg, Sweden
LACM—Natural History Museum of Los Angeles County,
California, USA
NHMB—Naturhistorisches Museum Berlin, Germany
NMNZ—Museum of New Zealand Te Papa Tongarewa,
Wellington, New Zealand
NSMT—National Science Museum, Tokyo, Japan
SMNH—Swedish Museum of Natural History, Stockholm,
Sweden
USNM—United States National Museum, Smithsonian
Institution, Washington (DC), USA
ZMO—The Zoological Museum, University of Oslo,
Norway
ZMUC—The Zoological Museum, University of
Copenhagen, Denmark.
Other abbreviations
CPD—critical point dried.
FST—Fine Science Tools (supplier of microtools).
HCl—Hydrochloric acid.
HMDS—Hexamethyldisilizane.
KOH—Potassium hydroxide.
MORIA—Microtool brand.
NaOH—Sodium hydroxide.
LaB6—Lanthanium hexaborite.
LCD—Liquid crystal display.
LED—Light emitting diode.
OsO4—Osmium tetroxide.
PVA—Polyvinyl acetate.
PVC—Polyvinyl chloride.
SCUBA—Self Contained Underwater Breathing Apparatus.
SDS—Sodium lauryl sulphate.
SEM—Scanning electron microscope, - microscopy, micrograph.
TEM—Transmission electron microscope, - microscopy, micrograph.
VPSE—Variable pressure secondary electron detector.
The workspace
Work with small molluscs is greatly facilitated by the use of
proper tools. It is perhaps not as important to use exactly one
model of something for a certain kind of work, but rather to
be familiar with a range of tools so some alternative options
are available.
When working with small objects, the timing of various
steps in a procedure is critical. Therefore, it is important that
tools and the workspace are clean and well organised. Also,
as in most laboratory situations, suitable precautions should
be taken when working with chemicals that are noxious,
toxic, flammable and corrosive (e.g., ethanol, formalin, HCl,
KOH, OsO4, HMDS: see Appendix, manufacturers’ Material
Safety Data Sheets). In respect of these concerns, a fume
hood with an extractor fan is an essential part of any
laboratory space.
TECHNIQUES FOR STUDYING SMALL MOLLUSCAN SPECIMENS
3
FIGURE 1. Selected SEM images of marine and freshwater micromolluscs illustrating their morphological diversity. A. Anatoma sp.
(Vetigastropoda: Anatomidae). B. Sinezona n. sp. Geiger unpubl. data (Vetigastropoda: Scissurellidae). C. Emarginula sp. (Vetigastropoda:
Fissurellidae). D. Biwakovalvata biwaensis (Prestion, 1916) (Heterobranchia: Valvatidae). E. Cingulina cingulata (Dunker, 1860)
(Heterobranchia: Pyramidellidae). F. Spirolaxis exornatus Bieler, 1993 (Heterobranchia: Architectonicidae). G. Amathina tricarinata
(Linnaeus, 1767) (Heterobranchia: Amathinidae). H. Cavolina sp. (Heterobranchia: Cavolinidae). I. Caecum gracile Carpenter, 1858, adult
(Caenogastropoda: Caecidae). J. Caecum sp., juvenile (Caenogastropoda: Caecidae). K. Orbitestella sp. (Caenogastropoda:
Orbitestellidae). L. Joculator ridicula (Watson, 1886) (Caenogastropoda: Cerithiopsidae). M. Microdaphnella trichodes (Dall, 1919)
(Caenogastropoda: Turridae). N. Triphora sp. (Caenogastropoda: Triphoridae). O. Epitonium sp. (Caenogastropoda: Epitoniidae). P.
Scaliola bella A. Adams, 1860 (Caenogastropoda: Scaliolidae). Q. Granulina sp. (Caenogatropoda: Cystiscidae). R. Parashiela sp.
(Caenogastropoda: Rissoidae). S. Stosicia incisa (Laseron, 1956) (Caenogastropoda: Rissoidae). T. Barleeia sp. (Caenogastropoda:
Barleeidae). U. Ringiculina doliaris (Gould, 1860) (Heterobranchia: Ringiculidae). Images: A–C, H, K–M, O, Q–R: DLG; D–G, I–J, N, P,
S–U: TS; C, H, L, M, O, Q, R: kind permissions of Henry Chaney and Kirstie Kaiser.
4
FIGURE 2. Automontage images of type specimens (NMNZ) of some New Zealand land snails (Heterobranchia: Pulmonata) (A,C,J,L:
dorsal views; B,D,F,G-I,K,M,O: apertural views; E,N: ventral views). Dimensions given are the maximum diameter. A. Phenacohelix
giveni Cumber, 1961, holotype, M.20254 (5.50 mm). B. Phrixgnathus murdochi Suter, 1894, holotype, M.88067 (5.60 mm). C.
Flammoconcha stewartensis Dell, 1952, holotype, M.5450 (2.10 mm). D. Fectola trilamellata Climo, 1978, holotype, M.47445 (2.85 mm).
E. Ptychodon takakaensis Climo, 1981, holotype, M.47451 (1.80 mm). F. Laoma spiralis Suter, 1896, syntype, M.83460 (2.90 mm). G.
Cavellia oconnori Dell, 1950, holotype M.4067 (3.85 mm). H. Helix pseudoleiodon Suter, 1890, syntype M.30484 (2.50 mm). I,N.
Climocella reinga Goulstone, 1996, holotype, M.129904 (3.02 mm). J. Phrixgnathus viridula caswelli Dell, 1955, holotype, M.6158 (2.38
mm). K. Allodiscus austrodimorphus Dell, 1955, holotype, M.6149 (5.10 mm). L. Suteria raricostata Cumber, 1962, holotype, M.16935
(6.70 mm). M. Charopa pseudocoma Suter, 1894, syntype, M.125163 (5.10 mm). O. Rhytida meesoni Suter, 1891, syntype, M.125139
(11.45 mm). Images: Raymond Coory (NMNZ) and BAM.
5
FIGURE 3. Selected micromolluscs illustrating their morphological diversity. A. Nucula declivis Hinds, 1843 (Taxodonta: Nuculidae),
shell length 3 mm. B. Nucula exigua Sowerby, 1833 (Taxodonta: Nuculidae), shell length 3.5 mm. C. Acila castrensis (Hinds, 1843)
(Taxodonta: Nuculidae), shell length 4 mm. D. Runcina coronata (Quartefages, 1844) (Cephalaspidea: Runcinidae). Field photograph of
live specimen with 28 mm lens reversed on bellows unit, illuminated with two flashes. At 8:1 magnification (animal approximately 3 mm
in length) depth of field becomes very shallow. E. Colpodaspis pusilla M. Sars, 1870 (Cephalaspidea: Diaphanidae, animal approximately
5 mm in length). Photograph of living animal with 50 mm macro lens on bellows unit, illuminated with two flashes. F. Cingula cingillus
(Montagu, 1803) (Caenogastropoda: Rissoidae) photographed in the field with bellows unit, extension ring, 50 mm macro lens, flash
illuminated. Shell length approximately 3 mm. Some blurring is apparent due to excessive closure of the diaphragm (f/11, fmax = f/4). G.
Julia sp. (Ascoglossa: Juliidae). Animal approximately 5 mm long. H. Murchisoniella sp. (Heterogastropoda: Pyramidellidae). 3 mm. I.
Discrevinia sp. (Caenogastropoda: Pickworthidae). Shell 2 mm long. J. Moerchinella sp. (Heterobranchia: Pyramidelloidea). Shell 1.8 mm
wide. K. (Caenogastropoda: aff Vitrinellidae). Shell 1.3 mm long. L. Gibberula sp. (Caenogastropoda: Cystiscidae). Shell 2.5 mm long.
Images: A–C: DLG, courtesy Paul Valentich-Scott; D–F: DLG; G–L: AW (courtesy Panglao 2004 Workshop/Philippe Bouchet).
Equipment
•
Important considerations include:
•
•
Keep tools clean and properly stored, to prevent
damage to their delicate tips.
A glass jar with paper on the bottom is good for storing
pipettes.
Fine paint brushes should be stored under cover in a jar,
with their handles resting on the bottom, to avoid dust
accumulation and deformation of the hairs.
6
•
•
•
•
•
Use some kind of rack or stand to keep them available
and ready for use.
Micro-pipette tips used in molecular biology (100–
1000 µl) make excellent tip and needle protectors.
Preferably use tools made of material that will not
deteriorate quickly in salt water (e.g., stainless steel).
For microscope work, a comfortable seat of the correct
height is essential. For fine manipulation, steady your
body by resting your elbows and wrists on the table, use
the back support of your chair, and place your feet
firmly on the ground. Consider breathing rhythm as it
moves the ribcage and arms.
Keep equipment clean to avoid deterioration and
contamination.
There are a number of suppliers of suitable equipment
who can also offer useful information (e.g., http://www.
finescience.com; http://www.mccronemicroscopes.com). We
do not illustrate most of the readily available tools, only
those that are custom made or enlarged views ordinarily not
shown (Fig. 4).
Sieves
For a more efficient examination of samples containing
a large proportion of sediment, the residues should be
divided into size fractions by using graded sieves (e.g., 10, 5,
2.5, 1.0 and 0.4 mm mesh size). If one wants all specimens,
including larval shells, 100 µm mesh is needed, while for all
adult species 0.4 mm is suitable. Fractions larger than 5 mm
can be examined with the naked eye. For 5–2 mm a low
power magnifier can be used, although a stereomicroscope is
preferable and gives a better yield as untypical molluscs are
more easily recognised. For smaller fractions a
stereomicroscope is essential. Commercially made sieves
and even shakers for banks of sieves are available, or screens
can be constructed from various sizes of wire mesh (Fig.
4H). Sieves can also be made by using short pieces of PVCpipe, 50–250 mm diameter (Fig. 4G). A piece of metal
(preferably stainless steel) mesh slightly wider than the pipe
can be placed on a piece of aluminum foil on a hot plate and
the pipe pushed down on it until the end of the pipe starts to
melt. At that point, put it on a cold surface, still with some
pressure, so the net does not separate from the soft plastic.
Trim off any surplus net and grind the edge to remove any
free wires. Instead of a net, a perforated sheet of stainless
steel can be used. It is a little more difficult to work with but
makes very sturdy sieves that are less readily clogged.
For fieldwork, collapsible nets with a fine mesh
(<0.5 mm), such as a cloth liner used in aquaria to rear
juvenile fish, or bags made from plankton netting are very
useful for initial removal of the finest sediments. Plastic or
metal kitchen sieves available from supermarkets are also
very useful and come in various mesh sizes, or sieves can be
made from commercially available wire mesh (Fig. 4H).
Sieving should preferably be done in water to avoid
unnecessary abrasion or damage to fragile specimens.
FIGURE 4. Selection of tools used for the study of micromolluscs.
A–C. SEM images of needles at identical magnification. A. Sewing
needle. B. Insect pin. C. Ultrasharp tungsten needle. Scale bars =
100 µm. D. Forceps made from bamboo. E. Modified Pasteur
pipette to dispense small droplets. The Pasteur pipette is connected
to a rubber tube that is closed at the distal end. Compression of the
rubber tube allows the dispensing of minute droplets of liquid. F.
Mesh scoop used to concentrate shell grit in situ. G. Sieves made
from plastic tubing and wire mesh screen. H. Set of nested screens
made from wire mesh available at hardware stores. The screens are
nested and fit into a tightly closing plastic container. 50 ml Falcon
tubes can be placed inside, which may hold specimens, ethanol or
MgCl2. A dishwashing brush used for rock-brushings is also shown.
Images: A–C, G–H: DLG; D: TS; E: AW.
7
Microscope
A good-quality dissecting microscope is essential. A
minimum magnification of 50x is desirable. In general, those
with stepped magnifications have better optical quality than
those with zooms. For illumination, traditional focusable
light sources, halogen fibre optic, or light emitting diode
(LED) lights can be used. The traditional lights can often be
more precisely positioned and a greater working distance can
be achieved because the lights can be focused, unlike
standard fibre optic lights, allowing more freedom to
manipulate objects. On the other hand, fibre optics lights
have a higher light output and optional focus attachments are
available. LED lights are similar to fibre optics, though a
little weaker. They are particularly useful for field work due
to being lightweight and in having long-lasting bulbs. A
substage illuminator, a tiltable mirror, or a dark-field base
can be helpful when searching for radulae in maceration
solution (see below). The base of the microscope can be
mounted in a hole in the working platform (bench or desk),
so that the surface area of the microscope is level with the
remainder of the desk.
Pipettes
Pasteur pipettes of glass with a rubber bulb are useful;
the diameter of the tip can be adjusted by cutting. Disposable
Pasteur pipettes of polyethylene do not deteriorate and can
easily be cut to tip diameters of up to 5–6 mm. Pipettors
(Eppendorf and similar brands) used in molecular biology
are considered by some to be bulky and difficult to
manoeuvre when used under a microscope, while others like
their precise flow control. There are also devices that
produce a constant vacuum on a very small bore to pick up
specimens which are released when the vacuum is broken
(Hemleben et al. 1988).
For a pipette to form very small droplets (e.g., for
radular work), use a commercially available disposable
pipette tip for µl work; insert it in a fitting polyethylene tube
50–80 mm long and seal the other end (Fig. 4E). The
stiffness of the polyethylene tube gives better control over
the quantity delivered. A ‘home-made’ capillary glass-tube
can also be used as a tip.
Forceps
There are five main types of forceps used for work with
micromolluscs.
1.
2.
Entomology forceps. Made of thin spring steel, they are
available with a variety of tip designs, which can be
further adapted using a sharpening stone. Some users
find they have good handling properties with a reduced
risk of breaking fragile specimens while others find that
the tips do not meet exactly, or are too slippery. With
inexperienced users, specimens, especially smoothshelled gastropods, can be catapulted some distance.
Watchmaker’s forceps are available with very fine tips.
Many qualities and shapes of tips are available ranging
from the expensive straight MORIA MC-40 model
which has the finest tips currently available and is made
3.
4.
5.
of stainless steel. There are a wide range of similar,
cheaper models available. Watchmaker’s forceps are
most suited for anatomical and radular work, but with
practice they can also be used for routine sorting,
including handling fragile specimens.
So-called ‘Iris forceps’. These very soft forceps have a
rather broad tip (for example, the MORIA MC-32 or
32B has a tip of ca 0.8 mm), which can be either
smooth or serrated. These forceps are excellent for
handling specimens 5–10 mm and smaller. The shape of
the tip can be easily modified with a file or sharpening
stone. They are almost as soft as entomology forceps,
but less flimsy.
Stub handling forceps. There are two basic types for
handling SEM stubs; one made for handling Cambridge
stubs, by holding it in a track in the edge and another
designed for gripping the pin of all 1/8” pin stubs.
Grind the tips of the former so they become more
slender for a less tight fit in the groove and of the latter
to a finer point so they can be more easily inserted
under the stub. The Cambridge stub forceps can be
modified to have narrower and less curved tips.
Bamboo forceps (Fig. 4D). One of us (TS) makes
forceps from two pieces of bamboo. The tips of the
bamboo pieces are shaped with a knife and sand paper,
which can be accomplished in a relatively short time.
Bamboo is softer than steel and is suitable for
manipulation of fragile specimens and anatomical
manipulations.
Microscissors
Microscissors come in a variety of models and a wide
price range. Spring loaded scissors are suitable in many
instances, which can be complemented with a couple of
cheap, slightly larger ones for standard work. For particularly
delicate work, a pair of extra fine ones (e.g., MORIA extra
fine) can be useful. They come with various tip
configurations (pointed, blunt, angled) and are rather delicate
and expensive.
Scalpels
Scalpels with a fixed blade are not recommended as
they need re-sharpening, are expensive and corrode easily.
The common types with a flat metal handle and disposable
blades are suitable for most purposes. There are many
different blades available; microsurgery scalpels (e.g., FST
10315-12) are excellent for opening very small bivalves and
any other work where regular scalpels are too large. A cheap
and very good alternative is to use a needle holder to hold a
broken piece of razor blade (see below). Different brands of
razor blades break in different ways.
Pin and needle holders
These come in different sizes and materials, some
having a small chuck that will hold the finest needles. The
handles vary (diameter, shape and texture) to suit different
preferences and can be colour coded for easy identification
of the various needles. Dismantle and clean the needle holder
8
after it has been immersed in corrosive chemicals. Needles,
pins or razor blade fragments can be glued or otherwise
attached to tooth picks or other wooden sticks. Heated
needles can be pushed into thin wooden strips or perspex/
plexiglass rods, but the heating makes the metal more
sensitive to corrosion.
Surgical needle holders are useful for holding small
needles, pieces of a razor blade (or anything else thin or flat).
There are also special ‘blade holders’ available for this
purpose.
Pins and needles
Needles are probably the most important piece of
equipment for radular and many other micromollusc
applications. The finest and most expensive needles are
made from electrolytically etched tungsten wire (USD/Euro
5–10 each) with a 1 µm point (Fig. 4C). These can also be
made using tungsten wire and suitable equipment (Hubel
1957). Their points can easily be deformed, which is
sometimes an advantage for certain types of manipulation.
Micro-pins used to pin small insects come in black or
stainless steel and in diameters from 0.1 to 0.2 mm. The
stainless steel needles are less prone to rust and the thicker
ones are stiffer but less pointed, with much variation in the
shape and quality of the point. The black steel pins are
sensitive to rust (Fig. 4B) although their life is prolonged by
rinsing and drying after use. Household pins are much
blunter and thicker but are often made of chromium plated
brass and are less sensitive to chemicals. Sewing needles are
made of chromium plated steel, available in many sizes and
can be useful for work on larger specimens (Fig. 4A). Most
of the ready-made needles for surgical use are inferior to
micro-pins and much more expensive.
All metal needles or pins can be bent by holding the
very tip with a pair of watchmaker’s forceps and bending the
outermost fraction of a mm to a suitable angle. This tool is
useful for moving radulae from one rinse to the next (see
below). Such needles, including those with a minute hooklike end, are also valuable for dissecting. A needle with its
point bent at 45° is excellent for picking small single valves
of small bivalves; turn the shell so the concavity is up and
‘hook’ the valve under the hinge. Needles with a 60° bent
point are good for pulling the animal out of coiled shells. The
needle point is inserted along the wall of the shell, then the
point rotated to hook the animal.
Hairs
For cleaning dust particles from mounted radulae it is
often better to use hairs (rather than a pin) glued to a small
handle of wood or inserted into a holder. Eyebrow hairs (if
straight) and eyelashes are commonly used but some animal
hairs such as the pointed and stiff whiskers of a cat make
very good tools. The stiffness of hairs decreases with
increasing length of the hair.
Brushes
One centimetre wide brushes are useful for
manipulating dry samples while the finest brushes can be
used for manipulating individual specimens. The diameters
of brushes are graded, with ‘0000’ being the finest, and may
be made of either synthetic fibre or natural hair. Synthetic
fibre is chemically more resistant, can be used with bleach
and hydroxides used in radular extraction and are usually a
little stiffer. Natural hairs are often better for picking up
small shells but are more expensive. It may be necessary to
shape the tip, or to increase the stiffness of the brush by
shortening the hairs. Many fine paintbrushes have one or a
few much thicker and stiffer hairs to act as a support for the
others. These can be cut off to avoid the risk of specimen
damage if the brush is used for cleaning.
Pliers
The smallest sizes of regular tool pliers are useful for
cracking and opening small shells. Locking vice grip pliers
will prevent the specimens being crushed. For most
microshells, dissolving the shell is a better method (see
below). Wire cutters for electronic use come with a variety of
cutting edges, pointed, blunt, straight, angled, etc. and some
are made of stainless steel. These are good for opening
medium-sized (>5 mm) shells from the aperture, by breaking
the outer lip. For cutting steel needles, use wire cutters with
tungsten carbide edges. Watchmakers forceps can be used for
cracking very small shells (see below for details).
Drills
Some power tools resembling a dentist’s drill have a
flex-shaft attachment and a variety of rotary tool bits,
engraving cutters and high-speed cutters that can be used to
open shells with minimal damage. Drill bits are available
down to about 0.7 mm diameter and can be used for grinding
a hole of >0.7 mm diameter in the back of the shell. A hole
can be made in thin-shelled species simply by scratching the
shell with a needle.
Tool sharpening
A few different types of very small files for jewellery
work are useful, both for keeping other tools in shape and for
filing holes in 3–5 mm (or larger) shells. Files rust easily so,
if in contact with seawater, they need to be rinsed with hot
fresh-water and wiped dry.
For the rough shaping of coarser tools, a bench grinder
with as fine a wheel as possible can be used. For more
detailed work use fine sand- or carborundum paper or
sharpening stones. For the final sharpening of forceps and
needles, use a fine-grained stone such as ‘Arkansas Stone’
(see also http://www.antiquetools.com/sharp/sharphistory.
html). With some practice, a good point can be achieved for
watchmakers and some other forceps. Start with (if
necessary) roughly bending the tips so they are parallel, then
grind them off to the same length and start sharpening on a
fine carborundum stone. The final grinding is done on an
Arkansas stone, by moving it back and forth in the direction
of the points. The finishing should be done under a
stereomicroscope for better control.
Bowls
There are two types of bowls that we find particularly
useful for working on microscopic animals. Square, solid
glass bowls (‘embryo-bowls’), ca 40 x 40 x 15–17 mm are
ideal. Use a square piece of glass cut to the same size as a lid.
The sides of the ‘bowl’ slant at an angle, so nothing is
concealed by a meniscus. They are easy to handle since the
outer sides are straight and allow a good grip, as opposed to
watch glasses. The lid usually stops evaporation, but some
can have irregularities that prevent a tight closure.
Depression slides (concavity slides) are available with
different sized depressions. Those with a depression about
18 mm diameter and 2.5 mm depth are good for cleaning
smaller radulae. As a lid, another depression slide with a
larger diameter depression can be used upside down. If a
regular flat glass slide or coverslip is used as a lid when
heating KOH in radular preparation, condensation will form
on the glass of the lid above the fluid, which will finally
connect with the fluid in the depression and may draw it and
the radula into the capillary space between the slides. The
larger ‘dome’ above solves this problem.
Larger dishes useful for sorting are discussed below.
Collecting
Field collection of micromolluscs requires some specialised
techniques. Most micromolluscs are difficult to see with the
unaided eye and usually cannot be identified in the field
without magnification, making targeted collecting for a
particular species difficult. Thus the likely habitats of the
target organisms, or a range of microhabitats in the case of
surveys, usually need to be sampled.
The collecting methods covered below are simple
techniques that require minimal equipment and can be
undertaken by hand in intertidal and other shallow-water
aquatic systems. The process of obtaining small specimens is
not limited to intertidal and SCUBA as the same or similar
methods can be used on a larger scale as, for example, with
samples collected by various remote-sampling devices such
as dredges, trawls, epibenthic sledges or grabs. In such cases,
equipment (such as sieves and containers) needs to be
scaled-up. In order to obtain a representative collection, a
range of techniques should be employed.
The choice of the final volume of the sample, and the
number of samples, depends on the question to be answered.
Small samples of 50–100 ml will reveal the dominant species
whereas samples of 10 litres and more may still miss rare
species (rarefaction effect).
All collecting, domestic or foreign, should be
conducted under appropriate and applicable permits.
However, because microscopic species cannot usually be
collected in a targeted fashion and substrate sampling is
essential, this needs to be appropriately covered. Some
authorities do not provide the option for substrate collection
and require a priori list of species and numbers of specimens
to be collected. These issues are best dealt with on a case by
case basis.
9
Hand collecting methods
Shell grit. Sediments are usually distributed nonuniformly. Some areas accumulate organic material and
biogenic carbonates. These shell-rich portions of the
sediment are often referred to as shell grit or shell sand. They
can simply be scooped up and processed like any other
sediment samples. They usually have mainly empty shells
and sometimes can be the only source of certain species.
The fine sand can be removed from these samples in
situ by washing it in a sieve or moderately fine mesh bag
(Fig. 4F). While the bag method works well for the larger
micromolluscs, it will lose many of the smaller species.
Algal samples. Algae are a habitat of many molluscs,
those with large fronds usually having fewer (but often
different) individuals and taxa than the heavily branched or
foliose species, such as many of the turfing algae. Kelp
holdfasts can also harbour different species. Certain molluscs
are found only on particular algae; for instance, sacoglossans
are generally found on green algae (Chlorophyta). In (ant-)
arctic waters, larger algal species harbour many
micromolluscs, particularly in holdfasts. Large algae such as
Laminaria sp. may lose their blades seasonally, thus at most
a single season of micromolluscs can be encountered on the
blade, while the holdfast may contain several seasons’ worth
of fauna. Algae can be processed on site or collected for later
processing; larger algal species can be placed in a bucket,
smaller species may fit into a ‘zip-lock’ bag. The most
durable freezer zip lock bags with slider closure mechanism
are also suitable for SCUBA collecting. One litre of algal
volume for turfing species usually produces a representative
sample.
The method of extraction of the molluscs from algal
samples removed from the habitat depends in part on the
intended use of the specimens. They can be extracted alive
(using a binocular microscope) for observations on living
material or to extract specimens for special fixation.
For bulk collection, the simplest technique is to
vigorously wash algae on the shore in a bucket or bowl filled
with seawater. The algal material is then removed and the
sample allowed to settle briefly. The water can then be gently
decanted, being run through a sieve to catch any floating
molluscs (e.g., opisthobranchs). Such samples are ideal for
collecting living specimens for later examination. More
thorough washing can be carried out by vigorously shaking
algae in a 0.5–1 litre jar half filled with water from the
environment and with a tightly fitting lid. Samples may also
be pre-treated with an irritant or narcotic (see below) to
ensure that tenacious specimens are released from their
substrate.
Leaves and other litter. Rich organic material provides
molluscan habitats particularly in mangrove and other upper
littoral and supralittoral habitats as well as terrestrial
habitats. Mangrove litter can be washed as for algae and is
ideal for collecting e.g., small ellobiids, truncatellids and
assimineids.
Rock washing. Smooth rocks may be hand-washed
with bare hands in a bucket. Byssally attached bivalves,
limpets, chitons and opisthobranch may be tenacious and
10
require some assistance to dislodge. The upper and
undersides of rocks are very different environments; algal
films or turf usually cover the upper sides, while colonial
animals such as sponges, tunicates and bryozoans usually
live beneath the rocks along with their specialised
carnivores.
Sculptured rocks or large pieces of dead coral can be
scrubbed with a brush (e.g., 9 x 25 cm oval brushes or a
round brush about 5 cm in diameter are effective). When the
rocks are lifted out of the water, they can be brushed in a
bucket. The residue in the bucket, particularly from coral
washings, may harbour potentially dangerous animals so
care is required.
Rocks buried in sediment often have an anoxic,
blackish or rusty underside; certain molluscs occur almost
exclusively just at the oxic – anoxic border (e.g.,
phenacolepadids, some rissooideans, marine valvatoideans
and galeommatoideans).
Rock brushing while using SCUBA is best
accomplished within a cloth bag—a pillowcase is ideal—or
in a plastic laundry basket with a plankton net lining. The
rock is placed into the container and brushed within it from
above with the specimens mostly falling into the container—
although opinions differ as to how many specimens float off
rather than sink into the tub. Alternatively, the rocks can be
collected underwater and placed, with as little disturbance as
possible, into a large bucket or cloth bag attached to a buoyline. The container can then be hauled slowly to the surface
by the boat crew and the rocks can then be scrubbed as
described above, minimising the risk of losing specimens.
Other methods
Hasegawa (2004) and Hickman and Porter (2007)
recently reported the collection of samples of Scissurellidae
using floating light traps. The use of attractants (light, bait)
may be worth exploring, particularly for micro scavengers
and predators.
Small grab samplers (e.g., Petite-Ponar, Wildco, NY,
USA: www.wildco.com), have been used for the collection
of micromolluscs (Geiger 2006a). At 14 kg weight it is
transportable as luggage on commercial airplanes and can be
deployed and recovered by hand from a small boat by a
single person. Sampling beyond normal SCUBA depth to
220 m has been achieved (B. Raines, pers. comm.) and,
unlike a dredge or benthic sledge, it requires line only as long
as the sampling depth, and recovers even the smallest species. However, the sampling area is very small (15 x 15 cm).
Air-lift pumps can be used as a very effective way of
sampling both hard surfaces and substrate and are also a
means of obtaining, with careful and targeted use, large
quantities of living specimens (e.g., Bouchet et al. 2002).
Dredging and benthic sledge can provide significant
amounts of material and sample a larger area than either grab
or air-lift pump. The benthic sledge is advantageous as it
only skims the top surface where most micromolluscs are
found, but infaunal taxa will largely be missed.
Some taxa are commensals or parasites and their hosts
need to be examined—for example Eulimidae on and in
echinoderms, pyramidellids on other molluscs, Epitoniidae
on Actinaria, Aeolidioidea and Solenogastres on Hydrozoa,
and Doridoidea on sponges and bryozoans.
Methods of collecting terrestrial micromolluscs include
sorting leaf litter and soil samples, beating foliage and
carefully examining specific habitats—bark, rocks, crevices,
logs etc.
Narcotisation and relaxation
The procedures and concentrations for narcotising
animals vary greatly, except for magnesium salts, where an
isotonic solution (7.5% in freshwater) must be used in order
not to disturb the osmotic balance of the animals. It is
recommended that as few narcotising agents as possible
(including water, cold and heat) be used and users should
aim to get to know them well.
There are two main reasons for narcotising animals:
•
•
To facilitate and improve the yield of shake samples.
To relax animals for detailed studies.
The first is somewhat simpler, as it is acceptable if the
animal retracts into the shell or curls up. An irritant such as a
small quantity of formalin, a small amount of detergent, or
some freshwater for marine and estuarine species can be
added to the sample. Many molluscs will retract into their
shells but remain alive. Limpets and chitons may not
necessarily fall off unless such a method is employed, but
non-shelled molluscs may be adversely affected. If
specimens are intended for histology, non-isosmotic
treatment is best avoided. A secondary shake in water with
the irritant after an initial shake in habitat water may produce
additional species in a sample. Note that byssally attached or
cemented bivalves (e.g., Mytilidae, oysters) and some
limpets can usually not be reliably collected other than by
physically removing them.
The whole sample may be pre-treated with magnesium
chloride to anaesthetise the animals (75 g MgCl2 per 1 litre
of freshwater).
The algae may also be placed in a closed bag in full
sunlight so that heat stress will kill the animals, or for
tropical samples, cooling in the fridge or freezer will have
the same effect. In these cases, a single shake per subsample
will be sufficient to extract the vast majority of the
specimens.
Relaxation of animals for soft-part studies needs to be
more controlled and depends on the particular species in
question. The most common method is by gradual addition
of a 7.5% MgCl2 solution in freshwater to the holding
container. Various molluscs respond differently to
magnesium chloride; some will hardly be affected while
others may immediately retract into the shell. Gradual
addition of the narcotic produces the most satisfactory
results. Introduce the solution away from the animal and
gently stir the water. Wait a few minutes and watch how the
animal reacts. Once the extended animal has stopped
moving, wait a little longer, possibly add a little more salt
solution as an overdose and then gently touch the animal
with a brush. Watch carefully for even the slightest
movement (especially on the tentacles if a gastropod). Once
the animal has completely ceased to move, transfer it to the
fixative of choice.
The second most common narcotic is low temperature.
Place the specimens in the fridge and wait till the animals
have completely stopped any movement.
Sea slugs are often difficult to narcotise as they will
frequently autotomise cerata and evert their genitalia. Both
MgCl2 as well as low temperature work sometimes either
alone or combined, but with a significant failure rate.
Experimentation with other invertebrate narcotics such as
drop-wise addition of ethanol, sprinkling of menthol crystals,
diethyl ether, lithium salts, de-oxygenated (boiled) water,
carbon dioxide, tobacco, MS222 and various barbiturates
may prove advantageous (see Appendix 1).
Wet micromolluscs
Wet specimens for anatomical study may be stored in a
variety of preservatives, whereas for molecular work strong
(>95%) ethanol or freezing in liquid nitrogen at -190ºC are
the best preservatives. The particular fluid medium (e.g.,
water, ethanol, formalin, glutaraldehyde) has little effect on
the mechanics of the handling techniques. However, the
different media present various health and safety concerns
(see Appendix 1). Live sorting (following sieving in
seawater) enables observation of living specimens,
microphotography (Fig. 4) and/or the use of special
relaxation and/or fixation methods for individual taxa.
Sorting from bulk samples
Wet bulk samples containing significant amounts of
plant or algal material will turn acidic very quickly so require
extra buffering (see below) and should be sorted as quickly
as possible. Proper preparation of a sample can significantly
increase the efficiency of sorting. Preparation falls into two
main categories, separation by elutriation and sieving.
•
•
Elutriation—carefully floating off lighter matter such as
plant material and silt while the shelled molluscs
remain in the bottom of the container. As a more
sophisticated alternative, flush water with a hose or
pipe from the bottom end of a tall transparent cylinder
holding the sediment. The water flow, which must be
carefully regulated, will start carrying off all debris.
When the water flow is increased, initially soft animals
and light shells are carried away and collected in a sieve
and, finally, only mineral particles remain. Elutriation
and flotation always give better results if the size range
of the particles is narrow, e.g., 0.4–1 mm or 2–5 mm.
Sieving—necessary because it is easiest to sort samples
that contain significant quantities of sediment if the
particle size is homogeneous. Sieving through a series
of screens (see Tools section above) can achieve this.
The finest fraction, which may contain larval shells and
11
juveniles and occasionally very small-sized adults,
should be checked using a microscope before being
discarded.
All but the largest fractions of micromollusc samples
should be sorted under a stereomicroscope. Sorting
techniques vary considerably and we describe here a few
methods that have proven reliable. In general, when sorting
specimens, it is better to work with too small a subsample
than one that is too large. Dishes made of any material
chemically resistant to the medium are suitable, including
those made of plastic. Petri dishes (glass or plastic) are ideal;
lids of rectangular polystyrene boxes are even better since it
is easier to keep track of what has been sorted, particularly
those with slanting sides, where it is easier to both see and
grab specimens close to the side. As a high-end option, black
metal sorting trays, with or without rulings, used to sort
foraminiferans are available from a few suppliers (e.g.,
Green Geological Supplies: http://www.geocities.com/
greengeology). Even cheaper, small tartlet or pie pans are
available in specialty kitchen stores. The viewing
background should be in a contrasting colour, black being
suitable in most instances.
In one approach, a sorting dish is covered with a single
layer of particles. Round dishes make the even distribution of
particles easy, whereas square dishes are more easily
searched systematically. Swirling motion concentrates the
particles in the centre of the dish, whereas back and forth
motion moves material towards the periphery.
Alternatively, a small amount of either wet or dry
material is placed in the centre of a round glass Petri dish and
spread into an elongate pile approximately 5 cm long.
Starting at a face of the pile, spread small amounts at a time.
Techniques for picking up specimens depend on the
type of specimens and personal preference/experience. Three
types of forceps are commonly used: watchmaker’s forceps,
fine-tipped ‘soft’ stainless steel entomological goose-neck
forceps and iris forceps (see Tools section above).
Older fluid-stored specimens may become soft or brittle
requiring extra care. Forceps with deformed tips are useful
for smooth and slippery specimens. Very delicate specimens
can be sucked up with a pipette or, if dry, with a damp fine
brush. After transferring the specimen, check the wall of the
pipette to make sure that the specimen is not stuck inside.
Some like to use Irwin loops, particularly for live
sorting (K. Barwick pers. comm.), which have been used for
other meiofauna work (e.g., Kristensen and Funch 2000). A
small brush can also be used to move wet specimens in a
dish.
When working with ethanol-water mixtures, the
concentration in the sorting solution and the specimen vial
should be the same, otherwise turbulence will be induced by
the fluid from the other container adhering to the tool.
Similarly, small volumes of fixed samples can be examined
in 30% ethanol to reduce thermal circulation of the fluid, but
should be avoided if tissue swelling is of concern (see
subsection Storage below).
12
Fixation
The intended use of the specimens should determine the
fixation and storage fluid. For fixation and storage for
specialised needs we recommend the following:
•
•
•
Molecular work—95–100% ethanol. See also ‘boiling
method’ below.
Histology—formalin, bichromate or mercury-based
fixatives, Bouin’s fluid or other histological fixatives.
TEM—ideally glutaraldehyde fixation. Formalin can
also be used but with inferior results.
Detailed and complicated descriptions and recipes for
fixation and preservation are available but are mostly
unnecessary for standard work. Also, most methods work
well within a wide range of concentrations and often one
kind of buffer can be replaced by another as long as they do
not interfere. For example, recipes often specify that 3.7%
formalin is to be used. That is simply because they used
formalin : water, 1:9, but good fixation with formalin can be
achieved as long as it is stronger than ca 2%.
For more general information regarding fixation and
preservation see Gohar (1937), Romeis (1948, 1989),
Mahoney (1973), Presnell and Schreibman (1997) and
Glauert and Lewis (1998).
When fixing shelled molluscs, the fixative must have
access to the tissues; a light cracking of the shell is usually
needed, except in limpets, chitons and gastropods with a
short, broad spire, small operculum and large aperture. To
crack small specimens may be difficult without crushing
them. A small pair of wire cutters for electronics is usually
good; some models are made of stainless steel. Also, for
larger specimens, a bench vice, locking vice pliers, or any
other tool where you can control the cracking is better, to
avoid crushing the shell. Power pliers with an extra joint for
increased power are usually good for larger specimens with
thick shells. Watchmaker’s forceps can also be used like a
nut-cracker. Insert the specimen about a quarter of the length
of the handle from the join, with one face of the forceps on
the table, and gently press the other arm of the forceps until
the specimen cracks. However, this method requires practice
as it is liable to crush the shell unless carefully controlled.
More drastic measures (e.g., a small hammer) may break the
shell into many pieces and reduce the animal to pulp.
Drilling a hole in the back of the shell (see above) and
injecting 95–99% ethanol is an alternative for larger species
(>3–10 mm), but is not as safe as cracking.
For most studies involving micromolluscs, the shell is
one of the most important sources of taxonomic information.
For this reason, if shells are cracked or removed prior to
fixation, it is important to keep an undamaged specimen for
reference purposes—even an empty shell will often suffice.
Because micromolluscs often have little shell material,
they are particularly prone to adverse effects by preservation
fluids. Acidic formalin or ethanol can quickly damage or
completely destroy shells. However, formalin is a good
general fixative for tissue preservation and samples can be
used for TEM, SEM etc. Its biggest downsides are that the
material cannot be used for molecular studies with current
techniques and it is carcinogenic. Marine samples may be
fixed in 5–10% formalin-seawater, which is sufficiently
buffered for short time fixation (1 day) at a pH of
approximately 7 (Anonymous 2006a). It is important with
any fixative to have an appreciably larger volume (factor of
at least 5–10) of fixative than the specimen. For formalin
fixation, a quick approach is to fill a container with molluscs,
add 1/10 of the jar volume in 40% formalin and top off with
water (seawater is preferred as it provides a more nearly
isotonic solution). Mix the solution well by inverting the jar
several times until there are no more streaks in the fluid. The
bodies of the animals will provide the remainder of the water
to make an approximately 5% formalin solution. To properly
buffer formalin, 1 g of borax per litre seawater formalin
gives a pH of 7.5–8.5 and is good for several years.
However, borax may clear tissue during prolonged storage
(>10 years: Anonymous 2006a) and may be considered
unsuitable, although some of us have not noticed any
detrimental effects. Excess sodium bicarbonate mixed with
formalin (allow it to settle for several hours) made up with
fresh or seawater gives a pH of approximately 8. If instead
sodium carbonate is used, the pH becomes approximately 10,
which is much too basic, it becomes histolytic and the skin
peels off within a few years. Other buffering agents include
powdered aragonite, which is more soluble than calcite, but
may recrystallise and interfere with shell material, and
household ammonia, which reacts strongly exothermically
with formalin to form hexamine (= hexamethylenetetramine,
methenamine) to pH 8.2 (Clark 1998). Hexamine decays and
has to be adjusted after one and six months, and then every
two years (Hemleben et al. 1988). This labour-intensive
procedure will be prohibitive for larger collections (See
Appendix 1 for safety notes). Recently several ‘formalin
free’ fixatives and preservatives have appeared on the
market. Some are based on phenoxetol, which does not
replace fixation for histology or SEM purposes.
Storage
The most commonly used storage medium is 70–80%
ethanol. Borax, powdered aragonite, or powdered calcite/
shells may be added to ethanol solutions to safeguard against
shell damage. Precise amounts have not been specified,
though some have expressed concern that borax and
aragonite may pose problems with recrystallisation; this area
needs further investigation. To reduce the problem with
dissolution of shells in water-ethanol mixtures, the
concentration of the alcohol can be increased. For histology,
tissue shrinkage is of concern and it is customarily advised to
use a graduated series (30%, 50%, 60%, 70% ethanol) when
transferring specimens from aqueous to alcoholic solutions
to minimise shrinkage. Glauert and Lewis (1998) question
this approach, because <70% ethanol solutions expand tissue
and may damage fine structures. Hence, direct transfer from
formalin to 70% ethanol is appropriate. Freezing
micromolluscs in 70% ethanol in a -80°C ultracold freezer
for five years does not appear to affect the shells, but will
adversely affect tissues of un-fixed specimens.
13
The alcohol concentration can be kept stable in wellsealed containers. Specimens in tubes with polyethylene
(never polycarbonate, which will disintegrate) closures or
cotton plugs should be immersed closed end down in ethanol
in larger, secondary containers of proven durability. Because
ethanol vapours consist of approximately 95.5% ethanol,
evaporation reduces the alcohol concentration in the medium
and evaporated liquid should be replaced with 95.5%
ethanol. Calcium carbonate is quite soluble in water, hence
shells may easily be dissolved in an ethanol-water mixture.
For example, scissurellids can become fully decalcified in as
little as 18 months in ethanol at less than 80% concentration
(DLG, pers. observ.). As it is impractical to monitor alcohol
concentrations in small vials every six months, it is advisable
to store some shells dry as vouchers. Storage of
micromollusc samples with large amounts of organic
material should be avoided. Ideally use high quality ethanol
free of impurities, although this is much more expensive.
Long-term storage in formalin is generally avoided
since formalin is on the list of suspected carcinogens and is a
well-known allergene. However, it has been successfully
achieved in at least one major collection (AMS), where 5%
seawater formalin buffered with NaHCO3 is used. Problems
with ethanol include evaporation and flammability, with
collections requiring regular maintenance and special
fireproof housing. For any preservative, the pH needs to be
kept below pH 8.5 to avoid tissue dissolution and above pH 7
to prevent shells and other exoskeleton parts from dissolving.
Such narrow tolerances require regular testing and
adjustments. The pH of ethanol-water solutions is difficult to
measure, though some specialty pH electrodes are available.
For field storage, unbreakable plastic containers, ideally
with screw tops, should be used. Eppendorf tubes (1.5 ml)
and Falcon tubes (10 or 50 ml) seal fairly well, as long as the
seal is free of dirt. Heat-sealed bags can leak when filled with
wet sediment samples, but are useful as secondary
containers. Although cheap, scintillation vials should be
avoided, because ethanol evaporates quickly from them.
Scintillation tubes are also potentially dangerous during air
transportation, since they do not close well and are not
intended for such use.
Switching storage media
When switching specimens from one solution (e.g., 5%
formalin) to another (e.g., 70–80% ethanol), the tissue
volume and other water filled spaces need to be taken into
account, as water they contain will dilute the preservative.
As an example, a jar half-filled with specimens and filled up
with 95% ethanol will eventually result in about a 50–75%
ethanol solution. Hence, for samples intended for molecular
work, it is important to replace the solution with 95–100%
ethanol within the first two days.
Boiling method
Dr H. Fukuda has kindly provided details of a method
that he has successfully employed for microgastropods
which is a modification of a method used by a number of
Japanese malacologists for large species and is known as
‘niku-nuki’ (e.g., Habe and Kosuge 1967). It has proved to
be particularly useful for instances where only one or two
individuals are available and intact shells and animals are
required.
A living individual is placed in a small beaker in
enough water (seawater if a marine species) to enable it to
extend and crawl. Add hot (70–100ºC) water, which will
immediately kill the animal with the head-foot extended.
After a few seconds (1–2 for minute species) in the hot
water, move the specimen to a smaller dish of cool water
under a stereomicroscope. The animal can be carefully
removed from the shell by gently pulling on the head-foot
with forceps, holding the shell with a second pair of forceps
and rotating in opposite directions. The animal removal can
be facilitated by squirting water into the aperture using a fine
syringe. The water temperature and the length of immersion
in the hot water vary according to the size of the specimen
and the thickness of the shell, with larger, thick-shelled
species requiring hotter water and longer times. The visceral
mass (digestive gland and gonad) becomes hard and loses
flexibility in high temperature and sometimes cannot be
removed from the upper whorls of the shell. Thus, the water
needs to be hot enough to separate the columella muscle
from the shell and cool enough to keep the visceral coil
pliable. More details on this method will be provided
elsewhere (Fukuda, Haga and Tatara in prep.). As DNA is
not broken down in 80 to 100ºC, tissue can be placed in 99–
100% ethanol for molecular work (Ueshima 2002).
Dry micromollusc shells
Two ‘diseases’, or more correctly, chemical processes, that
ultimately result in the destruction of shells, have affected
many type specimens in museum collections including AMS,
BMNH, NMNZ, NSMT, NHMB, USNM and ZMO among
others (Fig. 5C). Some collections are more affected than
others, with no clear pattern emerging. The collections in
GNM, SMNH and ZMUC have largely escaped it,
presumably by using different types of glass. Typical
instances are illustrated by Higo et al. (2001) in
micromolluscs such as triphorids and turrids. Two different
kinds of ‘disease’ are recognised, Byne’s and glass, but it is
sometimes difficult to decide which particular ‘disease’ is
responsible (e.g., Kilburn 1996). The manifestation of both
diseases is identical in that they first produce white
efflorescence on the shell, which eventually crumbles to
dust, but they differ in the cause.
Species described prior to 1960, and many thereafter,
were illustrated without the benefit of the SEM. Much
needed detail for species level identification (e.g.,
protoconch microsculpture) cannot be observed using light
microscopy. Therefore, many species cannot be positively
identified from the original descriptions or illustrations and
type material is the sole recourse to settle uncertainties.
Accordingly, it is a high curatorial priority to upgrade storage
systems of micromollusc material, particularly types, and to
14
engage in effective damage control. Proper initial specimen
preparation can avoid many problems later on.
Initial drying
Prior to dry storage, all specimens should first be
washed in fresh water to remove dirt, soften mucus and
dissolve salts, which are hardened and/or precipitated by
ethanol. It is very important to remove any salt because
NaCl2 is hydroscopic. To remove excess water, to dehydrate
animal remains within the shells and to speed up drying, the
material may additionally be washed with 80–100% ethanol.
If the dry specimens or total samples were not previously
washed, soak the material thoroughly so that mucous, dirt
and salt crystals dissolve. Specimens can then be air dried on
absorbent paper.
Insufficient drying will negatively affect long-term
storage of the specimens. Specimens washed with 100%
ethanol dry fastest and with the least possibility of retaining
any liquid. In diluted ethanol, the ethanol will evaporate
faster and may leave water behind. This water may soften
gelatine capsules, may smudge labels and may contribute to
mould growth. In dry climates, air drying is adequate but in
more humid environments, a drying chamber set to 40–50°C
may be advantageous. Simple drying chambers can be
constructed using incandescent light bulbs in a simple box
with some openings to allow for air circulation. Blow-dry
systems are unsuitable because the specimens are too easily
blown away once dry.
Handling
Dry specimens may be kept in glass vials, gelatine
capsules, or cardboard slides, with or without cushioning
cotton. Specimens prepared for scanning electron
microscopy (SEM) may either be stored mounted on the
SEM stub (see below for storage conditions), or may be
dismounted (see below) and placed in a standard container.
To properly view specimens, they usually need to be
removed from their glass vial or gelatine capsule, whereas
the glass window of a cardboard slide usually allows
adequate viewing. Specimens are best placed onto a metal,
glass, or paper surface of contrasting colour. Avoid plastic as
the static charge can make it difficult to move dry specimens.
To move individual specimens into separate containers,
a moist (but not wet) artist’s brush is safest, but forceps can
also be used with care (see Tools section above). Dip the
brush into clean or, preferably, distilled water or ethanol and
squeeze the bristles between two fingers or touch a piece of
paper. Some people use saliva but, apart from hygiene issues,
it can leave residues on the shell that are obvious under the
SEM. The specimens will adhere to the tip of the moist
brush. Deposit the specimen in the new container on the
inner lip or wall of the container; rotating the axis of the
brush while keeping the specimen touching the container
helps to transfer the specimen from the brush to the
container. When transferring specimens into gelatine
capsules, make sure that the brush contains as little moisture
as possible (gelatine is soluble in water) as specimens may
become glued to the wall of the capsule.
A dry, soft artist’s brush may also be used for the most
fragile specimens, particularly if the brush is somewhat
frayed. Specimens can be gently ‘speared’ so that they are
held between the bristles of the brush.
Multiple specimens can be poured into the storage
container from the sorting tray. If a square dish is used, the
corner may be suitable to concentrate and spout the
specimens into a container. Alternatively, pour or brush the
specimens onto a tightly folded piece of paper, then pour
them into the container using the angulation as a guide.
Gently tapping the paper may help move stubborn
specimens. Other techniques include making a funnel out of
paper or using a small glass or metal funnel (avoid plastic).
Storage
Specimen containers
First, the specimens, even those collected as empty
shells, must be properly cleaned and free of salt (see above).
This can be done by soaking them in clean (preferably
distilled) water for a few hours and then drying thoroughly.
The dried specimens are best kept in small containers:
ideally gelatine capsules within larger high sodium glass
vials, or cardboard mounts made from acid free, archival
quality board. None of the storage media is superior for all
storage conditions; each has its advantages and
disadvantages. The following discussion of the materials
assumes that the specimens are in direct contact with that
material. In many collections, specimens are contained in
gelatine capsules or glass microvials and placed within glass
vials that hold the label.
Glass vials are the most durable solution, but are also
costly.
Gelatine capsules do not degrade shells, but should only
be used in conditions with < 60% average relative humidity.
Shorter periods of 60–80% relative humidity do not seem to
have a negative impact. Gelatine is hygroscopic, thus, in
conditions of high humidity, and with insufficiently dried
specimens, the gelatine softens and may glue the specimens
to the wall of the capsules. Usually, a gentle push with the
stiff bristle of a fine artist’s brush will free the specimen.
Otherwise, the gelatine can be fully dissolved in water.
Specimens do not seem to be damaged by sticking to
gelatine. Insects can also eat the gelatine capsules if they are
left loose, leaving holes for shells to fall out.
Cardboard slides, also called geology micromounts or
microfossil slides, are very space efficient. However, they
are usually made from acidic paper and may negatively
affect specimen preservation, particularly in humid areas.
They release nitric acid from the celluloid, which is known to
dissolve foraminiferans (Barbero and Toffoletto 1996).
Cardboard slides may be custom made using archival quality
material (see below). Another drawback is that sliding the
glass window usually generates a static charge and
specimens become stuck to the glass. Fragile specimens may
become damaged when the glass window is pulled through
the slit in the cardboard. For sources and types of microfossil
slides
see
http://www.pangeauk.com
and
http://
www.ukge.co.uk.
Chlorine bleaching (Clapp 1987) of natural cotton wool
can release acid and may negatively affect the specimens,
therefore, artificial cotton is more suitable. Some of us use
medical grade cotton wool with glass tubes and have not
observed any shell degradation. Avoid any polyvinylchloride
(PVC) products, as they release hydrochloric acid from the
plasticiser. Most micromolluscs do not have sufficient mass
to damage one another and cushioning is not necessary.
Small plastic boxes are sometimes used and some
consider them superior to gelatine capsules (Kurtz 2005). We
are cautious about plastics because of the plasticisers used in
the material and different types of plastic are not easily
distinguishable. Transparent Polystyrene boxes seem
acceptable. Mylar is the only clear plastic film known to us
to be of fully archival quality—it is also widely used in the
fine arts business.
Plastic foam has been and, to a certain extent, still is
today, popular with shell collectors to hold specimens in
place within a plastic box. Plastic foam is the worst possible
storage for any shells and is particularly insidious for
micromolluscs. All foam disintegrates in one way or another
over a relatively short period of time (10–20 years). At best
foam crumbles and specimens have to picked out of the
material but usually it partially liquefies and becomes sticky.
It can be rubbed off larger shells, but micromolluscs can
usually not be adequately cleaned. The plasticisers in the
foam or the foam matrix itself can also be acidic and cause
an efflorescence akin to Byne’s or glass disease.
Some shell collectors use commercial plastic clays,
often referred to as mineral micromount, or tackless picture
mountants. These are supposed to not release any grease,
although they eventually do, leaving wet-looking patches on
shells. Their long-term stability is quite variable. Over a 15
year period in a collection of approximately 2000 lots, some
80% of the material seemed to be unaffected by the plastic
clay, although long-term contact left residues in the grooves
and lamellae of finely sculptured specimens: 10% of the
material became tacky and stringy and some 10% crumbled.
The cause for the material’s decay is uncertain; oiled shells,
and those containing animal remains, seem to be more
frequently associated with the tacky or crumbled clay
(Geiger 2004). As many micromolluscs are finely sculptured
and as the clay may change dramatically over relatively short
periods of time, this mounting medium is unsuitable for
microspecimen storage (Geiger 2004).
Specimens are sometimes stored on SEM stubs, though
most shells can be removed from them without problems
(see below). Usually, recommended storage of SEM stubs is
in sealed containers with silica gel to remove moisture and, if
possible, in an evacuated bell jar. These measures ensure that
the specimens will not outgas when placed in the high
vacuum of the SEM chamber, particularly those with field
emission and LaB6 guns, which require a vacuum at least an
order of magnitude greater than for tungsten filaments.
Hence, these storage requirements rather address operational
issues of the SEM and do not relate to specimen
15
preservation. Whether such storage conditions actually make
a difference will depend on local environmental conditions
and on the particular SEM used. The re-emergence of
tungsten guns, especially on variable pressure SEMs, makes
the above storage requirements unnecessary; protection from
dust and storage in a normal collection environment is
sufficient.
Labels
Ideally, the full-data specimen label should not be in
direct contact with the specimens, but contained in a
secondary container (usually a glass vial) housing the
gelatine capsule or glass microvial housing the specimen(s)
or on the backside of a geology micromount (as in the
LACM). In some collections, a tiny label showing only the
registration number is added to the specimen vial. Such
labels offer added protection against switching data between
lots. However, even small paper labels in direct contact with
microspecimens should be made from acid-free paper. We
are unaware of any issues relating to writing medium; pencil,
pigment ink and laser writer labels; all seem to work well.
There is anecdotal evidence that photocopied labels are more
durable than that of laser writers because the temperature at
which the toner is fused with the paper is higher in
photocopiers than in laser writers (H. Chaney, pers. comm.).
Heating labels in a direct heat oven improves the durability
of laser print; the toner changes from a powder-like
appearance to a shiny surface when examined under a
microscope; ‘cooking’ labels with microwaves does not
affect the durability of laser labels (Zala et al. 2005).
Elevated humidity has an adverse effect on laser printing,
because the temperature is lowered by the residual water in
the paper. As a safety measure, the catalogue number can be
written in pencil on the back of the label.
Paper as a material should be carefully considered in all
applications: labels, geology mounts, cardboard boxes. The
term ‘archival’ can be misleading, as a number of paper
products of variable long-term stability are issued with such
a descriptor (Clapp 1987; Turner 1998). Acid free paper
products are made from 100% cotton rags, not from wood
pulp, are usually not bleached and will not release any acids.
Buffered or pH-balanced products are made from wood pulp
and contain a pH-buffering substance, usually calcium
carbonate powder. The wood pulp will continuously release
acids, which at first is neutralised by the calcium carbonate
in the paper, but eventually the buffer capacity is exhausted
and acids will be released from the paper product. Synthetic/
plastic papers have not been available for a sufficiently long
period of time in order to assess their suitability as data
labels. A good start for internet searches is http://
www.universityproducts.com, http://www.archivalsuppliers.
com, http://www.archivescanada.ca/english/index.html and
http://library.amnh.org/conservation/suppliers.html.
The strongest paper we have found is ‘laundry tag
manila’. Coated papers in particular are unsuitable because
of the chemical coatings and filler materials, which will
eventually deteriorate.
16
Chemical deterioration
Byne’s ‘disease’
Micromolluscs are often fragile and prone to adverse
reaction with acids and salts. A serious destructive effect on
calcium carbonate shells is known as ‘Byne’s disease’,
named for L. St. G. Byne (1872–1947) a British amateur
shell collector who first described the phenomenon (Byne
1899a, b). Its manifestation is a white efflorescence covering
the shell, which eventually destroys the specimen
completely. Byne’s explanation that it was caused by butyric
and acetic acid was partly correct. It seems not to be caused
by shells with remaining animal tissue since the old
collections where it occurs often contain empty shells only
(pers. obs. by the authors). Tennent and Baird (1985)
identified the crystalline substance as a mixture of calcium
acetate and formiate and considered that the acetic and
formic acid originated from the oak-wood frequently used in
cabinets. The reaction can also be accelerated by low air
circulation and by high humidity, where the water molecules
act as an extractor and carrier for the acids. The necessity for
well aerated cabinets was pointed out as a precaution by
Byne (1899a, b). Some other organic substances are also
likely culprits and include cork, natural cotton, fibre-wood or
particle board, where formaldehyde-based glues are used, as
well as any surface treatment evaporating formalin as
applied for instance to some metal cabinets. All of these
substances should be avoided. A number of articles have
been written on various aspects of Byne’s disease (Kenyon
1909; Lamy 1933; Nicholls 1934; Nockert and Wadsten
1978; Padfield et al. 1982; Grosjean and Fung 1984;
Hatchfield and Carpenter 1985; Kolff 1988; Kamath et al.
1985; Davies 1987; Davies 1988; Hertz 1990; Pinto de
Oliveria and de Cassia da Silveira e Sa 1996; Stamol 1998;
Callomon 2000, 2003; de Prins 2005).
Glass ‘disease’
As in Byne’s disease, the so called ‘glass disease’
produces an efflorescence of calcium salts and, unless halted,
will lead to complete destruction of specimens in as little as
5–10 years (Fig. 5). It starts with the surface of previously
shiny specimens becoming dull, then powdery, then crystals
start forming. The shells start disintegrating and finally
crumble with only a whitish crystalline powder remaining. It
can occur in specimens that were originally perfectly dry and
repeatedly washed in fresh water and ethanol, including fatsolvents such as perchlorethylene and carbon tetrachloride,
stored in metal cabinets in tubes with plastic closures (no
cork or cotton) and acid free archival labels. The glass
disease has hardly been mentioned in the literature (except
Kilburn 1996). Glass can release sodium hydroxide when
interacting with moisture in the air and this NaOH and any
other leaching minerals interact with the calcium carbonate
of the shell leading to the formation of sodium carbonate
powder (see Birch 2000).
Our observations indicate that the worst offenders are
high quality, high silica, heat-resistant glasses, especially cut
sections of narrow bore tubing, although the extent to which
specimen preparation and environmental parameters play a
role in development of glass disease is an open question.
Cheap, high sodium carbonate glasses, such as disposable
culture tubes or blood test tubes are by far the better choice.
Although glass disease can be arrested in tubes containing
moisture absorbent silica gel sealed with plastic closures, by
far the simplest remedy is to avoid contact with glass by the
use of gelatine capsules if environmental conditions are
appropriate (see also above).
FIGURE 5. A. Six specimens of Skeneopsis planorbis (Fabricius,
1780) from the same lot and in the same vial from SMNH. The four
peripheral specimens are glued to paper, so the specimens were not
in direct contact with glass. The two central specimens are free,
they were in contact with the glass vial and show the white
efflorescence produced by the glass disease. Photograph AW. B.
SEM micrograph of shell affected by glass disease. Scale bar = 500
µm. Micrograph AW. C. Last surviving syntype of Anatoma
aedonia (Watson, 1886) (BMNH 1887.2.9.398); the remainder had
crumbled to dust. Uncoated specimen in variable pressure SEM.
Scale bar = 1 mm. Micrograph DLG. D. Detail of efflorescence
from specimen shown in B. Scale bar = 30 µm. Images AW.
Preparation of micromollusc shells for SEM
Cleaning
Whenever possible, select the cleanest shells available
for SEM. Delicate specimens can be cleaned with a fine-
tipped artist’s brush. Two main methods are outlined below
that can be used to remove stubborn dirt.
Cleaning with bleach. Specimens can be immersed for
one to two minutes in strong commercial bleach and/or 10–
20% hydrogen peroxide in order to remove the periostracum
or to remove tissue remnants from the inside surface of
bivalves. Excessive treatment for more than a few days
should be avoided, because minute crystals cover the shell
surface and become difficult to remove. Commercial bleach
contains meta-silicates that are added to stabilise dirt in
suspension so it does not precipitate on the surfaces
supposed to be cleaned. These silicates may precipitate in an
irreversible reaction when water evaporates and the pH
changes as a result of formation of NaOH, which then reacts
with air to produce sodium carbonate. Shells must be washed
in water repeatedly, especially in spirally coiled gastropods
in which bleach is trapped within the inside of whorls. As
these treatments dissolve tissue and tanned proteins, the
conchiolin matrix within the shell will also be weakened,
rendering the shells more brittle, although breakage rarely
occurs. More diluted bleach or hydrogen peroxide is less
destructive and may save the periostracum and nacre, so start
with weak solutions until you gain experience of your kind
of material.
Ultrasonic cleaning. Sturdy specimens can be cleaned
using ultrasound. Heavily incrusted dry specimens should be
soaked first overnight or even longer to loosen the dirt.
Additional soaking steps between sonication treatments may
be necessary. Cleaning of dry or wet stored shells is best
accomplished in a mild detergent solution; a few drops of
dishwashing liquid or a pinch of trisodium phosphate powder
in 100 ml of water. The detergent should be neutral or
slightly alkaline, some are too acidic. Shells can be first
wetted with ethanol to break the surface tension, which helps
to avoid trapped air bubbles, and then immersed in the
cleaning solution in a glass specimen tube, embryo bowl, or
watch glass and left to soak for a few hours or overnight. The
container is then put in the water bath of an ultrasonic
cleaner. The duration of sonication depends both on the
power of the sonicator, the water level in the bath and the
specimen itself. In some instances, even one second
sonication can break a delicate specimen, yet with some
specimens 60 second sonications are harmless and necessary.
Because thin-shelled specimens may break during
sonication, particularly if air bubbles are lodged within the
shell, the sonicator should be tested using similar,
expendable specimens. Air bubbles can be removed by
putting the submerged specimen under moderate vacuum but
avoid the water boiling (boiling point of water ~0.01 bar at
25°C). Some consider the use of a sonicator too
unpredictable and too often damaging to the specimens.
Thus, it is advisable to not use ultrasonic cleaning on rare or
unique specimens.
After sonication, shells should be washed in distilled
water—preferably two or three times. It is safer to change the
fluid in the glass container than to handle the specimen. After
the last wash, remove the larger water drops with a paper
tissue or paper towel and air-dry the specimen. Alternatively,
17
the specimens can be stored in ethanol and then placed on
blotting paper just prior to mounting; the remainder of the
ethanol will quickly evaporate. The dried specimens are now
ready for mounting.
Mounting
Multiple specimens can be mounted on one stub but it
is imperative to make a ‘stub map’ to keep track of the
specimens. A sculptured orientation marker (e.g., a dab of
colloidal graphite, some silver paint or a groove in a double
sided carbon tab, that is easily visible in the SEM) is needed
because coating will obscure pencil or pen marks. Different
sets can be separated by marks and/or numbered separately.
It is advantageous to mount specimens that may be confused
mixed with easily identified ones, which serve as landmarks.
Mounting of specimens can be achieved in a variety of
ways. Ideally, multiple standardised views should be
obtained with minimal remounting, because any handling of
specimens increases the risk of damage or loss. Typical
standardised views for coiled gastropods are apertural,
apical, umbilical. Mounting the specimen on the periphery of
the last whorl opposite the aperture serves this purpose with
a typical SEM stage that usually allows approximately 90°
unidirectional tilt and 360° rotation. Unfortunately, it is also
one of the least stable orientations. For bivalves, interior and
exterior views, with the shell outline in the image plane and,
possibly, enlargements of the hinge and an umbonal view
showing the prodissoconch, are typically obtained. Most of
us remount specimens at least once to obtain all views
necessary from the same specimen.
At least four main mounting media are available
depending on the object being mounted and each has its
advantages and disadvantages. Consideration of the
electrical conductivity of the mounting medium is essential
to reduce or eliminate ‘charging’ (see below).
Colloidal graphite. A small dot is applied to the stub. If
a specimen is placed into fresh colloidal graphite, capillary
forces will pull the mounting medium into the sculpture of
the shell. In order to avoid this problem, either use a more
viscous suspension of colloidal graphite, or wait for the
surface of the colloidal graphite to become silvery. The
specimen can then be placed with a moist artist’s brush and
held until the medium has sufficiently dried to hold the
specimen. Colloidal graphite is mostly used with relatively
large and heavy specimens, which are not sufficiently held in
place by double-sided carbon tape or stickers (see below).
Silver paste. This is normally more viscous and avoids
the problems associated with capillary forces. However,
silver paste also dries more slowly, making secure
orientation of the specimen more difficult. It is mostly used
for larger specimens to paint conductive wires on specimens
(see below).
Double-sided tape and double-sided carbon stickers.
These are less adhesive than colloidal graphite and silver
paste but can be used to mount smaller (< 2 mm) coiled
gastropods on the periphery of the last whorl opposite the
aperture. For other, more stable specimen positions, there is
effectively no size limit. Carbon tabs can be shaped with a
18
blunt-ended tool to make a ridge or a mound against which
the specimen can rest, or can be deposited in such a fashion
that the material will form waves or wrinkles. These
sculptural elements of the mounting medium offer additional
bonding surface to the shell (and opportunities for specimen
identification; Fig. 6). However, part of the specimen will be
obscured and 90° tilts of the SEM stage show less of the
apical and basal surface, making re-mounting necessary if all
views need to be achieved with a single specimen. The
mounting with carbon tabs is somewhat flexible; after an
initial placement the orientation can be adjusted a little by
pushing the specimen gently in the desired direction.
FIGURE 6. A gold coated atlantid heteropod mounted for SEM.
Due to the keel of the atlantid shell, it can not be mounted on its
periphery. The double sided carbon tab material was shaped into a
mound with old forceps and the specimen leaned against the
material. The aperture was put horizontally by a combination of
stage rotation and tilt, and the image of the specimen was kept
horizontal using digital image rotation; note the slope of the stub
from lower left to upper right corner. For publication, the specimen
can easily be cut out from the background. SEM operated in
variable pressure mode at 30 Pa, 20 kV, 200 pA, at 10 mm working
distance. Specimen courtesy Roger Seapy. Scale bar = 1 mm. Image
by DLG.
Carbon stickers and tape are supposed to be electrical
conductors, but some brands are only conductive along the
surface, not through the cross section of the material (R.
Burns, pers. comm.). Some, such as NEM tape (Nisshin EM
Co. Ltd., Tokyo) is conductive through its cross section.
Clear, double sided office tape can also be utilised, although
it is not conductive. Other double-sided tapes (copper,
aluminium, nickel) available from SEM supply vendors offer
various adhesive properties with which one can experiment.
Glues. Non-permanent spray glue (e.g., 3M™) is
possibly the most flexible mounting medium. The spray is
designed for temporary attachment and remains plastic for a
prolonged time. It is applied by spraying a thin layer of glue
onto the stub. The thickness of the layer can be varied
depending on the specimens. The glue surface is less even
than that of the carbon tabs. While it is most useful for
specimens of more than 1 mm in size, it can be used even for
larval shells. Attempts to use spray glue with 1 mm
specimens by placing them on top of glue-covered pin heads
were unsatisfactory because the surface of the glue was too
sculptured. The spray glue is stable in high vacuum (10-4 bar)
and in the electron beam. The major advantage of the spray
glue is its flexibility, even after coating. The glue itself is
non-conductive and charging can be more pronounced than
with carbon tabs. Pre-sputter coating the glue makes the
surface non-sticky. Letting the glue dry overnight, or putting
it under vacuum, will make the glue more viscous prior to
mounting. This can be advantageous with very fragile
specimens as they will adhere with less surface to the glue.
For large specimens, the glue can be shaped into mounds as
detailed above for carbon stickers.
Polyvinyl acetate based (e.g. Elmer’s) glue is good for
mounting radulae and opercula, but should be avoided for
shells since its acidity will quickly (within hours) corrode the
shell. It is, however, suitable for grounding CPD specimens
and can easily be drawn out to form small conductors.
Preventing charging
Stub mounted specimens undercut all around relative to
the stub (e.g., a gastropod mounted on its periphery, or a
bivalve valve mounted concave side uppermost) will not
receive conductive metal coating on the side in ‘shadow’ and
will commonly ‘charge’ under the SEM. Once mounted,
such specimens can be coated by tilting the stub at a suitable
angle or may additionally require careful painting of a ‘wire’
of conductive material between the stub and the periphery of
the specimen. This is most easily achieved by placing a small
blob of carbon paint on a narrow wedge of paper and
inserting it between the specimen and the stub, being careful
to ensure that the material does not run up onto the surface to
be viewed.
Some of us consider the use of conductive glue such as
colloidal graphite or silver paste more hazardous than worth
while, because of the difficulty in applying the material at
small scales and capillary forces which can pull the glue over
the surface to be viewed. The often tricky painting of a
conductive wire onto the shell can be avoided with more
modern SEMs, low accelerating voltage (0.5–2 kV), variable
pressure operation, reduced probe current/spot size and using
frame integration as opposed to line integration as noise
reduction technique for imaging. Specimens may also be
sputtered multiple times for short periods of time at different
angles to coat the under surface, or a sputter coater with a
slanted and rotating specimen holder can be employed (e.g.,
Cressington 108SE with rotary/planetary/tilting sample
stage; Quorum Technologies SC7640 with RotaCota stage
RC7606). The thickness of the metal coating is 1–10 nm and
even excessive coating will not interfere with the detail in
normal shell and radular work.
SEM parameters
Most SEMs allow a multitude of imaging techniques,
with modern designs adding additional features. We
encourage the users to explore the parameter space to obtain
the best images possible. Figure 7 shows some options
applied to a rather difficult specimen. Charging effects are
accentuated, because the specimen is partly corroded, which
by itself usually makes charging worse. Additionally,
because the specimen is corroded, it is also more fragile and,
consequently, could not be thoroughly cleaned without
risking breakage. The remaining dirt also increases charging
19
problems. Last but not least, the specimen is globular with a
single, small attachment point to the stub. Three different
detectors were utilized: the usual secondary electron detector
(Fig. 7A,C) and the backscatter detector (Fig. 7B–D) both in
high vacuum and, in a variable pressure environment, the
variable pressure secondary electron detector. Figure 7C
shows the effect of signal mixing, where the signal from two
backscatter detector quadrants is used to balance the
directional lighting effect of the secondary electron detector.
Probe current can also have a significant effect on charging
(Fig. 7E: 100 pA. Fig. 7F: 200 pA).
SEM imaging of uncoated material is usually carried
out in either low voltage (<1 kV) or in variable pressure
mode in which air or dry nitrogen is introduced into the
specimen chamber. Preliminary experiments using
environmental SEM technology, in which water vapours
instead of air is introduced into the chamber, shows promise
with particularly difficult specimens (DG, pers. observ.).
SEM parameters may be optimised differently
depending on the type of SEM used (tungsten vs. field
emission gun, high vacuum vs. variable pressure), detector
type and arrangement (secondary vs. back-scatter detector,
side-mounted vs. semi-in-lens design of secondary electron
detector) and the particular model of the instrument.
Goldstein et al. (1992) provided a thorough general
introduction and Reimer (1998) explained the underlying
physics. Tungsten guns in general allow a quicker specimen
change as the vacuum requirements are not as stringent as for
field emission (and LaB6) instruments; variable pressure
instrument are usually fitted with a tungsten gun. Field
emission guns in contrast emit more electrons, and therefore
have better signal to noise ratio, particularly at higher
magnification (>10’000 x). As the SEM parameter-space is
multidimensional, changes in one parameter show various
effects depending on the sample and the particular
instrument used. Additional constraints may be imposed by
detectors other than the usual secondary electron detector
used, as many require a 15–20 kV minimum accelerating
voltage. It is beyond the scope of this contribution to provide
optimum conditions for all specimens and all instruments;
the investigator is encouraged to explore the parameter
space, or to provide some indication to an operator not
familiar with the type of specimen. Table 1 gives some
indications on the various factors and some of the common
effects.
While most of the effects do not require further
explanation (resolution, charging, depth of field), sample
penetration requires consideration. Secondary electrons are
generated due to the interaction of the electrons of the
electron beam with electrons from the electron shells in the
atoms of the specimen. This interaction does not only occur
at the surface of the specimen, but with increasing
accelerating voltage, the penetration depth increases. Thin
materials such as thin opercula and radular teeth may be
completely penetrated by the electron beam. The materials
appear translucent and occasionally, the irregularity of the
underlying mounting medium may be visible. A reduced
accelerating voltage will remedy the problem, at higher
magnification (>~3000x) at the expense of resolution.
TABLE 1. Effect of changing SEM parameter. Only the most
common factors are listed and only the major effects are noted.
SEM
parameter
Accelerating
voltage
Spot size/
probe current
Working
distance
Chamber
pressure
high value
low value
higher resolution
lower resolution
more charging
less charging
more sample
less sample
penetration
penetration
narrower field of view wider field of view
more signal
less signal
lower resolution
more depth of field
higher resolution
less depth of field
less resolution
wider field of view
less charging
more resolution
narrower field of
view
more charging
less resolution
more resolution
Note that the commonly cited parameter of
“magnification” depends on the size of the output (35 mm
and 4x5 inches as common settings in SEM preferences),
whereas the field of view (e.g., 100 µm) provides a constant
reference point. A field of view of 100 µm is magnified 350x
on 35 mm, but 1250x on 4x5 inches. Given the wide variety
of instrument types and possible operating conditions, the
familiar magnification ranges used here are adequate.
A special technique to enhance surface sculpture is
demonstrated in Figure 8. Most SEM images are taken using
the secondary electrons. The usual side mounted secondary
electron detector (in contrast to semi-in-lens designs) is
usually at an angle of approximately 45°, producing an
apparent illumination angle of 45°. Specimens with very
subtle surface sculpture may not show it sufficiently. In
photography, flat lighting just grazing the surface of the
specimen may be employed, however, the vertical position of
the SEM detectors is fixed.
The electron beam—specimen interaction produces a
number of different electrons, the two most important ones
being the secondary and the backscatter electrons. The two
differ in their electron energy; secondary electrons have up to
-50 V, whereas backscatter electrons have between -50 V and
up to slightly less than the accelerating voltage (usually 10,000–20,000 V). The more energetic backscatter electrons
produce straighter trajectories than the low energy secondary
detectors. As a result, slight surface irregularities can be
better visualized with the higher energy backscatter
electrons. As the regular backscatter electron detector is
usually mounted near-coaxial with the electron beam and
parallel to the specimen, it does not aid in this situation.
20
21
FIGURE 7. Illustration of different SEM imaging techniques applied to the same specimen sputter coated with gold. The shell is somewhat
eroded, which often leads to greater problems with charging. As eroded specimens are often also more fragile, the specimen could only be
superficially cleaned, with the remaining debris further enhancing charging problems. All images taken with Zeiss EVO40XVP at 20 kV,
10 mm working distance, and 100 pA probe current (except F). A–D. High vacuum; integration of 25 frames, total imaging time 1.5
minutes; line integration produced heavy charging artefacts (not shown). A. 100% secondary electron detector. Notice uneven illumination
in upper right portion of shell. B. Two lower left-hand quadrants of backscatter detector. C. Signal mixing of 75% secondary electron
detector from A and 25% backscatter detector from B. Notice the more even overall illumination as the backscatter signal is used to
brighten up the dark portion of the shell. D. 100% backscatter detector with all four quadrants active. E, F. Variable pressure mode with
chamber pressure at 30 Pa, line integration for 1.5 minutes. E. Variable pressure secondary electron detector (VPSE) with 100 pA probe
current. F. VPSE with 200 pA probe current. Notice greater charging effects at suture. Anatoma proxima (Dall, 1927) (USNM 449418),
Scale bar = 1 mm. Images DLG.
The secondary electron detector attracts the negatively
charged secondary electrons with a bias voltage of around
+300 V (Fig. 8A). If the bias voltage is set to –50 V (Fig.
8C), the secondary electrons are repelled, while the highenergy backscatter electrons with straight trajectories can
overcome the slight bias barrier. With more negative bias
voltage the effect can be further enhanced (Fig. 8D). As the
electron yield is smaller, a higher probe current is usually
necessary. The contribution of secondary and backscatter
electrons can be varied continuously, by adjusting the bias
voltage between 0 and –50 V.
Specimen removal from stubs
Specimens can be removed from carbon tabs and
double-sided tape in dry condition if necessary, but the bond,
particularly between carbon tabs and shells, is rather strong
and specimens may break when attempting to remove them
from the tab. This tendency becomes more pronounced once
the carbon tab has been exposed to high vacuum. Specimens
can be removed from the carbon tabs using 95–100%
ethanol, either to remount specimens for additional views, or
to be returned to the specimen vial. Cleaning colloidal
graphite from a shell requires multiple washes in isopropanol
or 80% ethanol. Uncoated carbon tabs can be reused a few
times (e.g., when imaging type specimens). Repeated exposure to ethanol makes the glue less sticky. Specimens can be
remounted on coated tabs if the mounting spot is rubbed with
a blunt pin to remove the gold coat on its surface.
Spray-glued specimens can easily be removed from the
stub and returned to the original lot. The glue can be
dissolved in butyl acetate or acetone (neither of which are
very toxic nor do they evaporate too quickly), or chloroform.
These solvents are also used for cleaning used stubs.
Separating the valves of minute bivalves
Small wet or dry bivalves may be difficult to open since
they are firmly stuck together. Avoid trying to open these
when dry. The specimens, with or without the animal, should
be initially passed through ethanol to break surface tension,
then soaked for several hours or even a day or two in water
containing a little detergent. Some additional techniques are
outlined below, the choice depending on the degree of
overlap at the valve margins and whether or not a little
damage to the ventral margin of the valves can be accepted.
•
•
•
•
•
Do in a vacuum, where enclosed air may exert
sufficient pressure to push the valves open slightly.
If there is little or no valve overlap, view under a stereomicroscope (and moisten periodically), while placing
the specimen with one end against the thumb and the
other against the forefinger (right handers) of the left
hand, with the anterior or posterior end uppermost and
the ventral margin facing right. In this way the valves
may separate slightly, making it easier to insert the
point of a scalpel or mounted razor blade fragment (a
blade spreads forces over a wide area, unlike needles or
forceps which are more liable to break or damage the
valves). As soon as the tip is inserted, the bivalve can
be held with the blade and cautiously pushed with its
back against the finger and, if a preserved specimen, the
adductor muscles cut.
Cutting the adductor muscles can also be achieved by
carefully moving the specimen towards a fresh blade
held vertically between the thumb and two forefingers
of the right hand, the objective being to place the
ventral meeting point of the valve margins squarely
against the blade.
Soaked specimens can be placed in a specimen tube
half filled with water and connected by a tube passing
through a closure to a standard, water-driven,
laboratory vacuum pump, repeatedly applying vacuum
via a valve or hose clamp, with the aim of parting the
valves sufficiently for them to be opened with a blade.
Very small or very fragile bivalves can be soaked in
warm, diluted bleach so that a bubble of chlorine will
push the valves open. Use a paint brush (with artificial
hair!) or a fine pipette for handling the specimens. This
method is unsuitable for highly nacreous shells, which
are sensitive to bleach. Instead use weak hydrogen
peroxide with a trace of KOH added to make it basic.
Note that sodium lauryl sulphate is not suitable as it
takes longer and does not open the valves, it only
dissolves the soft tissues, not the ligament composed of
tanned proteins.
SEM imaging
Specimens should be illustrated in standardised views.
For gastropods, the coiling axis of the shell should be parallel
or at a right angle to the image plane. In apertural view,
showing a little of the outside of the outer lip allows better
22
determination of the position of the shell, facilitating
comparison. Protoconchs should be shown in apical view, at
right angles to the coiling axis or parallel to the coiling axis
with the transition proto- to teleoconch in the centre.
Bivalves should be shown with the outline of the shell
parallel to the image plane and the prodissoconch (with at
least long axis parallel to image plane) should be shown in
umbonal view.
SEM preparation of animals
External anatomy can provide many useful features. The
animal should be preserved in as natural a state as possible,
ideally following relaxation. In general, formalin or
glutaraldehyde fixed animals dry better than those fixed in
ethanol only. Additional post-fixation may be carried out
with osmium tetroxide (OsO4), particularly if specimens are
viewed in older SEMs that necessitate high accelerating
voltages as OsO4 will add conductivity to the specimen (see
Appendix 1). Post-fixation is not necessary for more modern,
low voltage or variable pressure/environmental SEMs.
Whether low voltage operation (< 1 kV), possibly with
increased spot size, or variable pressure/environmental mode
gives better results depends on the particular model of SEM
and the particular specimen and it is worthwhile
experimenting with various settings (see also above). Much
information can be obtained (Fig. 9), even using older SEMs
and crudely preserved (‘vodka’) specimens, with the results
usually surpassing visual examination under a
stereomicroscope.
Preliminary inspection
Wet specimens preserved in glass tubes do not need to
be removed from the tube for a quick assessment. The glass
tube can be immersed completely in the same preservative
and the distortion caused by the glass will disappear. An air
bubble in the container will also greatly reduce the distortion.
For quick approaches suitable for common and
abundant species, see ‘Removing the shell the fast way’. If
the specimen is rare or unique as much information as
possible should be obtained from it, including SEM of the
shell, information on external animal morphology etc.,
before removing the radula or other relevant internal
structures. The following techniques require some
experience and practice with common species is
recommended.
It is generally easier to extract bodies from dried
gastropod and bivalve specimens that have been rehydrated
as opposed to continuously fluid preserved specimens. In the
latter, the body is more firmly attached to the shell and it may
even be useful to dry the specimen and then soak it if other
methods are unsuccessful.
Limpets
Bodies can usually be separated from the shell, or dried
in the shell (see below: Tissue preparation for SEM), which
will protect the body from any damage when physically
removing the body from the shell. The shell with body can be
scanned or the body can be separated from the shell when it
has been dried. To separate the body from the shell with
minimal damage use a micro-scalpel or a piece of razor
blade, not a needle. After SEM documentation, the body can
be rehydrated in water for radula extraction.
Coiled gastropods
Because dried bodies are difficult to remove, the shell
should
be
photographed
(SEM
or
standard
macrophotography) before trying to remove the body in case
the shell is damaged. Tightly coiled species are more difficult
than those with few whorls and a large aperture. From those
with a large aperture, the body can usually be extracted with
a fine needle with a small hook by inserting it at the
columellar side after rehydration in weak buffered formalin
and traces of a neutral detergent. Rehydration may take an
hour to a couple of days depending on the condition of the
body and the size. To speed up rehydration, evacuate the air
with a vacuum pump; use a small container to avoid
implosion by the glass breaking. The water may suddenly
start boiling when the pressure drops. A regular 20 ml glass
jar with a rubber stopper, connected to a water-jet pump with
a transparent polyethylene hose works well. In this way airbubbles trapped inside the shell are replaced with water.
FIGURE 8. Different approaches to visualize surface texture using SEM as shown on a fossil micromollusc kindly made available by Mike
Vendrasco and Christine Fernandez (phosphatic internal mold of Mellopegma sp., Middle Cambrian, Georgina Basin, Australia). All images
were taken on a Zeiss EVO40XVP with an accelerating voltage of 20 kV and rather high probe current of 300 pA at a working distance of
10 mm. Scale bar = 100 µm. A. Secondary electron detector (SED) with bias of +300 V. This is the usual operating condition of the SED. B.
SED with bias of ±0 V. C. Secondary electron detector with bias of -50 V. All secondary electrons are repelled, and the SED operates as a
backscatter detector. D. SED with bias of -250 V. SED operating as a backscatter detector, excluding the secondary electrons as well as the
lower energy backscatter electrons. E. All four quadrants of QBSD. Macroscopic relief is completely obscured, and only microscopic
irregularities are visible. F. Single quadrant of four quadrant scintillating backscatter detector (QBSD). It shows slightly more macroscopic
contrast than G, but also has a slightly lower signal to noise ratio as shown by the somewhat more granular image. G. Two adjacent quadrants
of QBSD with normal polarity producing positive image. H. Two adjacent quadrants (opposite ones from G) of QBSD with inverted polarity
producing negative image. I. G and H combined. Notice fine detail of surface is best visible in A and E. The macroscopic undulations of the
shell are best seen with the SED used as a backscatter detector (C and D), The combination of normal polarity and inverted polarity signals
from the QBSD highlights the macroscopic undulations, while smoothing the minor surface irregularities. Images DLG.
Usually the body cannot simply be pulled out straight,
but is ‘unscrewed’ while removing. Often the body is
retracted too far to be extracted as described above. Make a
23
small hole in the shell about 1.25 whorls up the spire from
the outer lip (see under Tools: Drills for methods). Carefully
drilling the hole will not destroy the shell and good SEM
24
pictures can still be obtained. Then cut the body, particularly
as much of the columellar muscle as possible, using a needle.
The lower body can then be pushed out either directly, or
indirectly by inserting small pieces of wet tissue paper or
cotton wool through the hole with fine forceps or a needle.
Some of the visceral coil will be probably be left in the shell,
but the head-foot is usually rather easily removed.
Species with a very tightly coiled shell may be difficult
to process without severe damage to the shell. Such
specimens can be broken in two at mid-height, soaked and
the soft parts flushed out by inserting the apical part of the
lower half into a fine pipette. Let the water drain into a fine
mesh that can be examined under the stereo-microscope if
the body is fragmented. Sometimes the radula can be
obtained even from almost completely decayed and
fragmented remains of a poorly preserved or rotten
specimen. The two remaining pieces of the shell can usually
be glued together. Dilute the glue if it dries too fast.
The easiest way to manipulate the specimens is to hold
the specimen between index finger and thumb of your hand
with less dexterity (usually left) and moisten the specimens
and the fingertips with the immersing fluid using a fine
brush; use gloves if necessary (or work with harmless
chemicals) and a stereo-microscope as needed. With your
right hand, apply the appropriate tools (pins, forceps) to
remove the body. This technique is generally superior to
manipulating the specimen fully immersed in a suitable dish
with two instruments (needles, brush, forceps). Attempts to
construct specimen cradles with pins in a wax tray are
disappointing. The finger-method works with specimens
down to less than 1 mm in size.
Opercula
Many microgastropods produce opercula of a variety of
forms and sturdiness: from strong calcified ones to wafer
thin varieties. It is usually still attached to the foot and may
be retracted into the aperture of the shell. To avoid damage to
the operculum when extracting the body, try removing the
operculum by inserting under it either a micro-scalpel, a pair
of fine watchmakers forceps, or a fine needle. If the
operculum falls off at the first touch, the specimen is
probably more or less decayed; there may be little detail
available in the soft parts and the radula may need extra care.
During the process of extraction, hold the shell under the
microscope between thumb and index finger (as described
above). Opercula are easily imaged by SEM when still in
position in the aperture if the animal is not too far retracted
and very thin opercula are often better imaged in this way.
Contrast of structural details such as growth rings may be
indistinct when mounting thin corneous opercula on doublesided carbon adhesives. If separate mounting of thin opercula
is desirable, mount them only with part of the operculum
touching the carbon adhesive, or place the moist operculum
on dry PVA glue.
Removing the shell the fast way
As an alternative to the above methods, the shell can be
decalcified in dilute hydrochloric acid (HCl: 2–5%), which
should only take a few minutes. However, some bubbles will
form and the procedure may rupture the tissues when there
are internal deposits of carbonates. An alcoholic solution of
HCl is less damaging to the soft parts as less carbon dioxide
is generated and the bubbles are smaller due to the lower
surface tension of the ethanol. Decalcification with ethylene
diamino-tetraacetic acid (EDTA) is possible in aqueous
solutions (5–20%, pH 7.0 adjusted with 1N NaOH), and is
favoured by some for animal preparation for histology (not
further covered here), but is very slow. A mixture of formic
or acetic acid and formalin can be used to fix and decalcify in
a single step (as in Bouin’s fluid). Cracking the shell is
another simple option to get access to the animal (see section
Storage above).
Once the shell is removed from the body, several
options are available: investigation of external morphology
by light microscopy or SEM (see Tissue preparation for SEM
below); investigation of internal anatomy by dissection or
histology or radula extraction.
Tissue preparation for SEM
Once the animal has been removed from the shell, it
must be dried prior to further inspection using the SEM. In
some instances, it is advisable to dry the body inside the shell
and examine the exposed head-foot characters visible on the
relaxed and nicely extended animal. Drying of tissue from
aqueous or alcoholic solutions directly leads to severe tissue
shrinkage and makes detailed inspection of the external
morphology impossible.
Proper drying of animals can be carried out by three
methods: critical point drying (CPD: Fig. 9);
hexamethyldisilizane (HMDS); and freeze drying (Sasaki
1998). CPD and freeze drying require specialised equipment.
HMDS, on the other hand, can be used at room temperature,
although a fume hood is necessary for safe handling of the
liquid. Both CPD and HMDS often give suitable results
although there are sometimes unexplainable failures. In both
cases, the specimen has to be taken through a graded ethanol
series to pure, undiluted electron microscopy grade ethanol.
‘Pure’ ethanol used for storage of specimens is actually only
approximately 95% and is unsuitable for tissue dehydration,
and most problems arise due to insufficient dehydration.
From pure ethanol, the ethanol has to be replaced by either
CO2 in the case of CPD, or HMDS, through several fluid
changes. For HMDS, better results are obtained if the liquid
is evaporated more slowly in a covered dish overnight, as
opposed to an open one in a few minutes. For freeze drying,
the specimen is placed in t-butyl alcohol and the freeze
drying machine automatically applies a vacuum to the cooled
specimen vessel. The advantage over CPD is that typically
the equipment is all automatic, not requiring the operator to
fill and empty the specimen reservoir with liquid CO2. There
are some semiautomatic CPD. The results of CPD and freeze
drying are comparable (see also Goldstein et al. 1992:
chapter 12.5.4, fig. 12.9). Specimens from historical
collections are often suitable for SEM tissue preparation
(Fig. 9).
25
FIGURE 9. Comparison of critical point dried historical specimen with recently collected material. A, B. Old specimen: Puncturella
noachina (Linnaeus, 1758), SMNH old catalogue #116, Pröven, Greenland, 29–73 m, Leg O. Torell. A. Entire animal. Scale bar = 2 mm. B.
Part of gill enlarged. Scale bar = 200 µm. C, D. Specimen collected two weeks before CPD, May, 1996. Emarginula crassa Sowerby I, 1813,
Koster Area, Sweden, approximately 50 m. C. Entire animal. Scale bar = 2 mm. D. Part of gill enlarged. Scale bar = 200 µm. Images AW.
26
Extraction of radulae from micromolluscs
•
The radulae of micromolluscs can be very small, often
making manipulation daunting. However, with some practice
and patience, radulae a fraction of a millimetre long can
routinely be successfully mounted. If possible, use an adult
specimen unless an ontogenetic study is specifically carried
out, as the radulae of many species change morphology with
age (Warén 1990). In some cases so-called generic characters
are obtainable only from adult radulae.
It is recommended that specimens used for radular
preparation should be photographed prior to radular
extraction, especially if there is any doubt as to identity. The
shell may be destroyed when attempting to remove the body
and, if chemical treatment is used, tissue-dissolving agents
contribute to the deterioration of the shell. In some cases,
species-level identification requires the observation of
minute details such as protoconch microsculpture that cannot
be observed with a light microscope and necessitates the use
of SEM.
While the radula can be dissected from larger
microgastropods, there is a danger of damaging it. A safer
method is to dissolve the buccal mass or even the entire
animal.
There are a number of methods used for dissolving the
tissue surrounding the radula. The simplest and quickest
methods can be used for most gastropods. Gentler, more time
consuming and more complicated methods may be necessary
for some of the groups with delicate radulae or radular
membranes. The latter include:
•
•
•
Patellogastropoda. Damaged by strongly alkaline
agents; mineralised cusps fall off and the remaining
parts are partly dissolved.
Monoplacophora. Teeth damaged by strong alkali.
Lepetelloidea. Teeth may get distorted and crack in
strong KOH.
Vetigastropoda.
o
In some with thin and slender teeth (e.g.,
calliostomatids, trochaclids), the teeth become
softer and tend to stick together. These should,
after rinsing and cleaning, be soaked in 50%
ethanol and mounted in at least 80% ethanol, to
reduce the risk of the teeth sticking together. In
ethanol the teeth become stiffer and there is less
surface tension.
o
In some fissurellids (Cosmetalepas) the teeth fall
off the radular membrane when treated with strong
alkali.
We advise against trying to dissolve the animal inside
the shell. Strong NaOH or KOH will damage the organic
matrix in the shell (Strasoldo 1991) and the remaining
hydroxide will react with aerial carbon dioxide, to form a
crystalline or powdery coating that cannot be removed (Fig.
10). However, with some sturdy gastropods, such as
marginellids, no adverse effects have been reported (Coovert
and Coovert 1987) and short periods of maceration of tissue
inside the shell can be tried. Shells of more fragile
gastropods, such as scissurellids, will break when sonicated
after such treatment, whereas they are stable before radular
extraction. Shells exposed to tissue dissolving agents also
deteriorate over time and can be completely broken down in
as little as 10 years, while enzymes in detergents can destroy
a shell in a few hours, due to the low pH. Proteinase K,
commonly used in DNA extraction from tissues, is most
destructive to the shell. Whereas hydroxide treatment usually
leaves a recognisable shell behind, proteinase K will
fragment shells.
FIGURE 10. A. Protoconch of shell (Scissurellidae) after sodium hydroxide treatment. B. Similar protoconch without hydroxide
treatment. Note the tunnelling and recrystallisation in A. Scale bars = 100 µm. Images DLG.
Standard method
The tissue of the body can be dissolved in 5–40%
NaOH, KOH, bleach, or by using proteinase K as part of
DNA extraction. Sodium dodecyl sulphate (SDS = sodium
lauryl sulphate) provides a further alternative. The hydroxide
concentration indicated in the literature is quite variable;
higher concentrations are advocated by those who like to
speed up the dissolution of the tissue (e.g., Coovert and
27
Coovert 1987), which can be additionally accelerated by
heating specimens up to 100°C. Such drastic methods are
usually not necessary for micromolluscs and usually lower
hydroxide concentrations (5–10%) and no or low heat (30–
50°C) are suitable. Lindberg (1977) noted adverse effects of
heating on patellogastropod radulae, particularly the
contraction of the radular membrane and also the separation
of teeth from the radular membrane.
Bleach offers a low cost option but dissolving power
varies from brand to brand. Test treatments should be carried
out before risking rare material. Some brands of bleach
dissolve soft tissue in a shorter time and cause fewer
artefacts on radular teeth than KOH. Some of us consider
bleach too aggressive and favour the inexpensive SDS, while
others (AW, WFP) prefer KOH, as the specimen can be left
in the solution for several days if necessary. With dissected
patellogastropod radulae, to avoid destruction TS and AW
add bleach drop by drop until maceration is visible.
Never place paper identifying tags in the extraction
solution, because crystal deposits will form on the radula
(e.g., Geiger 1999: figs. 11, 12). It is not clear what the
chemical composition of the deposit is, but it may be formed
as a precipitate from the hydroxide and the filler substance
used in most papers. Always label the containers on the
outside.
Manipulation of the radula and the body can be
achieved by a variety of implements; forceps and needles,
paint brushes, Pasteur and Eppendorf pipettes (see Tools
above).
Gentle methods. Patellogastropod radulae can be
extracted by dissolving the head or body in a large quantity
of very weak (ca 0.1%) KOH at room temperature. A small
body requires several days, but the radula is not damaged.
The body will not dissolve completely, but after a few days
the buccal mass with the radula can usually be dissected out
with no great risk of damage. After removal, the radula
should be soaked even longer in a new bath of the same
solution to remove most of the soft tissue, although usually
some remains. If necessary, the remainder can be removed
by quickly dipping the radula in lukewarm, diluted (1:3–1:5)
commercial bleach for a few seconds and then vigorously
rinsing it. Make sure that the radula can be retrieved quickly
if accidentally dropped into the bleach.
A strong solution of sodium lauryl sulphate is very
gentle but takes a few days. It is difficult to get the radula
completely clean but rinses in hot water or diluted bleach as
above can be used. For larger radulae a fine paint brush can
be used as a starter.
Risso-Domingue (1961) discussed other amines useful
for radular extraction, particularly in cases where the radular
membrane is weak and hydroxide treatment results in teeth
becoming isolated, rather than remaining attached to the
radular membrane.
Maceration
For the maceration process, three different types of
vessels can be used. All dishes should be made of glass;
plastic charges statically and metal surfaces produce
precipitates with hydroxide solutions.
•
•
Medium specimens (2–6 mm). Covered square embryo
bowl is suitable, filled to about half its depth. When
moving, be careful to avoid the fluid entering between
the bowl and the lid.
Small (< 2 mm). Use a depression slide, 5–6 mm thick,
with a depression as deep as possible and
approximately 15 mm diameter. As a lid use a
depression slide with a wider depression of any depth
(as described above). Never use a regular, flat slide or
cover slip because condensation will enter the space
between the lid and bottom and the two slides will stick
together and sometimes also contact the fluid in the
bowl. Avoid jerky movements (and hence slop) when
transporting the slide sandwich, or when removing the
lid.
Eppendorf tubes are not recommended, despite the
advantages of a tight fitting lid and the possibility of
spinning down the solids (i.e., radula and shell), because it is
very difficult to remove the radula from the narrow tube.
Heat incubation is best accomplished in a small
incubator that allows for precise (within 2–3°C) temperature
control. Cabinet incubators, slide warmer plates and dry bath
incubators are suitable. The incubator should be situated
close to the microscope preparation area to minimise risks
associated with transport.
It is important to cover the container during maceration,
particularly if heating the specimen, because crystals of
sodium carbonate (flocculent material of Mikkelsen 1985)
may otherwise form due to absorption of carbon dioxide
from the air. These do not redissolve when adding small
amounts of water and make it very difficult to find a tiny
radula. The formation of insoluble crystals has been
observed particularly when macerating the animal within the
shell, but here a calcium compound may be involved.
Sometimes different water-soluble crystals are formed after
heating.
A number of chemicals can be used for maceration.
Among the hydroxides, KOH is better than NaOH since it is
less hygroscopic and reacts less quickly with aerial carbon
dioxide. Always use analytic grade and preferably the kind
that comes as small spheres or half-spheres because the
powdered form reacts faster with aerial carbon dioxide. Do
not use a stock solution but prepare it fresh in situ with
distilled water, to avoid unnecessary precipitates. The KOH
tablets should be semitransparent-porcellanous; they become
white and dull with age because they react with aerial carbon
dioxide. Thus, keep the KOH in an airtight container.
Proteinase K is a safe cleaning agent of radulae from
fresh, frozen, or alcohol fixed material and will not result in
any damage to the radula (cf., prolonged exposure to
hydroxides). Radula and operculum can be collected from
the filter of spin columns (Fig. 11). Cut the spin column just
above the plastic ring retaining the filter and carefully
examine the filter as well as the plastic wall holding the filter
under a stereomicroscope. If the pieces cannot be found,
28
slowly rotate the column bottom while carefully pulling the
filter from underneath the retention ring from one segment
after the other. Wash the radula in water and mount. Retrieval
rate is approximately 80–90%. Proteinase K, though, does
not work well on formalin fixed material, in which proteins
of the tissue have been cross-linked (Holznagel 1998).
FIGURE 11. Recovery of the radula from a spin column used for DNA extraction. A. Cut the column above the retainer ring for the filter.
B. Oblique view of the cut column. The arrow highlights the radula. C. Enlargement of B with the radula visible near the edge of the filter
retainer ring. D. The radula has been removed with fine forceps and is placed into a gelatine capsule for temporary storage. Images DLG.
Cleaning the radula
To find the radula after maceration, examine the
container under the stereo-microscope, using the substage
illuminator. Incident light may make particles shine and hide
the radula. Horizontal illumination through the sides of the
glass vessel may be employed if a substage illuminator is not
available (pseudo dark field), but will only be effective if the
fluid is clear.
The radula has to be washed in water and then possibly
in ethanol. Two methods are outlined below.
In the first, fluid is exchanged, minimising handling of
the radula. The fluid can be removed with a Pasteur pipette
or a pipettor and discarded into a separate container in case
the radula is inadvertently removed with the fluid. The
remaining fluid film can be blotted with a fragment of folded
paper tissue, keeping it a safe distance from the radula. A
small radula that is stuck to the paper will usually be lost
because it is difficult to distinguish the radula from the paper
fibres. It is easiest to move the radula when dry by touching
it with a moist tungsten needle, or very fine, moist insect pin.
Eyelashes or other hairs are too flexible for radular
manipulations.
In the second method, the radula is transferred in a
succession of fluids, which prevents it from drying. The
radula is picked up with fine entomological forceps, a bent
needle, or with a pipettor and transferred into a series of
washing solutions. Do not use a paintbrush to transfer a
radula, as it can easily get entangled in the bristles and lost.
Minute radulae can be washed in drops of water on a glass
histology slide. Clean radulae can be stored in tubes in 80%
ethanol.
Sometimes the maceration solution is very dirty and it
may help to dilute it with distilled water or more KOH
solution. Heat may also help if nothing else does. As a final
resort, the contents of the container can be poured through a
very fine sieve with low edges. The mesh should be a
29
maximum of 1/10 of the estimated length of the radula (as a
general guideline, the radula is approximately one tenth to
one third the length of the shell/body). Rinse with hot water,
which will usually dissolve the remaining grease, tissues and
particles.
Medium size radulae may be brushed with a very fine
paint brush (#00 or #000, preferably artificial hair), while
holding the end of the radula with a needle pressed against
the glass.
With very small radulae there are few possibilities for
further cleaning; they can be moved in the water or scratched
with a fine needle against the glass so it makes small
vibrating jerks. Use a needle with its tip bent close to 90° and
scratch with the tip at right angle to the glass. If the radula
still looks dirty, try a new KOH bath. If nothing else helps,
try diluted, warm bleach (rather than cold, more concentrated
bleach, as less hypochlorite is carried over to the rinse).
Neogastropod radulae have a sheath of thin, transparent
cuticle surrounding the part not in use. This should be
removed by a quick dip in commercial bleach and
subsequently rinsed, or with a fine needle when it has been
glued and is drying. Do this at the same time as the lateral
teeth are unfolded. Do not be concerned about the radular
membrane extensions that under-lie the anterior part of the
radula in vetigastropods and taenioglossate caenogastropods.
They may even facilitate mounting.
In general, do not use an ultrasonic cleaner, unless there
are spare radulae. It is a good method for cleaning many
taenioglossate radulae, but in some cases teeth may fall off,
get entangled, or the radula may disintegrate entirely. Not
even experienced practitioners can predict the result.
Orientation
Identifying the orientation of the radula can be difficult.
Use the highest magnification of the dissecting microscope.
If the radula is too small to see individual teeth, rely upon the
appearance. The anterior margin of the radula with
completely formed teeth has two flaps. The teeth there are
facing outwards from the curve. The posterior end of the
radula is tapered and thinner than the anterior part.
The upper and lower surface of the radula can be
difficult to distinguish but they reflect light differently; the
underside has a more glassy appearance, whereas the top is
more sparkling. The long axis of the radula tends to curl with
the base inside; the curling of the radula along its long axis is
often impossible to see. It is easy to distinguish which way
up a radula is lying by transferring a wet radula to a fragment
of a cover slip on a slide and examining it under low-power
using a compound microscope. The radula can then be
mounted directly on the cover slip. For very small radulae, it
may be difficult to judge whether it has been properly
mounted. To ensure suitable results, several radulae can be
mounted, or the radula can be folded in a L or V shape so
that both sides are facing upwards (this method is especially
useful for elongate radulae and for species with short teeth).
Tilting the stub towards the light will usually show the
reflection pattern better.
Special techniques for small radulae
Several of techniques have been successfully used to
mount very small radulae. These include the use of different
surfaces such as:
•
SEM mounting of micromollusc radulae
Mounting can be achieved by a variety of techniques, the
choice mostly depending upon size and the type of radula
(Fig. 12). Techniques for light microscopy are not discussed
here as they are detailed elsewhere (e.g., Mikkelsen 1985;
Coovert and Coovert 1987; Bradner and Kay 1995).
Hickman (1977) discussed the types of information to be
gained from both light and electron microscopy of radulae
and stressed the complimentary nature of both techniques.
Large radulae (>1 mm long) can be mounted on doublesided sticky tape, or double-sided carbon tabs. Medium sized
radulae can be mounted on double-sided sticky tape and
manipulated in a drop of water or ethanol (Fig. 12D). Some
workers also like to roll the radula onto pins or conical
mounts as the information gained increases the more the
radula is folded or twisted (Bradner and Kay, 1995). The
slow evaporation method of Moretzsohn (2004) seems to
work well for larger radulae, however it is not clear whether
it would work as well if applied to the preparation of small
radulae. Minute radulae can be dried directly onto a piece of
coverslip (see below). Consideration of storage options and
times should also be taken into account when determining
the mounting method used.
•
•
Double-sided sticky carbon tabs (Fig. 12C). Other
sticky substances include double-sided tape and wet,
blackened, photographic paper. Place a small drop—the
amount of water that sticks to the head of a fine pin—
on one of these surfaces mounted on a metal stub. Place
the wet radula into this drop and orient it.
Radulae of species with many teeth per row (e.g.,
rhipidoglossate, ptenoglossate, many pulmonates) can
be placed directly on a glass cover-slip glued onto a
stub. Orientate the radula in a small drop of water and
let the water evaporate. Excess water can be removed
with great care with a fine strip of filter paper (see
above). Alternatively, pull the radula out of a drop of
water and let it dry in place. The radula will adhere to
the glass without any need for adhesive. Some narrow,
taenioglossate radulae may tend to spring off the glass
and may need to be mounted on carbon tape. Radulae
mounted on cover glass can, with some practice, be
removed with water even after viewing in the SEM.
Glue a histology cover-slip (round, 12 mm diameter) to
the mid-point of the histo-slide with a very small
amount of saliva. Let it dry well in an incubator. Cover
part of the cover-slip with a thin layer of <1:1 diluted
polyvinyl acetate glue in water. A horseshoe shaped
glue patch works well but it can be modified according
to needs. The glue should be applied thicker for larger
radulae than for smaller ones. Let the glue dry
30
•
thoroughly, preferably overnight in an incubator. When
transferring radulae onto the cover slip, be careful to
prevent water getting under the cover slip as it will
unstick it.
The glass cover slip is released from the histo-slide base
by applying some water to the edge of the cover slip.
Capillary forces will pull the water under the cover slip
and dissolve the glue. The loose cover slip is then
mounted on a SEM stub with new glue.
Rhipidoglossate (Vetigastropoda) and docoglossate
(Patellogastropoda, Polyplacophora) radulae have
overlapping lateral and marginal teeth, which obscure
other teeth. To mount these types of radulae, glue some
thin pieces of wire, or hair, or needles of a diameter
close to the width of the radula and cover them with
glue (Fig. 12A, B). Cover slips may also be prepared
with a series of wires etc. of different widths. Radulae
can then be mounted longitudinally, on top of these
wires with the marginal teeth bent outwards and
downwards. This affords a better view than mounting
radulae across the wire (e.g., Strasoldo 1991), but
requires some practice. Breaking up a flat-mounted
radula will also give the necessary data.
The orientation of a mounted radula can be doublechecked under a light microscope with a 25 or 40x objective
although care is needed as the working distance is only 1–0.2
mm, depending on the lens. For radulae mounted on glass
slides, commonly available transmitted light microscopes
can be employed. Stub mounted radulae can be checked with
compound microscopes equipped for (or improvised) epiillumination or with high power stereomicroscopes.
A radula that has been accidentally improperly mounted
can often be released from the mounting surface.
•
•
For PVA glue, add water with a paint brush or a fine
pipette, soak the radula, remove and remount it. Do not
disturb the glue excessively, as it may invade the radula.
If mounted on double-sided carbon tabs or tape, a
radula can be released by soaking it in a large drop of
water for a minute and peeling it off the carbon tab from
one end. Then, the radula may be remounted and dried
again. Once the radula has been sputter coated and
viewed in the SEM, the radula is more firmly stuck to
the carbon tab. Water will usually not be sufficient to
release the radula, but ethanol usually works. Coated
radulae are usually stiffer and more brittle than fresh
ones.
Very small specimens
The following variation of the method above has been
used for very small radulae (e.g., those of cimids with a
length of ca 60 µm) or post-larval gastropods. Soak the
specimen in a small quantity of distilled water: 1–6 drops
with a fine pipette in the depression slide or 1/3 or less of the
depth of the solid watch glass. Dissolve 1/4 of a tablet of
KOH to 50 µl (= 1–2 drops) of water. The tablets can be
readily split using a small stainless wire cutter. Heat the
solution in the incubator at 50°C for 15 minutes or a little
more. Usually the specimen will not dissolve but remain in a
lump of clarified tissues. Transfer the lump with a needle
(bent 90°), or a pair of forceps (an inferior method because
more fluid is transferred) to a drop of distilled water on a
cover-slip. Usually the lump of tissue will dissolve in a
fraction of a second. Add a drop of distilled water, close to,
but separated from, the point where the specimen was
dissolved and pull the radula over to this without allowing
the two drops to merge. Remove the dirty water with a small
piece of lint free tissue curled around the tips of a pair of
forceps and pinched in position by a little rubber band
around the upper part of the forceps. Wash the radula a
couple of times more—the radula should now be in very
clean water.
Manipulation of radula
Manipulation techniques vary with the mounting
surface chosen. Mark the position of the radulae, because
they are often easier to spot with a light microscope than in
the SEM.
For carbon tabs, the radula can be manipulated in a
small (preferably distilled) water drop using a pair of fine
tungsten needles. The water will evaporate at room
temperature in one to two minutes (ethanol evaporates too
fast for many small radulae, although it works very well with
larger radulae and has less surface tension than water [but see
remarks on some vetigastropod radulae above]). The surface
tension of the water will help in flattening the radula along
its long axis. At the point when the radula is still moist,
but when there is no free water around the radula (a period of
about two to three seconds), gently spread the radula out with
the needles. The outer rows of radular teeth tend to fold
over the central field when they dry, therefore it is important
FIGURE 12. A, B. Radulae mounted on long axis of pins. Scale bar = 2 mm. B, enlarged view of A. Scale bar = 200 µm. C. Assorted radulae
mounted on double sticky tape/carbon tab. Scale bar = 2 mm. D. Radula of Scissurella mirifica (A. Adams, 1862) showing tear in centre. Scale
bar = 200 µm. E. Enlargement of D showing full exposure of base of lateral and marginal teeth. Scale bar = 20 µm. F. Same radula as D. in
area not torn; note less exposure of the base of the teeth. Scale bar = 20 µm. G. Dikoleps nitens (Philippi, 1844) [Skeneidae] juvenile. Scale bar
= 10 µm. H. Dikoleps nitens adult, approximately 1 mm shell diameter. Scale bar = 10 µm. I-K. Haliotis discus hannai Ino, 1953 seven days
old larvae from a culture in Japan. I. Entire radula. Scale bar = 10 µm. J. Full width of row enlarged. Scale bar = 10 µm. K. Larval shell of
animal from which radula was obtained. Scale bar = 100 µm. Images: A–C, G–K: AW; D–F: DLG.
31
32
to physically spread the radula open. If the spreading does
not succeed, place a drop of (distilled) water on the radula
and try again. Only one part of the radula needs to be in a
good orientation without too much damage. Breaking up the
radula into segments and separating some teeth will increase
the information available (Figs 12D, E, G). For this fine
manipulation holding one’s breath helps, because ribcage
movement induces movement in the arms and hands. Mark
the radulae by drawing a circle around them with an old
needle or pair of forceps.
On glass slides, start by straightening out the radula. If
it is long, you may cut it to get a better view of tooth bases.
Small radula may be cut with two needles, crossing them to
imitate a pair of scissors. Start with the front of the radula,
because the posterior part of the radula is usually easier to
work with, as the teeth are normally not fully formed. Pull
out the front half with a needle to the edge of the water,
preferably tooth side up and at a right angle to the edge.
Quickly pull it out from the water with the needle, across the
glass and up on the glue bed. This requires some practice.
The intention is that the wet radula will soak the dry glue
enough to make it stick and firmly attach the radula. Wet
glue would invade the radula due to capillary forces.
While the radula is drying, after it has started sticking to
the substrate but before it is dry, spread out the lateral teeth,
preferably so they laterally stick to the glue. Use a needle
bent like an ice-hockey stick. You can also use a moist triple
zero paint brush, or a cat’s whisker in a holder works well for
smaller radulae. Small radulae usually stick without glue.
The position of the radula can be marked with a fine tipped
pen or dots of PVA glue.
A dry, flat-mounted radula can either be torn in the
outer part of the central field, approximately in the middle of
the ribbon, or a few rows of the radula can be cut with a
scalpel (Figs 12D, E, G) or torn with needles. Tearing a
radula can provide a better view of the basal plates of the
teeth and the teeth in the central field of Patellogastropoda.
In vetigastropods, it will often also show diagnostic teeth at
the boundary of the lateral and marginal tooth fields (lateromarginal plate), which are typically obscured one behind the
other. Although the result of such destructive approaches is
often unpredictable, it is recommended. Curving of the
radula (e.g., by using wire mounting) will also provide good
visual access to the base of the teeth, but a physical
separation of rows by tearing the radula is usually superior.
The merits and problems of wire mounting and tearing are a
trade-off. Bradner and Kay (1995) suggested removing a few
rows at the end of the radula for a better view of the teeth but
we suggest that a tear in a more central to slightly anterior
portion is more desirable, because of wear at the very
anterior end of the radula. Manipulation of the radula other
than standard mounting is not necessary for most
neogastropods, because there is little overlap of the teeth.
Transfer the radula from the last cleaning step to a drop
of water on the chosen mounting substrate with a hooked
needle. This has to be done fast, so the radula hanging on the
needle does not dry. It dries faster in pure water than in
KOH. A very small radula may dry in two to three seconds,
depending on aerial humidity. If it dries, the radula will stick
to the needle and may be difficult to remove, or it may
crumble more or less irreversibly.
Keep radular mounts in progress covered by an upsidedown glass bowl to avoid dust. To handle SEM subs, use
designated stub forceps; for stubs with a peg (e.g.,
Cambridge), rather use the model for grabbing the peg, as
opposed to the rim of the stub.
Radula, histology and X-ray computer tomography
In cases where the maximum information should be
obtained from a single specimen, a conscientious choice
must be made as to which data is most important. External
morphology can be obtained from histological sections
through three dimensional reconstruction (e.g., Amira 3.1®),
but radular structures are too fine to be accurately
reconstructed; at most the major radula types (e.g.,
docoglossage, rhipidoglossate, stenoglossate) can be
identified.
The most promising method for future radular studies
seems to be X-ray computer tomography (e.g., Hagadorn et
al. 2006), but so far the resolution (ca 1 µm: http://
www.microphotonics.com/skymto.html) is too coarse for
micromolluscs. Regardless, it requires the most sophisticated
instrument of its kind plus a cyclotron and involves much
work on reconstruction.
Three-dimensional reconstruction of animals is an
exciting new avenue. Given that even basic anatomical
features are difficult to observe in dissected micromolluscs,
the only alternatives are histological serial sections with
subsequent computer assisted reconstruction. Some of us
have begun to apply these techniques to some specimens
with 1 mm shells, the animals being half that size. We show
here (Fig. 13) a section through the head region of Sinezona
rimuloides (Carpenter, 1865) and the reconstructed pairs of
odontophore cartilages, the pedal ganglia and the partially
embedded statocysts (Fig. 12). The procedures are very
labour intensive and require appropriate computer hardware,
including a graphics tablet. However, it is possible to take
cross-sections and to view the anatomy in any orientation. A
dorsal view is shown in Figure 12. We expect software
developments to make the techniques easier in their
application as well as more reasonably priced.
Optical photography
Some specimens may not be placed in the high vacuum
environment of the SEM and also, if colour is required, light
optical imaging must be employed. Two main approaches
can be pursued.
SLR camera (film or digital)
A general review on shell photography has recently
been provided elsewhere (Geiger 2006b); Häuser et al.
(2005) provided an overview on digital imaging of biological
type specimens. Marco lenses usually provide 1:1
magnification (occasionally only 0.5:1), which covers an
area of 24 x 36 mm of film, or with 2/3 digital sensors an
area of 16 x 24 mm; hence, micromolluscs cannot be
photographed full frame with regular macro lenses.
Magnifications of at least 3:1 (5 mm specimen on 2/3 digital
sensor) to 24:1 (1 mm shell on 35 mm film) are required.
With standard photographic equipment (bellows, normal
50 mm lens reversed) magnifications of approximately 5:1
can be obtained (Fig. 3D–F). Some lenses (e.g., Canon 65
33
mm macro) can provide magnifications from 1:1 up to 6:1
without extension rings or bellows. With short focal length
macro head lenses (e.g., Zeiss Luminar series) on long
bellows units, magnifications of up to 12–20:1 can be
reached, although procedures are quite tedious. Dedicated
microphotography systems are available (e.g., Microptics
Inc.: www.microptics-usa.com). Mirror lock-up is advisable
to minimize the effect of shutter and mirror vibrations.
FIGURE 13. An example of histology and three-dimensional reconstruction from Sinezona rimuloides (Carpenter, 1865). The body axes in
the plane of the image are labelled. A. A histological semithin cross section in the head region. The anterior-posterior body axis is at right
angle to the image plane. Plastic embedded specimen sectioned at 2 µm and stained with multiple stain (Polysciences). B. Threedimensional reconstruction of select organs in anterior portion of body. The model was rotated in the computer by 90°: the dorsal-ventral
axis is at right angle to the image plane. Images DLG.
With most digital cameras, images of equivalent quality
to those taken with film can be obtained. However, the
optical principles still need to be observed. Avoid zoom
lenses, macroconverters, teleextenders and diopter lenses.
With digital cameras, it is usually more difficult to reverse
lenses and program automatisation may not take in to
account the special considerations of extreme macrophotography. The problems are not the digital capture
mechanism per se, however, modern cameras have many
automatic functions that are transmitted through electrical
contacts on the lens. Once the lens is reversed, the
information flow is interrupted and the automatic closure of
the f-stop when the shutter is pressed is often no longer
available. In that case, the f-stop has to be closed up front,
darkening the image significantly, which makes focusing and
composition much more daunting.
The current tendency to focus on the number of
megapixels (MP) of digital cameras is exaggerated. With the
vast majority of intermediate cameras, publication-quality
images can be produced. A single specimen is at most shown
at 1/4 page size, approximately 7 x 10 cm. At the usual
printing resolution of 300 dpi, a file size of 3.3 MP is
required and it is only for special large format images that
larger file sizes will be necessary. Dynamic range (Dmax),
bit depth of output file (preferably 16 bit per channel) and
file formats (tif, RAW, ProPhoto; not jpeg) are more
significant imaging attributes.
Focusing aids such as microprisms and split image on
the focusing screen are darkened at higher effective f-stops,
i.e., also at higher magnifications on bellows and fully open
diaphragm, because the f-stop increases due to the spreading
of the light beam in the bellows unit. Camera systems with
interchangeable focusing screens are therefore advantageous
to allow fine matt or even clear screens to be employed.
However, clear screens often used on autofocus cameras
make accurate focusing in the low magnification close-up
range more difficult and autofocus rarely focuses where
intended. As a consequence, manual focus adjustments in
these situations is more difficult than with traditional matt
focusing screens with microprisms and split image. Make
sure that the viewfinder is optical (i.e., made of a glass
prism) and not a LCD screen. LCD screens built into
cameras do not show sufficient detail for critical focus.
Accordingly, manual focus override is equally important.
34
Most digital SLR cameras can be connected to a computer
and the monitor image provides adequate resolution to verify
focus after the image has been taken.
Intermediately priced compact digital cameras with
supposed macro capability generally produce inferior results.
As an example, the Nikon Coolpix 8000 has a
macrofunction, flash and adjustable f-stop. However, the
maximum f-stop is f/8 (as opposed to f/22 on all dedicated
macrolenses), making the depth of field very shallow; at
closest focus, the flash does not illuminate the image area,
because the lens barrel produces a shadow; and the LCD
screens on the camera as well as in the view finder do not
permit accurate focus adjustments.
Improper file manipulation (e.g., working on .jpeg
rather than .tif/psd files, or in CMYK rather than RGB/Lab
colour space, or in 8 bit rather then 16 bit per channel, if
available) will produce inferior results. Please consult
appropriate works on digital imaging for further information
(e.g., Davies and Fennessy 2001; Sedgewick and Sedgewick
2002). The little known Lab colour space offers particular
advantages for un-sharp masking of colour images. In the Lchannel, the sharpening will only have effects on the
brightness value of the pixels, while not affecting their
colour value stored in the a and b channels (see Margulis
2005). In RGB, the brightness and colour values are a joint
value in each of the R, G and B channels and sharpening can
lead to colour artifacts.
Furthermore, particularly in flash-photography, the
exposure meter assumes 18% reflection, hence, exposure
compensation is often required. For instance, when
photographing a white shell against bright background, the
automatic exposure will assume that 18% of light is reflected
and produce a dull-grey image. Thus the photographer has to
instruct the camera to overexpose the image to obtain the
true white of the shell (see Geiger 2006b for step-by-step
instructions). The black box of matrix metering may increase
the percentage of acceptable images, but will inevitably lead
to failures. A thorough understanding of exposure and
exposure compensation is imperative. Many of these
adjustments can also be accomplished afterwards with digital
image manipulation, but the final result will be affected by
the quality of the source files.
The Bayer pattern of most digital cameras (CCD,
CMOS sensors) is a significant issue for digital colour
photography, as 2/3 of the colour information in each image
is interpolated. Three layer photo-sensors such as the Foveon
X3, currently only available in Sigma cameras, and co-site
sampling technique as implemented in the Zeiss microscope
camera Axiocam HRc, have overcome this limitation. The
opinions on the Foveon X3 chip are divided, as it has a lower
resolution compared to current CCDs and CMOS sensors,
while on the other hand, the larger pixels have a better
signal-to-noise ratio and capture all three colours at each site.
The signal-to-noise question also applies to the issue of 2/3
vs full-size digital sensors; at the same number of pixels, a
larger chip has larger pixels and a better signal-to-noise ratio.
Three chip cameras also avoid the Bayer pattern problem,
but light intensity reaching each sensor is only one third of
the original intensity because of the beam-splitter. The lower
light levels will cause somewhat elevated signal-to-noise
ratios. The latter can be improved by cooling of the imaging
chip. The two main cooling methods are fan and Peltier
stage. Because Peltier devices have no moving parts, they
cannot produce any vibrations, in contrast to a fan.
Stereo-microscope
For magnifications above 5x, a stereo-microscope with
photo attachment is advisable. Models with trinocular heads
or a dedicated photo-tube are preferable over ocular-mounted
systems. All photo-ports of modern stereo-microscopes use
only one of the light paths and, as the two light paths are at
an angle for stereoscopic viewing, lateral image shifts occur
when changing focus regardless of whether the
stereomicroscope is of Greenough or Telescope design.
Some instruments can counteract this image shift with
special attachments (e.g., Zeiss Discovery V8, V12 with
objective slider), which moves the objective so that the lightpath is in line with the photo-tube.
There are some older models designed for photography
(e.g., Wild M400 series, Zeiss Tessovar system). These
microscopes look like a stereomicroscope, although they
only have a single light path, hence true stereoscopic viewing
of specimens is impossible. As they are primarily intended
for imaging, this design feature should rather be viewed as an
asset than a deficiency.
Lenses for stereo-microscopes come in many different
quality ranges. Plan-apochromatic lenses produce flat images
and are fully colour corrected, but are also expensive. Plan
lenses are corrected to produce a flat imaging plane, but may
show pronounced colour fringes (= lateral colour, e.g., Leica
Plan 1x with yellow/blue fringes). In some cases, the image
plane is distinctly curved, resulting also in apparent
distortion of the object. One can test the image flatness and
distortion by photographing graph paper with 1 mm ruling;
ideally the image is sharp from the center to each corner and
the lines are exactly parallel to the edge of the image.
Lighting
Some of us prefer continuous light (incandescent,
fluorescent, LED) with long exposure times. The lighting is
more predictable, because the effect of any changes can be
observed in real time. Issues with colour temperature of the
light can either be addressed with colour filters or with a
custom white-balance in digital systems. Some of us prefer
flash photography because the ultra short exposure time
eliminates any possibility of vibrations (shutter and mirror in
SLR cameras, fan of fiber optics illuminators, person moving
in room) which may deteriorate the image sharpness and the
colour temperature is a well-defined. With some experience,
the results are equally predictable and, with digital capture,
rapid assessment of the results is possible. Use a high-power
flash unit, as the flash duration is proportional to the
discharge proportion: at 10% discharge, the flash duration is
around 1/10,000 s, whereas a full discharge will take
approximately 1/200 s. Some portable flashes have built-in
focusing lights and studio strobe systems generally have both
35
modeling lights and strobe tubes along with available
lighting modifiers (tubes, gabos, diffusers). Studio strobes
are bulky and fine adjustments are difficult to execute.
Depth of field
Light-optical systems are limited in the depth of field
that can be obtained. Depth-of-field should not be enlarged
excessively by closing a diaphragm because diffraction will
blur the image. Diffraction affects the diameter of the Airydiscs of two adjacent image points; when the Airy-discs of
two points separated by the circle of confusion (usually
0.3 mm) touch, then the two image points are no longer
distinct and the image appears blurry. The maximum
advantageous f-stop (fmax) under normal circumstances
(circle of confusion = 0.3 mm, 8 x 10” = 20 x 25 cm image at
reading distance) is 32/(magnification + 1). At 5:1, fmax =
32/(5+1) = 32/6 = f/6.3, at 9:1, fmax= f/3.2 (for details see
Geiger 2006b and references therein). On camera lenses, the
f/stop units are indicated, however on stereomicroscopes
with diaphragms, these values are not available and
resolution data are given for a fully open diaphragm.
Occasionally, fmax, is confused with the f-stop
producing the most highly resolved image. Whereas fmax
balances depth of field against loss of sharpness, the latter
only concerns the maximum sharpness as measured by the
modulation transfer functions (see Geiger 2006b for details).
Maximum sharpness is usually attained when any
photographic lens is stopped down two to three f-stops from
fully open.
The available depth of field decreases with
magnification. With high-resolution digital cameras, it is
possible to take an image at a low magnification using only a
part of the sensor and crop the image. Currently the largest
digital sensors are 16.7 MP in size. 35 mm film can be
scanned at 5400 dpi producing a 41.7 MP file. The
information content in fine grain film is still un-surpassed,
although the highest-resolution film (Kodak Tech-Pan) has
been discontinued. These theoretical calculations also omit
the resolution ability of the lenses, which are at
approximately 80 lines/mm.
Computer image processing (e.g., Automontage, Fig. 2)
allows the generation of an image with great depth of field
from a stack of images in a through-focal series, a so-called
z-stack. Although the programs can compensate for some
lateral image movement, best results are obtained if all
images in a z-stack are aligned. Specimens with many welldefined edges produce better results than those that are
featureless. Some programs are more susceptible to uneven
vertical intervals in the z-stack; motorized focus can be
helpful under certain conditions, but is usually not critical.
Usually a stack of five to nine images is sufficient to produce
a good quality combined image regardless of specimen size
and ten-20 will produce excellent results. Some of us have
had difficulties with specimens 1 mm and smaller and SEM
has been more successful in those cases. Automontage-like
programs have become routine applications to generate high
quality images. Good results are obtained in many situations
(e.g., Fig. 2; NMNZ type-collection on-line: http://
collections.tepapa.govt.nz/).
Positioning
To position dry specimens, they are usually mounted
with a slightly tacky substance (e.g., plasticine, malleable
silicone, beeswax). For transparent specimens the SEM
mounting medium Leit-C plast (Neubauer Chemikalien) is
suitable. Check that the specimen can be easily removed
from the mounting medium and that no residue is left on the
shell. For fluid immersed specimens, wax cradles, glass
slides, stainless steel nuts and pins inserted into a wax base
can be used.
Chemicals
A list of some chemicals regularly used for narcotisation,
fixation, preservation, preparation and cleaning of
micromolluscs is provided in Appendix 1. Many more
fixatives were used before biologists started using formalin
routinely (see, e.g., Romeis 1948).
The handling of chemicals requires knowledge and
experience. Clean equipment and chemicals of good quality
should always be used. For most chemicals, the CAS
(Chemical Abstracts Service numbers, http://www.cas.org)
number is provided to facilitate Internet search for further
information. Some chemicals mentioned below are regulated
by local authorities; rules and regulations vary from country
to country. Transport and importation regulations should be
carefully followed when travelling.
Acknowledgements
We thank the various technicians and lab directors at SEM
facilities, including Alicia Thompson (University of
Southern California, Los Angles, California, USA) and Sue
Lindsey, Ian Loch and Alison Miller (Australian Museum,
Sydney, Australia). Rüdiger Bieler (Field Museum of
Natural History) provided some information on
macrophotography. AW wants to thank Olle Israelsson and
Ylva Lilliemarck for information on chemistry and histology.
Mike Vendarasco and Christine Fernandez (University of
California, Santa Barbara) kindly made some of their fossil
micromolluscs available. The constructive criticism of James
McLean (Los Angeles County Museum of Natural History),
an anonymous reviewer and Jean-Claude Stahl (NMNZ)
helped to further improve this contribution. This study was in
part supported by NSF grant MRI 0402726.
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Appendix 1.
Alphabetical list of chemicals used for work on
micromolluscs. CAS numbers (Chemical Abstracts Service
numbers, http://www.cas.org) are provided to facilitate
further inquiries. List compiled by AW.
Acetic acid, CAS no 64-19-7. Acetic acid is one of the oldest
fixatives on record: in the eighteenth century vinegar
(4–10% acetic acid content) was used to preserve
hydras. It does not harden tissue; actually, it prevents
some of the hardening that, without it, might be induced
by subsequent alcohol treatment. In some techniques,
however, acetic acid must be avoided because it
dissolves certain cell inclusions, such as Golgi and
mitochondria and calcareous material. Many lipids are
miscible with acetic acid or are soluble in it. It neither
fixes nor destroys carbohydrates. Acetic acid is used as
a component of many fixatives, e.g., Bouin’s fluid. Its
usefulness lies within its fixation of nucleoproteins, i.e.,
good nuclear fixation. Acetic acid (5–50%) can be used
for decalcification.
Acetone. CAS no 67-64-1. Water-free acetone causes strong
shrinkage and is therefore not often used. It can be used
for fixation of smears and unfixed sections. For
histology, acetone is only used in combination with
other fixatives, such as formalin and sublimate.
Acetone is often used as a medium for critical point
drying, but should be avoided because of its shrinking
effect. Dry specimens directly from 100% ethanol,
which is also soluble in carbon dioxide (drying
medium). Acetone is useful for cleaning fat and
remains of glue from shells , since it is a comparatively
harmless chemical.
Amylocaine hydrochloride (Stovaine). CAS no 532-59-2.
Used for narcotisation. Slowly add 1% solution in
water, drop by drop, to 100 ml of water with animals.
For each drop give the chemical time to disperse and
animals time to react.
Benzamine compounds. Used for narcotisation. A 1–2%
solution in water is slowly added, drop by drop, to
100 ml of water with animals. For each drop give the
chemical time to disperse and the animals time to react.
Bichromate. See chromic acid.
Bleach. See sodium hypochlorite.
Borax (disodium tetraborate). CAS no 1330-96-4. (Na2B4O7
MW 201.2 and Na2B4O7, 10 H2O MW 381.4) is easily
soluble in water, non toxic and its aqueous solutions are
basic. Used for buffering formalin.
The solubility of borax in 37% and 3.7% formalin
is slightly more than 50 g /1000 ml, but varies ± 5 gram
depending on the quality of the formalin (mainly due to
concentration of methanol in the formalin).
When buffering formalin, the variety with crystal
water should be used because it dissolves much more
easily. If too high concentrations of borax are used, it
may recrystallise due to changed conditions in the
39
solution, for example temperature or concentration of
other substances. Such precipitations may be difficult to
dissolve, but warm water usually helps. One gram per
10 litres of 37% formalin will raise the pH to 7.5–7.8
and is enough for several months; do not use more than
10 gram/litre. Dilution of 37% formalin buffered in this
way to 3.7% will raise the pH about 0.9–1.0 units. For
field work it may be useful to know that one teaspoon
of borax contains 4.2 gram, one tablespoon, 11.5 gram.
Concentrated solutions of borax interfere with carbon
dioxide from the air and boric acid (H3BO3) may
precipitate, as milky clouds of very fine needles (easily
visible at 12x) in the solution. This is a result of the
equilibrium Na2B4O7 + CO2 + 6H2O <==> 4H3BO3 +
2Na+ + CO32-. Such precipitations can be avoided by
not adding more borax than necessary.
Bouin’s Fluid. A common fixative used in histology. It is a
mixture of 375 ml saturated aqueous picric acid, 125 ml
stock formaldehyde (37% w/w), 25 ml glacial acetic
acid. See under individual ingredients for hazards and
safety precautions.
Butyl acetate. CAS no 123-86-4. A good substitute for
benzene and many other unpleasant non-polar organic
solvents since it is less harmful. Hygienic limit for short
term allowable air concentration is 150 ppm =
0.11 g/m3. It has a strong smell so you are certain not to
stay anywhere with too much of it in the air. It can be
used to dissolve various organic glues (not based on
polyvinyl acetate) as well as for air-drying animals with
some dermal skeleton, such as echinoderms and insects,
with much less shrinkage. Transfer animals to butyl
acetate via 95% and 100% ethanol. Even critical point
dried specimens of soft-bodied animals can be cleaned
or removed from a stub by dissolving the glue in butyl
acetate, usually with little or no damage to the
specimen, thanks to its low surface tension.
Calcium carbonate. CAS no 471-34-1. Has a solubility
constant of 3.36 x 10-9. This means that a saturated
aqueous solution contains 5.8 mg/l, but due to
formation of hydrocarbonate ions with aerial carbon
dioxide, the solubility becomes higher with time.
Calcium carbonate (as dolomite) is used for buffering
commercial concentrated formalin, providing so-called
neutral or histology quality. These attributes, however,
cannot be stored for more than a couple of years
because of the low solubility of dolomite. After that
time the neutralizing capacity is exhausted. Calcium
and magnesium salts are not good as buffers for
specimens intended for histology because insoluble
salts have a tendency to recrystallise in tissues (Quay
1974).
Calcium phosphate. CAS no 7758-87-4. Has a solubility
therefore
constant
of
2.07
x
10-33 and
sodiumphosphates are, not very good as buffers when
fixing molluscs, since phosphate may replace carbonate
in the shells.
Carbowax. See Polyethylene glycol.
40
Carbon dioxide, CO2. CAS no 124-28-9. Used for
narcotising, by bubbling the gas through the water with
specimens or by placing animals in water saturated
with it; mainly for fresh-water organisms.
Chloral hydrate (= chloretone). CAS no 302-17-0. Soluble
in water and alcohol. Used for narcotising by slowly
adding a 0.1% solution to the animals. Chloral hydrate
is a scheduled prescription drug in some countries.
Schroll (1968) indicates that nicotine may be used as a
substitute.
Chloretone. See Chloralhydrate.
Chloroform. CAS no 67-66-3. Used for narcotising, by
sprinkling a small quantity on the surface of container
with animals. Repeat if necessary. Note that chloroform
is poisonous and flammable and is a restricted
substance in some countries.
Chromic acid (chromium trioxide). CAS no 1333-82-0.
Chromic acid and its salts, chromates or dichromates
are valuable fixatives, but the acid is considered
carcinogenic and the salts are allergenic.
Cocaine hydrochloride. CAS no 53-21-4. Used for
narcotising. Excellent for chitons and heterobranch
gastropods. Slowly add a few crystals to the container
with animals. The possession of the chemical is
generally illegal and it is difficult to obtain permits for
its use.
Cold. Many animals, especially tropical species, will die
when the temperature approaches freezing point.
Usually they do not retract to cooling so it can be
combined with the addition of some narcotising agent.
Do not allow the animals to freeze, which will destroy
most histology (unless quick frozen, e.g. in liquid
nitrogen). Freezing is suitable for DNA sequencing
work.
Cyanoacrylate, methyl. CAS 187-05-3, super glue. Can be
used for repairing small, broken shells, but only in
cases where two pieces need to be glued together and
you can do it without much adjustment, since you only
have a second for this. If using it on a scale larger than
milligram, consult safety information since it is highly
toxic, much more than formaldehyde, but also more
treacherous since its smell is less deterring. It glues by
polymerisation induced by moisture, and sticks to skin
and any other tissue.
Dichromates. Fixatives. See chromic acid.
Diethyl ether. CAS no 60-29-7. Used for narcotising, by
sprinkling a small quantity on the surface of container
with animals. Repeat if necessary. A good solvent for
many organic compounds but due to the tendency to
form explosive peroxides it should be stored in a dark
bottle, preferably cold. It is highly flammable.
EDTA (Ethylene diamine tetra acetic acid). CAS no 60-00-4.
A 0.1 M (MW = 292.24) solution has been used for
narcotising purposes. Also used for decalcifying when
development of carbon dioxide may rupture tissues.
The process is very slow, up to several weeks. For
molluscan shells use 1–5% hydrochloric acid in 80%
ethanol instead. It takes a few minutes to a day only and
leaves the tissues in very good condition. EDTA is a
permitted food additive in small quantities, and not very
poisonous.
Ethanol. CAS 67-17-5. Has been used for storage of
biological material for more than 300 years (Boyle
1666; Waller and Strang 1996). It is by far the best
storage medium for material for general use and the
only one for long term use (Jones and Owen 1987: 60;
Levi 1966).
Avoid denatured ethanol for museum storage; the
denaturants will accumulate as the ethanol evaporates
and the jars are refilled. Many denaturants have severe
side effects on the stored material, ketones and
aldehydes react with the tissues; finally the poor human
that has to work with the denatured ethanol is affected
as is the intention with the denaturation. Some
denaturants
(methanol,
glycerol,
isopropanol,
aldehydes) contribute to dissolution of micro shells by
forming complex ions with calcium. Most governments
accept use of tax-free ethanol for museum purposes
although the bureaucracy may be intimidating. Ethanol
is only sold as >99.5% solution and no manufacturer
guarantees 100% concentration because ethanol is
hygroscopic. When we refer here to 100% ethanol, we
indicate the purest form available. Hygroscopic beads
may be added to ultrapure ethanol, which will bind
excess water in the ethanol.
Micro shells may be affected by storage in
ethanol. This is not caused by acidity and buffering
does not help. (Actually you cannot even properly
measure the pH in alcohol since the ion product of [H +]
x [OH-] is no longer 10-14.) The reason for the
dissolving power is formation of a complex ion. A
calcium ion surrounded by five ethanol molecules is
slightly water soluble and the calcium no longer stays
precipitated as calcium carbonate. Since the complex
ion is water soluble, this effect can be reduced by
storing in 80% ethanol instead of the usual 70%. When
stored in 95% the effect is not noticeable (an advantage
with saving specimens for DNA (Carter 2002)), but
regrettably the specimens become brittle and less useful
for anatomy. At SMNH 80% ethanol is used as standard
for this reason. Presence of other organic compounds
like fat can give the same result, which is why tubes
with micromolluscs should not be stored with large
specimens.
Ethanol may be used for narcotising animal by
slowly adding it to the animals.
Ethanol is flammable and ignites at 363°C. Its
flash point is 13°C and it forms explosive mixtures with
air. The density of ethanol vapour is 2.1 g/l, 1.6 times
that of air, which means that you can walk in explosive
concentrations without smelling it, since it is
accumulated along the floor. The upper hygienic limit is
usually given as 1000–5000 ppm (1.9–3.8 g/m3). It can
be smelled already at 10 ppm (0.02 g/m3). Even
41
working in the highest allowed concentration does not,
contrary to common belief, cause intoxication by
absorption of ethanol via the lungs (Travis 2001). The
minimum lethal dose known for an adult human
corresponds to 100 grams of pure ethanol. See USI
Chemicals (1981) for details.
Be sure to know the difference in concentration
between percent by volume and percent by weight;
normally when biologists talk about 95% or 70%
ethanol, it is by volume. Tables for preparation of
different concentrations can be found in most histology
text books, e.g., Romeis (1989) but for simplicity see
Tables 1 and 2.
TABLE 1. Dilution of 1000 ml water to produce a certain strength of ethanol (Lide 1997). Based on Romeis (1989) and Lide (1997).
Required
strength vol %
Ml 95%
ethanol
Corresponds to
weight %
Add ml water
(Romeis 1989)
Density
(Lide 1997)
Total
volume
95%
1000
0.808
90%
1000
86%
64.1
0.8284
1047
85%
1000
80%
133.3
0.8436
1107
80%
1000
74%
209.5
0.8581
1179
75%
1000
68%
295.2
0.8724
1265
70%
1000
62%
381.5
0.8865
1342
65%
1000
58%
502.2
0.8958
1460
60%
1000
52%
630
0.9095
1584
55%
1000
47%
779.9
0.9205
1730
50%
1000
42%
958.9
0.9311
1906
.
TABLE 2. Dilution of ethanol to produce 1000 ml solution of a desired strength.
Required vol
Required strength
ml 95% ethanol
ml water
1000
by volume
1000
90%
955
61.2
1000
85%
903
120.4
1000
80%
848
177
1000
75%
790
233
1000
70%
744
284
1000
65%
684
344
1000
60%
631
398
1000
55%
578
451
1000
50%
521
507
Ethylene glycol. CAS no 107-21-1. A 50% water solution is
sometimes used for storage (Lincoln and Sheals 1979:
136), but should be avoided for sensitive molluscs,
because it speeds up dissolution of shells.
Eucaine. See Benzamine compounds.
Formaldehyde. CAS no 50-00-0. A water-soluble (max.
52%) gas, commercially sold as a 35–50% water
solution called formalin. For general information see
Anonymous (2006b). It is strongly recommended to
abandon the poorly founded practice to call the
commercial solution ‘100% formalin’ and the 3.5–4%
solution normally used for fixation ‘10% formalin’
(e.g., Pritchard and Kruse 1982; Simmons 1991). The
reasons for this are the variation in strength of the
commercial solution, and the fact that this is against all
normal practice for other chemicals.
Since the late 1800s formalin has been routinely
used in zoology for fixation of animal tissues. Fixation
takes place by chemically forming links between nearby
protein chains. The optimal pH range for this reaction is
7.5–8.0. Formalin should never be used unbuffered
(Presnell and Schreibman 1997: 21).
Formaldehyde and its aqueous solutions are
poisonous and highly irritating to the eyes, nose and
42
throat, even at very low concentrations. It is
carcinogenic and regular handling frequently causes
allergy (short term exposure maximum concentration
1 ppm). For some people allergic reactions may appear
after only a short time, for others it never happens, but
great caution should always be taken to avoid contact
with the fluid or inhaling the vapour. Preserved material
and the solution should always be stored in ventilated
cabinets or a fume hood, never in a closed cabinet. If
stored enclosed, formaldehyde vapour will accumulate
and the concentration may rise to dangerous levels due
to evaporation.
It is fairly cheap and simple to test laboratories
and storage space for presence of formaldehyde in the
air and the test has sensitivity of less than one
thousandth of the maximum allowed concentration. Test
badges (GMD systems 570 series), available at http://
www.scottinstruments.com/products/product_list.cfm
or their local representative are placed at various
locations, a protective cap is removed, the badge is left
in place for 24 hours then resealed and sent to a certified
laboratory for analysis. Quantities down to less than
0.001 ppm are registered. Few (or none) of people in
charge of collecting or collections seem to have
accepted the warnings of Simmons (1991) about the use
of formaldehyde, although many of his statements
about formaldehyde are obviously wrong or
exaggerations.
Addition of a small quantity of ammonia to
formalin fixed specimens has been used to remove the
smell, but the procedure is based on the formation of
formamide (HCONH2, CAS number 75-12-7) and
hexamethylene tetramine (CAS number 100-97-0)
which are odourless, but the practice is deceptive since
the formalin smell is a good warning to improve
ventilation or do the work in a more suitable place. For
similar reasons, decalcification of large specimens in
formalin with hydrochloric acid must be done in a fume
hood because of the formation of phosgene (carbonyl
chloride, COCl2 CAS no 75-44-5), also highly
poisonous (threshold limit for allowable air
concentration 0.05 ppm.). Therefore, formalin fixed
specimens must be well rinsed before they are
processed.
Many different qualities of formalin are available
on the market. The cheap qualities are fully functional
for normal fixation if used only for a short time and
adequately buffered. Even so-called acid-free qualities
which are buffered by addition of dolomite (calcium
carbonate) change their pH after one or two years
because the solution cannot dissolve enough dolomite
to buffer for a long time.
Formation
of
Paraldehyde
and
Paraformaldehyde [(CH2O)n (n = 6–50)] takes place
when formalin is stored at temperatures below +5–
10°C; the conversion is faster at lower temperature,
high concentration of formaldehyde and high pH. The
polymer is insoluble in water, ethanol, xylene, acetone,
but can be destroyed by treatment with bleach. If such
deposits have precipitated on shells, it is usually
possible to get rid of them by soaking them in warm
water and brushing off the deposits. It is said that the
polymerised form can be dissolved by autoclaving.
Polymerisation may also occur directly if formalin is
allowed to evaporate, so it is important to rinse formalin
preserved specimens intended to be dried.
One advantage with formaldehyde fixation of
specimens intended for dry collections is that they are
much less likely to be eaten by insects. Specimens
preserved only in alcohol are as likely to be attacked by
insect as those simply dried without any fixation
(common with land snails).
The commercial solution of formaldehyde usually
contains 5–20% methanol, to prevent generation of
formic acid by the so-called ‘Cannizzaro reaction’,
where an aldehyde produces equal amounts of the
corresponding alcohol and acid. This lowers the pH
towards 3 in old, low grade formalin. This can lead to
destruction of sensitive shells in less than a day. To
avoid this the formalin should be buffered, preferably
with sodium tetraborate, which is a nontoxic, cheap and
stable substance. This neutralizes the formic acid and
raises the pH to ca 7.5–8.0 in 40% and to 8.0–9.2 in 4%
formalin.
The solubility of borax in formalin (37% and
3.7%) is slightly more than 50 g/litre at room
temperature (lower at lower temperatures). Addition of
this much borax, however, will increase the risk of
formation of paraldehyde and (1 tablespoon) 10 g / litre
37% formalin is more than enough to ensure a stable pH
above 7.5 for two years.
Prolonged storage of organic tissues in borax
buffered formalin must be avoided because of the onset
of histolysis at this high pH, but this need not be
considered for several months.
To measure pH in formalin is simple and may be
done with indicator paper (preferably special types for a
narrow range, of which several are available, for
example pH 6.0–10.0) or with an electric pH meter.
A general conclusion of this and existing literature
is that it is not advisable to store, only to fix, molluscs or
any animals with a calcareous skeleton in formalin. The
formalin needs buffering, at least to a pH of 7.5–8.0 to
prevent damage of calcium carbonate. At this high pH
there is a risk of polymerisation. For tissue samples a
storage at the isoelectric point of proteins, pH 6–7, is
considered advantageous (Steedman 1976) because at
this level they have the minimum of solubility. At this
low pH, however, the protein chains are believed to
become more brittle and anatomical details, from cilia
to legs, break off more easily (Steedman 1976). More
information on formalin can be found in Carter (1997).
Formic acid. CAS no 64–18-6. Formic acid is often added to
fixatives for combined fixation and decalcification.
Gooding and Stewart's fluid is an aqueous mixture of
formic acid and formalin used for decalcification. It is
quite a bit slower than nitric and hydrochloric acid, but
tissues can be left in it for much longer. After 5 or 6
days, good nuclear staining with alum hematoxylin can
still be obtained.
Glycerol. CAS no 56-81-5. Used for storage at a
concentration of 50%, mainly of histological specimens
(Horie 1989:24; Presnell and Schreibman 1997: 331).
However it is sticky and difficult to clean. Addition of a
small amount (ca 5%) to ethanol preserved specimens
has been used and is said to reduce the hardening effects
of ethanol, at the same time it may save the specimens if
the jar should dry out. The reason for this is that
glycerol evaporates much more slowly and will keep
the specimens moist for a long time after the ethanol is
gone. There is, however, a risk that mould may start
growing on the remains before the alcohol is totally
gone (Jones and Owen 1987). Naticid egg collars can be
immersed in glycerol and then ‘dry’-stored without
becoming brittle. Glycerol speeds up dissolution of
calcareous material and must be avoided for storage of
small molluscs. Since it is a complex formation this is
true also for small quantities. (Threshold limit for
allowable air concentration 50 ppm; MW 92.09, boiling
point 290 °C, vapour pressure <0.01 kPa, flash pt.
199°C, ignition temp. 370°C).
Heat. Some animals will slowly die from exposure to
supernormal heat. Be careful; often the animals produce
large quantities of mucus and the histology of epithelia
is destroyed. Another application is to drop small
animals into as hot fixative as possible, e.g., boiling
chromic acid or formalin (Gohar 1937). The diameter of
the thickest part of the animal should not exceed a
centimetre. If larger, the animals will have time to
contract before fixation. The fixation, however, is
usually excellent. A less noxious variety of this method
is to immerse specimens in 70–100ºC water for a few
seconds, or up to a minute for large specimens. This
will detach columellar and adductor muscles from the
shell and make it possible to ‘unscrew’ the soft parts of
gastropods enough to ensure unhampered access of the
fixative. If preserved in ethanol afterwards, the
specimens can still be used for DNA. A spring-loaded
tea strainer is excellent for keeping the specimens under
control.
Hexamethyl disilizane = HMDS. CAS no 999-97-3. It is
used as a intermediate solvent to desiccate tissues
without shrinkage, particularly for SEM applications.
Tissue is brought to 100% EM-grade ethanol through at
least three changes of 100% ethanol. After three
changes of HMDS, the animal is slowly air-dried
overnight in a loosely covered dish. The last change of
ethanol should be carried out shortly before transfer to
HMDS because ethanol is hygroscopic (attracts water)
and trace amounts of water will produce inferior results
later.
HMDS has a high vapour pressure, is waterreactive, foul smelling and poisonous. Use only in well-
43
ventilated areas and use gloves when handling it.
Consult MSDS before use.
Hexamethylene tetramine. CAS no 100-97-0. White
crystalline powder. Jones and Owen (1987: 58)
recommend 200 g/litre of 40% formalin, for buffering
collections stored in formalin. At a price five times that
of borax and a recommended use of 10–20 times that of
borax, this is relatively expensive. In addition, the
substance is a skin-, eye- and respiratory irritant and is
flammable.
Hydrochloric acid. CAS no 7647-01-0. Sold as a 35–38%
solution (concentrated) of hydrochlorine. Develops
fumes and heat when diluted. Diluted (3–5%), it is good
for decalcifying tissues if development of carbon
dioxide does not rupture the tissues. Such damage can
be reduced by using a mixture of one part concentrated
acid and nine parts of 80–95% ethanol.
Hydrogen peroxide. CAS no 7722-84-1. Hydrogen
peroxide is a good substitute for bleach in many cases,
since it is less damaging. It can be used for cleaning
shells and radulae, also shells with nacre and it does not
damage the periostracum as much as bleach. Use it at a
strength of 1–10% and render it slightly alkaline by
adding a trace of sodium hydroxide. 30% Hydrogen
peroxide can be used as stock solution and must be
stored cold; stronger hydrogen peroxide should be
avoided since it can decompose explosively.
Isopropanol. CAS no 67-63-0. At a concentration of 40–
60% it is used for storage of zoological material
because it is not taxed like ethanol (Wagstaffe and
Fidler 1955: 173; Pearse 1960). Storage in isopropanol
makes specimens unsuitable for most histological work
(Jones and Owen 1987 and references therein).
Isopropanol speeds up dissolution of calcareous
material and must be avoided for small mollusc shells.
Since it is a complex formation this is true also for the
small quantities used in some denaturations of ethanol.
Lithium salts. Schroll (1968) lists 2–4% lithium solution as
a narcotic for molluscs.
Magnesium chloride. CAS no 7791-18-6. At a
concentration of 72.3 gram per litre it is isotonic with
(3.5%) sea-water. This is an excellent tranquilliser for
marine animals (Tullberg 1891). It works by changing
the magnesium-calcium balance of the motor end-plates
of the nerves, inhibiting the transfer of impulses to the
muscles. Magnesium chloride is added slowly; start
with 1/10 of the sea-water volume and check if the
animals retract. A few minutes later add another tenth.
Continue with larger doses. It may be necessary to drain
a part of the fluid with the animals. The process is
reversible. By transferring the specimens to fresh seawater the animals will recover and the process can be
started over again if the animals get disturbed and
retract. See also magnesium sulphate.
Magnesium chloride is a fairly harmless and cheap
substance. Ingestion of several grams causes diarrhoea.
It is sold as a fairly expensive anhydrous crystal, or as
the cheaper hexahydrate on which the weight above is
44
based.
Magnesium sulphate (Epsom salts). CAS no 10034-99-8.
(MgSO4 x 7H2O, MW = 246.48). Sometimes used
instead of magnesium chloride. The chloride is easier to
handle since it is less hygroscopic and requires smaller
amounts (131.5 g/litre of the sulphate). The
physiological mechanism, blocking synapses, is the
same, as is the procedure. Smaldon and Lee (1979)
presented 12 variations for the use of magnesium
sulphate and chloride, for use with marine invertebrates,
concentration between 0.1% and 20%.
Menthol. CAS no 2216-51-5. Used for narcotising mainly
fresh-water molluscs, by slowly adding a solution in
ethanol or a few crystals to the surface of the water.
May be irritating for skin, eyes, or respiratory organs,
but is not considered very dangerous. Smells of
peppermint.
Mercury chloride. CAS no 7487-94-7 (sublimate). A
component in many good fixatives, e.g., sublimate
alcohol (Romeis 1989). Due to mercury being a severe
pollutant and its toxicity, even at skin contact, these
fixatives are less in use nowadays. Residues must not be
discarded, but saved and labelled for special treatment
according to local laws.
Methanol. CAS no 67-56-1. Used for denaturing (5–20%)
ethanol, stabilizing formalin (1–20%, by the reaction 2
CH2O + H2O ⇔ CHOOH + CH3OH) and storage of
biological specimens (Horie 1989: 21). Its other
drawbacks are that it is poisonous (threshold limit for
allowable air concentration 200 ppm) and speeds up
dissolution of calcareous material.
MS 222TM (Tricaine methanesulphonate). CAS no 896-6-2.
Used for narcotising, usually fish, but also
invertebrates.
http://www.argent-labs.com/
argentwebsite/ms-222.htm. Has the advantage of giving
reversible narcotisation and may be used on fish
intended for later consumption (FDA approved). For
invertebrates, slowly add some crystals to a few ml of
water.
Nembutal (= sodium pentobarbitone). CAS no 57-33-0.
Used for narcotising by slowly and repeatedly adding a
5% solution. Test 1 ml solution per 100 ml sea water.
Osmium tetroxide. CAS no 20816-12-0. Used for
stabilising and contrasting tissues to be used for TEM
and SEM of critical point dried specimens, when very
high resolution is needed. It has not been found
necessary for SEM of critical-point dried specimens,
even at 10.000 times magnification.
Osmium tetroxide should always be handled with
utmost care, used in a well-ventilated area (under a
fume hood) and special care should be taken to avoid
eye and nasal contact. Osmium tetroxide vapours will
react with any proteins, including the cornea of the
human eye, where black deposits may be formed. The
solution penetrates poorly (maximum 1 mm) and leaves
the tissue soft and difficult to use for wax sectioning.
When fixation is complete, excess osmium tetroxide
must be washed out of the tissue or it will reduce to an
insoluble precipitation of metallic osmium during
treatment in ethanol.
Paraformaldehyde.
CAS
no
30525-89-4.
See
Formaldehyde.
Phenoxetol® (propylene phenoxetol = phenoxy isopropanol
= ‘nipa ester’ = B-phenoxyethylalcohol = propylene
glycol monophenyl ether). CAS no 122-99-6. Used as
1–2% solution, often with 2–5% propylene glycol to
dissolve it more easily, for storage of zoological
specimens (Lincoln and Sheals 1979: 136; McKay and
Hartzband 1970; Mahoney 1973; Steinmann et al.
1975). Phenoxetol requires heavy fixation a priori.
Phenoxetol is commercially used in disinfectants,
preservatives and cosmetics. It is not considered
harmful. More information at www.clariant.com.
After having poured out several squids so stored
30 years ago (after proper fixation), I am not fond of the
method. Propylene phenoxetol seems more promising
as a tranquilliser and has been used by slowly adding a
0.5–2% solution to sea-water.
Picric acid. CAS no 88-89-1. Picric acid is an excellent
protein coagulant, forming protein picrates that have
strong affinity for acid dyes. However, it penetrates
slowly, causes extreme shrinkage and offers no
protection against subsequent shrinkage. It is used in
Bouin’s fluid (see above).
Picric acid crystals are explosive, but need a heavy
shock to explode. However, salts with heavy metals,
e.g., iron, are shock sensitive and heavy metals must not
come in contact with picric acid. To store it more safely,
it is commercially handled in water. Be sure your
supply has not dried and top it up with water if
necessary.
Polyvinyl acetate. CAS no 9003-20-7. Together with
polyvinyl alcohol CAS no 9002-89-5 this forms the
basis for many brands of glues used for wood and paper.
They are usually white and may be thinned with water.
A dry surface of such glue is good for mounting wet
radulae and other small wet objects, where the moisture
will soak the surface enough to make it sticky. Do not
try to mount small objects in an excess of glue; the glue
will use every possible crack to soak your specimen by
capillary forces. Small dots of glue are good for
opercula on a SEM stub, but will corrode small shells.
For the same reason, an organic based glue should be
used for repairing shells, not PVA glues which are too
acidic and will cause damage. Use glues based on
nitrocellulose or other polymer dissolved in acetone,
ethyl acetate, butyl acetate or similar solvents.
Potassium hydroxide. CAS no 1310-58-3. Used for radular
preparation and maceration of tissues. Be sure to use
analytical grade to avoid unwanted precipitations of
impurities. Potassium hydroxide is said to be less
hygroscopic than sodium hydroxide, which is an
advantage, as only small quantities are needed and a jar
lasts for a long time. The quality with pellets is
preferable since it has a smaller surface and therefore
reacts more slowly with carbon dioxide from the air.
Prepare the solution directly in the vessel to be used by
adding a few pellets or a quarter of a pellet when small
quantities of water, e.g. 2–6 droplets, are used.
Potassium- and sodium hydroxide are highly
corrosive and all tools that have been in contact with
them should be cleaned as soon as possible. Even in
small quantities it is harmful on the skin, starting with
redness and a burning sensation. It destroys clothing.
Rinse with lots of luke-warm water, even with 1–2%
acetic acid or vinegar, then further washing. A splash in
the eye should be rinsed immediately with lukewarm
water for several minutes; then obtain medical
attention. Strong hydroxides quickly dissolve fat, which
is the main reason why strong hydroxides are at least as
dangerous as strong acids. It is a good preventive
measure to apply hand lotion before preparing radulae,
to reduce the effect of small splashes.
Dissolution of potassium and sodium hydroxide in
water develops enough heat to cause explosive boiling
if large quantities are dissolved in a narrow vessel.
Never dissolve it in warm water!
Propylene phenoxyethanol. See phenoxetol®.
Propylene glycol. CAS no 107-21-1. Can generally be used
instead of ethylene glycol (Presnell and Schreibman
1997: 9), but is much more expensive. Propylene glycol
is sometimes added (2–5%) to ethanol preserved
specimens because it is assumed to reduce brittleness
(Boase and Waller 1994). It contributes to rapid
dissolution of micromolluscs.
Sodium hydroxide. CAS no 1310-73-2. See potassium
hydroxide, which is used for similar purposes and
presents the same hazards.
Sodium hypochlorite. CAS no 7681-52-9, commercial
bleach. Commercial bleach is a solution of sodium
hypochlorite in water, usually with some silicates added
for preventing suspended particles sinking to the
bottom. Often perfume is added to conceal the smell of
chlorine. Due to the presence of the silicates, all
equipment, shells and radulae must be thoroughly
rinsed; otherwise the silicates will form insoluble
precipitations.
Commercial bleach, diluted with 1–5 times as
much water, is a good oxidant for destroying organic
dirt. Weak heating (30–50°C) speeds up the process. At
the same time the high pH facilitates removal of the
dirt. During the process chlorine (gas) is produced,
causing a distinctive, unpleasant smell. Working with
millilitres of the solution this does not present a danger.
Be careful not to get splashes in your eyes; if this
happens thoroughly rinse with lukewarm water; get
medical attention if problems remain. Read instructions
on bottle.
Sodium lauryl sulphate. CAS no 151-21-3 (=sodium
45
dodecyl sulphate). Used as a 5–50% solution for
dissolving tissues and cleaning inorganic material. Can
be used for radular preparation when potassium
hydroxide
disintegrates
the
radula,
e.g.,
patellogastropods and chitons. This is a longer
procedure and takes up to a week at 30–50°C. Much
tissue remains afterwards and cleaning with a fine
paintbrush in warm water and a short (a few seconds)
rinse with bleach before the final rinsing is
recommended. Due to the dissolution of fat and cell
walls it should be handled with care.
Sodium salts. Sodium phosphates are not good for buffer
use in combination with sea water or calcareous
material since the calcium in sea water will precipitate
as phosphate, or recrystallisation of calcareous tissues
may take place. Sodium salts of organic acids seem
poor as buffers for protection of calcareous elements
since the organic ions often form complex ions (even
chelates) with calcium ions. The carbonates seem
acceptable (see below), but the resulting pH of sodium
bicarbonate alone is high enough to precipitate
paraldehyde in formalin.
Sodium carbonate. CAS no 497-19-8. MW 105.989 (also
deca- and monohydrates exist) and sodium hydrogen
carbonate, CAS no 144-55-8, sodium bicarbonate,
(MW 84.007) have been used extensively for buffering
formaldehyde and give a pH of 8 and 11 respectively in
a 0.25M solution over a rather wide range of
concentrations.
In an aqueous solution 21 g/l (= 0.25 M) NaHCO3
has a pH of 8.0; 26 g/l (= 0.25 M) Na2CO3 gives a pH of
11.4. In formalin, however, a range of mixtures
carbonate:bicarbonate 1:10–1:1 produce a pH of 9–10,
which soon starts precipitation of paraformldehyde and
means that sodium carbonate – hydrogen carbonate
buffers should be avoided for buffering formalin.
Sublimate. See mercury chloride.
Superglue. See cyanoacrylate.
Urethane (ethyl urethane, ethyl carbamate). CAS no 51-796. Used for narcotisation by adding 1% solution of
urethane in sea water. (Urethane is actually the name of
the whole group of carbamates. For use as narcotising
agent, the ethyl ester is usually employed: Dudich and
Kesselyák n.d.). Not recommended since it is classified
as a carcinogen and easily replaced by less dangerous
compounds.
Water. Freshwater is often recommended for narcotising
marine animals, especially for echinoderms. Slowly
(1/10 per minute) add it to animals in sea water until
they stop reacting. Leave them for 5–10 minutes and
fix. This method destroys all epithelia and is suitable for
animals saved for identification only. Water used for
washing material for use with the SEM should
preferably be distilled.
46
Index Micromollusks
Pages in bold refer to illustrations.
3D, see three dimensional
Acetic acid ......................................................................... 16, 24, 39
Acetone ........................................................................ 21, 39, 42, 44
Acid........................................................................ 11–12, 15–16, 18
Acid free paper, see Paper, types of Adhesive, see Carbon tab,
Colloidal graphite, Glue, Leit-C plast, Silver paste
Air bubble ...................................................................................... 17
Air dry................................................................................ 14, 17, 43
Air-lift pump .................................................................................. 10
Aldehyde, see also glutaraldehyde...........................................40, 42
Algae .......................................................................................... 9, 11
Allergene............................................................................ 13, 40, 42
Amines ........................................................................................... 27
Amira .......................................................................................32, 33
Ammonia........................................................................................ 12
Amylocaine hydrochloride............................................................. 39
Anatomy, external .............................................................. 22, 24, 32
Animals, see SEM, animals; live animals
Anoxic............................................................................................ 10
Aragonite........................................................................................ 12
Archival paper, see Paper, types of
Arkanas Stone ..................................................................................8
Automontage.............................................................................. 4, 35
Bag, cloth ....................................................................................... 10
Bag, heat seal ................................................................................. 13
Bag, zip lock ....................................................................................9
Bait................................................................................................. 10
Barbiturates .................................................................................... 11
Basket............................................................................................. 10
Beeswax ......................................................................................... 35
Bench grinder...................................................................................8
Bench vice...................................................................................... 12
Benzamine...................................................................................... 39
Bichromate, see Chromic acid
Biodiversity..............................................................................2, 3–5
Bivalve .......................................................................7–8, 17, 21–22
Bivalve, opening ............................................................................ 21
Bleach ........................................................ 17, 21, 26–27, 29, 43, 45
Blow dry......................................................................................... 14
B-phenoxyethylalcohol, see Phenoxetol
Boiling method................................................................... 11, 13, 43
Borax............................................................................ 12, 39, 42–43
Bouin’s fixative ............................................................ 12, 24, 39, 44
Bowls, see also Embryo bowls ........................................................9
Box, plastic .................................................................................... 15
Box, polystyrene ............................................................................ 15
Brush ....................................................5, 6, 8, 10, 17, 21, 28–30, 32
Brush, dry....................................................................................... 14
Brush, moist ............................................................................. 11, 14
Buccal mass ................................................................................... 16
Buffer, see Paper, types of; Formalin, buffered
Butric acid ...................................................................................... 16
Butyl acetate....................................................................... 21, 39, 44
Byne’s disease .................................................................... 13, 15, 16
Byssus.........................................................................................9–10
Caenogastropoda ............................................................................29
Calcium acetate ..............................................................................16
Calcium carbonate, calcite .........................12–13, 15–16, 39–40, 42
Calcium phosophate .......................................................................39
Capillary forces ........................................................9, 17–18, 30, 32
Carbonate, biogenic..........................................................................9
Carbon dioxide .......................................................24, 26–27, 39–40
Carbon tab/tape, double-sided..............17, 18, 20, 21, 24, 29–30, 31
Carbon tetrachloride .......................................................................16
Carbonyl chloride ...........................................................................42
Carborundum paper ..........................................................................8
Carbowax, see Polyethylene glycol
Carcinogen..............................................................12–13, 40, 42, 45
Cardboard slide, see geology micromounts
Celluloid .........................................................................................14
Charging, see SEM, charging
Chemicals, see also under particular compound ................35, 39–45
Chiton, see Polyplacophora
Chloral hydrate, chloretone ............................................................40
Chlorine gas....................................................................................45
Chloroform ...............................................................................21, 40
Chromic acid, .....................................................................12, 40, 43
Clam, see bivalve
Clay, plastic ....................................................................................15
Cleaning....................................................................................16–17
Cocaine hydrochloride ...................................................................40
Cold, see also Freezer, freezing..........................................10–11, 40
Collecting ...................................................................................9–10
Colloidal graphite ...............................................................17–18, 21
Concavity slide, see depression slide
Conchiolin ................................................................................17, 26
Conductive wire .............................................................................18
Conductivity, electrical.............................................................17–18
Container for specimens ...........................................................14–15
Cork ................................................................................................16
Cotton wool ........................................................................13–16, 24
Cover slip ...........................................................................29–30, 32
Critical point drying, CPD......................................18, 24, 25, 39, 44
Cyanoacrylate .................................................................................40
Decalcification........................................................24, 39–40, 42–43
Dehydration ..............................................................................24, 43
Depression slide ...................................................................9, 27, 30
Detergent ......................................................................10, 17, 21–22
Diethyl ether .............................................................................11, 40
Digital imaging.........................................................................32–35
Dirt......................................................................................14, 17–18
Dish ................................................................................................11
Disodium tetraborate, see Borax
Dissecting microscope, see stereomicroscope
Dissection .........................................................................2, 8, 24, 26
Distortion..................................................................................22, 34
DNA extraction ..................................................................26–27, 28
Docoglossate ............................................................................30, 32
Dolomite ...................................................................................39, 42
Dredge ........................................................................................9–10
Drill ......................................................................................8, 12, 23
Dry bath incubator..........................................................................27
Drying ............................................................................................ 14
Drying chamber ............................................................................. 14
Efflorescence...................................................................... 13, 15–16
Elutriation ...................................................................................... 11
Embryo bowl........................................................................9, 17, 27
Epibenthic sledge ....................................................................... 9–10
Eppendorf tubes .......................................................................13, 27
Epsom salt, see Magnesium sulphate
Ethanol ....................................................... 11, 17, 28–30, 40–43, 45
Ethanol 30%............................................................................. 11–12
Ethanol 50–60%.......................................................................12–13
Ethanol 75–80%............................................... 11–14, 21, 28, 40, 43
Ethanol 95%........................................................... 11–14, 21, 40, 43
Ethanol 100%............................................. 11–14, 21, 24, 39–40, 43
Ethanol vapour ......................................................................... 12, 43
Ethanol, vodka ............................................................................... 22
Ethyl acetate................................................................................... 44
Ethyl carbamate/urethane, see Urethane
Ethylene diamino-tetraacetic acid, EDTA ...............................24, 40
Ethylene glycol ........................................................................41, 45
Eucaine........................................................................................... 41
Falcon tubes ................................................................................... 13
Files..................................................................................................8
Fixation, fixative .............................................. 11–12, 39, 41, 43–44
Floating off..................................................................................... 11
Flocculent material......................................................................... 27
Foam, plastic .................................................................................. 15
Forceps ........................................ 6, 7, 11–12, 14, 21, 24, 27–28, 30
Formaldehyde ..........................................................................41–42
Formalin ..................................................... 10–13, 22, 24, 27, 41–45
Formalin, 37–40%.............................................................. 12, 39, 41
Formalin, buffered ........................................... 12–13, 22, 39, 41–43
Formalin – seawater.................................................................12–13
Formamide ..................................................................................... 42
Formiate ......................................................................................... 16
Formic acid ........................................................................ 24, 42–43
Freshwater............................................................................ 2, 10, 45
Fridge, see also Cold................................................................ 10–11
Freeze drying ................................................................................. 24
Freezer, freezing, see also Cold ...............................................10–12
Funnel ............................................................................................ 14
Gastropod ......................................................... 17, 21–22, 26, 29, 40
Gelatine capsule ............................................................................. 14
Geology micromounts.................................................................... 14
Glass disease ...................................................................... 13, 15, 16
Glass, types ........................................................................ 13–14, 16
Glue................................................................................................ 24
Glue, polyvinyl acetate .................................... 18, 24, 29–30, 32, 44
Glue, spray ...............................................................................18, 21
Glue, super see Cyanoacrylate
Glutaraldehyde................................................................... 11–12, 22
Glycerol.................................................................................... 40, 43
Gooding and Stewart’s fluid .......................................................... 42
Grab sampler.............................................................................. 9–10
Habitat.................................................................................... 2, 9–10
Hairs .....................................................................................8, 30, 32
Hammer.......................................................................................... 12
Handling, see Specimen manipulation
47
Hazards .........................................................2, 11, 13, 24, 35, 39–45
Heat ..................................................................10, 13, 26–28, 30, 43
Hetrobranchia .................................................................................40
Hexamethyldisilizane, HMDS..................................................24, 43
Hexamine, hexamethylene tetramine .................................12, 42–43
Histology ....................................2, 10, 12, 24, 32, 33, 39–40, 43–44
Histolysis ..................................................................................12, 42
Holotype, see Specimen, types
Humidity.............................................................................14–16, 32
Hydrochloric acid ...................................................15, 24, 40, 42–43
Hydrogen peroxide .............................................................17, 21, 43
Hypochlorite, see Bleach
Illumination, see Lighting
Incubator, see also Dry bath incubator ...............................27, 29–30
Insects .............................................................................................42
Irritant ...............................................................................................9
Irwin loop .......................................................................................11
Isopropanol.........................................................................21, 40, 43
Ketone ............................................................................................40
KOH, see potassium hydroxide
Labels, see also paper ...............................................................14–15
Land snail, see Pulmonate
Laser writer.....................................................................................15
Lateromarginal plate.......................................................................32
Leaf litter ....................................................................................9–10
Leit-C plast .....................................................................................35
Lens, photographic ...................................................................32–33
Lens, microscope............................................................................34
Lepetelloidea ..................................................................................26
Light sources ..............................................................................7, 28
Lighting ........................................................................28, 30, 34–35
Light microscope ......................................................................29–30
Light trap ........................................................................................10
Limpet, see Patellogastropoda
Lithium .....................................................................................11, 43
Live animals ...............................................................................9, 11
Maceration......................................................................................27
Magnesium chloride .....................................................10–11, 43–44
Magnesium sulphate.......................................................................44
Mangrove..........................................................................................9
Manipulation, see Specimen manipulation
Marine ..............................................................................................2
Menthol ....................................................................................11, 44
Mercury ..........................................................................................12
Mercury chloride ......................................................................39, 44
Metal.........................................................................................14, 27
Methanol.............................................................................40, 42, 44
Methenamine, see hexamine
Methods, failed .............................................2, 12–13, 18, 24, 26–27
Method, best .....................................................................................2
Microfossil slides, see geology micromounts
Micromount, mineral......................................................................15
Microscissors....................................................................................7
Microscope, see stereomicroscope
Microscope, light, see Light microscope
Microwave......................................................................................15
Molecular work ......................................................12–13, 26, 40, 43
Monoplacophora.............................................................................26
48
Mother of pearl, see nacre
Mould .......................................................................................14, 43
Mountants, tackless picture............................................................ 15
MS222...................................................................................... 11, 44
MSDS...............................................................................................2
Mucus.......................................................................................14, 43
Mylar.............................................................................................. 15
Nacre ........................................................................................17, 21
NaOH, see sodium hydroxide
Narcotisation .................................................. 2, 9–11, 39–40, 43–45
Needle holder ...............................................................................7–8
Needle, see also Pin .....................................8, 21–22, 24, 27–30, 32
Needle, tungsten......................................................................... 8, 30
Nembutal........................................................................................ 44
NEM tape ....................................................................................... 18
Neogastropoda ......................................................................... 29, 32
Nicotine, see also Tobacco............................................................. 40
Nipa ester, see Phenoxetol
Niku-nuki ....................................................................................... 13
Nitric acid.................................................................................14, 43
Nitrocellulose ................................................................................. 44
Nitrogen, liquid .............................................................................. 11
Ontogeny..................................................................................26, 31
Operculum.............................................................. 12, 19, 24, 27, 44
Opisthobranch ............................................................................ 9–10
Optics .......................................................................................33–35
Organic material................................................................. 11, 13, 26
Osmium tetroxide..................................................................... 22, 44
Osmotic balance............................................................................. 10
Paper, bleaching ............................................................................. 15
Paper, filter..................................................................................... 29
Paper, heating................................................................................. 15
Paper, label, see Labels
Paper, surface ................................................................................. 14
Paper towel......................................................................... 27, 29–30
Paper, types of..........................................................................14–15
Paraldehyde, paraformaldehyde...............................................42, 45
Patellogastropoda....................................... 10, 12, 22, 26–27, 32, 45
Pencil........................................................................................15, 17
Perchlorethylene ............................................................................ 16
Periostracum ............................................................................16, 43
Permits ....................................................................................... 9, 35
Personal preference ..........................................................................2
Petri dish ........................................................................................ 11
pH.........................................................12–13, 17, 24, 26, 39–42, 45
Phenoxetol, phenoxy isopropanol.................................................. 44
Phosgene ........................................................................................ 42
Photographic paper, blackened ...................................................... 29
Photography .............................................................. 4, 5, 11, 32–35
Photography, optics, see Optics
Picric acid.................................................................................39, 44
Pie pan............................................................................................ 11
Pigment ink .................................................................................... 15
Pin, see also Needle .................................................. 6, 8, 29, 31, 35
Pin holder, see Needle holder
Pipette ........................................................... 6, 7, 11, 21, 27–28, 30
Plankton net ................................................................................... 10
Plastic box, see Box, plastic
Plastic foam, see Foam, plastic
Plastic, surface..........................................................................14, 27
Plasticine ........................................................................................35
Plasticiser........................................................................................15
Pliers ...........................................................................................8, 12
Polycarbon......................................................................................12
Polyethylene ...................................................................................12
Polyplacophora .......................................................10, 12, 30, 40, 45
Polystyrene .....................................................................................15
Polyvinyl acetate, PVA see Glue, polyvinyl acetate
Polyvinyl alcohol............................................................................44
Polyvinylchloride, PVC..................................................................15
Positioning......................................................................................35
Posture ........................................................................................6, 32
Potassium hydroxide ........................................21, 26–30, 32, 44–45
Preservation ....................................................................................11
Prodissoconch.................................................................................22
Propylene glycol.......................................................................44–45
Propylene glycol monophenyl ether, Propylene phenoxetol, see
Phenoxetol
Protein, tanned..........................................................................17, 21
Proteinase K .............................................................................26–28
Protoconch....................................................................13, 22, 26, 31
Ptenoglossate ..................................................................................29
Pulmonata .................................................................................29, 42
Radula.................................................................................19, 22, 24
Radula extraction....................................................24, 26–29, 44–45
Radula mounting ................................................................29–32, 31
Rarefaction .......................................................................................9
Razor blade.............................................................................7–8, 21
Recrystallisation ...........................................................12, 26, 39, 45
Reference specimen, see Voucher
Rehydration ....................................................................................22
Relaxation...........................................................................10–11, 22
Rhipidoglossate ..................................................................29–30, 32
Rock washing .............................................................................9–10
Saliva ........................................................................................14, 29
Salts ..........................................................................................14, 16
Sampling...........................................................................................9
Sand paper ........................................................................................8
Scalpel ..................................................................................7, 21, 24
Scintillation vial .............................................................................13
Scissors .......................................................................................7, 32
SCUBA.......................................................................................9–10
Sediment...........................................................................................9
SEM, animals .....................................................................22–24, 25
SEM, charging..........................................................................17–19
SEM, detectors .............................................................19, 20, 21, 22
SEM, environmental.......................................................................19
SEM, gun types ........................................................................15, 19
SEM, images ................................................................... 3, 5, 20, 23
SEM, mounting .............................................................17, 29–32 35
SEM, parameters ................................................................18, 20, 23
SEM, radula, see Radula
SEM, sample penetration ...............................................................19
SEM, signal mixing............................................................19, 20, 22
SEM, stub ...................................................14–15, 17, 21, 30, 32, 44
SEM, stub map ...............................................................................17
SEM, technique......................................................12, 14, 16–25, 20
SEM, variable pressure ................................................ 15, 18, 20, 22
SEM, uncoated materials ........................................................ 16, 19
Separation ...................................................................................... 11
Sharpening .......................................................................................8
Shell ........................................................................................... 2, 12
Shell, coiled....................................................................8, 17, 21–24
Shell, cracking..........................................................................12, 24
Shell, crushing................................................................................ 12
Shell damage ........................................12–17, 24, 26, 40–41, 43–44
Shell dissolution...........................................8, 12–14, 17, 26, 43–45
Shell drying .................................................................................... 14
Shell grit...........................................................................................9
Shell-less ..........................................................................................2
Shell manipulation, see specimen manipulation
Sieve............................................................................. 6, 6, 9, 11, 28
Silica gel......................................................................................... 15
Silicone .......................................................................................... 35
Silver paint ...............................................................................17–18
Size, specific .................................................................... 2, 6, 11–12
Slide, histology .................................................................. 28–30, 35
Slide warmer .................................................................................. 27
Snail, see gastropod
Sodium bicarbonate ........................................................... 12–13, 45
Sodium carbonate................................................... 12, 16–17, 27, 45
Sodium dodecyl suphate, SDS..................................... 21, 26–27, 45
Sodium hydrogen carbonate .......................................................... 45
Sodium hydroxide ............................................ 16–17, 24, 26, 43, 45
Sodium hypochloride, see Bleach
Sodium lauryl sulphate, see Sodium dodecyl sulphate
Sodium pentobarbitone, see Nembutal
Sodium phosphate .......................................................................... 45
Sodium tetraborate, see Borax
Sonication, see ultrasound
Sorting............................................................................................ 11
Sorting tray..................................................................................... 11
Specimen, drying ........................................................................... 14
Specimen manipulation............................................................ 11, 14
Specimen, types ................................................ 2, 13, 16, 21, 32, 35
Species, number of...........................................................................2
Spin column .............................................................................27, 28
Sputter coating ......................................................................... 18, 30
Static charge .............................................................................14, 27
Stenoglossate.................................................................................. 32
Stereomicroscope ............................... 6–7, 11, 22, 24, 27–30, 34–35
Storage .....................................................................................12, 29
Storage, field .................................................................................. 13
Storage, medium ................................................................ 13, 43–44
Stovaine.......................................................................................... 39
49
Streaks ............................................................................................12
Stub, see SEM, stub
Sublimate, see Mercury chloride
Sunlight ..........................................................................................10
Surface tension ...................................................................17, 21, 30
Swelling..........................................................................................11
t-butyl alcohol.................................................................................24
Taenioglossate ................................................................................29
Tape, office, copper, aluminum, nickel ..........................................18
Terrestrial................................................................................2, 9–10
TEM......................................................................................2, 12, 44
Thermal circulation ........................................................................11
Three dimensional reconstruction ............................................32, 33
Timing ..............................................................................................2
Tissue clearing................................................................................12
Tissue desiccation.....................................................................24, 43
Tissue dissolution .................................17, 21, 26–27, 29–30, 44–45
Tissue hardening.......................................................................43, 45
Tissue shrinkage ..........................................................12, 24, 43–44
Tissue swelling .........................................................................11–12
Tobacco, see also Nicotine .............................................................11
Tools .........................................................................................2, 5–9
Tooth pick.........................................................................................8
Transmission electron microscope, see TEM
Trap.................................................................................................10
Tricaine methanesulphonate, see MS 222
Trisodium phosphate ......................................................................17
Tungsten needle, see Needle, tungsten
Turbulence ................................................................................11–12
Type material, see Specimen, types
Ultrasound ..........................................................................17, 26, 29
Urethane .........................................................................................45
Vetigastropoda ....................................................................26, 30, 32
Vial, glass ...........................................................................13–14, 22
Vice.................................................................................................12
Voucher...............................................................................12–13, 26
Wash, shell................................................................................13, 17
Wash, radula .......................................................................27–28, 30
Water...................................................................................11, 28, 45
Water, distilled............................................14, 17, 27–28, 30, 32, 45
Water, hot..................................................................................13, 29
Wax tray....................................................................................24, 35
Wire ..........................................................................................30, 32
Wire cutter ................................................................................12, 30
Wood.........................................................................................15–16
Workspace ..........................................................................2, 5–6, 27
X-ray tomography ..........................................................................32
Xylene ............................................................................................42
Zip lock bag......................................................................................9
50
About the authors
Daniel L. Geiger is Research Curator of Electron
Microscopy at the Santa Barbara Museum of Natural
History. His award-winning dissertation from 1999 on
abalone systematics and evolution was overseen by coadvisors Dr. Russel Zimmer and Dr. James H. McLean at the
University of Southern California (USC) in Los Angeles,
California. Following a post-doctoral fellowship in
molecular systematics at the Los Angeles County Museum
of Natural History and teaching appointments at USC, he
moved to his current position. He is working on systematics
and evolution of Vetigastropoda, currently focusing on the
exclusively minute Scissurellidae, Anatomidae and
associated families, using light and electron microscopy on
shells and radulae, 3D reconstruction of histological
sections, as well as molecular phylogenetics. His field
experience ranges from the Irish Sea to Papua New Guinea;
photography and digital imaging are keen interests. He is
associate editor with Molluscan Research and the senior
editor for Mollusca with Zootaxa. He is the organizer of the
symposium on micromolluscs at the Unitas Malacologia
2007 conference in Antwerp, Belgium.
Bruce Marshall is Malacologist and collection manager of
Mollusca at Museum of New Zealand Te Papa Tongarewa,
Wellington, which he joined in 1975. He specialises in the
fauna of New Zealand and surrounding areas and has
published extensively on various groups of small gastropods
and monoplacophorans. He is an associate editor with
Molluscan Research.
Winston Ponder retired in 2005 after many years as a
Principal Research Scientist at the Australian Museum,
Sydney. He carried out a major study of rissooidean
microgastropods in New Zealand for his Master of Science
degree. After finishing his PhD on neogastropods he moved
to the Australian Museum in 1968 after briefly taking up a
position in the National Museum of New Zealand. He was
awarded a DSc in 1992 and is the author of over 200
publications on molluscs, many of them on micromolluscs,
particularly marine and freshwater Rissooidea. His current
primary interests include the taxonomy, distribution and
conservation of freshwater and estuarine molluscs, higher
systematics of gastropods and building interactive keys. He
is also writing a book on molluscan biology and evolution
together with Prof. David Lindberg and is the Managing
Editor of Molluscan Research.
Takenori Sasaki is a curator of paleontology and zoology at
The University Museum, The University of Tokyo. He
received his master’s degree in Prof. Okutani’s laboratory at
the Tokyo University of Fisheries with work on
patellogastropod systematics and a Ph. D. degree in the
paleobiological laboratory at The University of Tokyo
carrying out cladistic analyses of ‘archaeogastropods’. After
a post-doctoral fellowship at The University of Tokyo, he has
been in the current position since 1999. His main interests
are: comparative anatomy and phylogeny of whole
molluscan groups, serial sectioning of soft parts, larval shell
morphology and shell microstructure, biodiversity studies of
Japanese molluscs, taxonomic revision of patellogastropods,
and faunal research on deep-sea chemosynthesis-based
biological communities (hydrothermal vents and seeps).
Since 2002 he has especially worked on deep-sea molluscs
as a visiting scientist of JAMSTEC (Japan Agency for
Marine-Earth Science and Technology).
Anders Warén is senior curator at the Swedish Museum of
Natural History in Stockholm. He undertook his doctoral
studies at Göteborg University and has worked extensively
on eulimid gastropods. He has recently worked on deep sea
molluscs of the North-Eastern Atlantic and the Arctic as well
as hydrothermal vent taxa.

Molluscan Research: Techniques for collecting, handling, preparing

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