The World of Copepods
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Virginia Museum of Natural History
1001 Douglas Ave.
Martinsville, VA 24112 USA
Workshop on Taxonomic Techniques for Copepods
Prepared by Janet W. Reid
This information briefly describes some of the most commonly used methods for treating copepod specimens, primarily for taxonomic purposes. Most of these methods are discussed in greater detail in the references listed (Initially developed in 1992; most recent revision April 2000).
- Initial treatment of specimens
- Narcotizing agents
- Staining for sorting
- Recovery of dried specimens
- Microscopic examination
- Mounting Media; Temporary & Permanent media
- Procedure for examination and mounting
- Manipulation and Dissection
a. Narcotizing agents:
Narcotizing agents can be useful to avoid flexion of the body and the antennae, and to aid retention of egg sacs and gut contents during fixation. The agent is usually added slowly, drop by drop.
Oxygen-depleted water: boil water 5-10 minutes and cool.
CO2 excess: produced by a) bubbling CO2 through water; b) pouring soda water into sample slowly; or c) adding pieces of dry ice. CO2 excess: produced by a) bubbling CO2 through water; b) pouring soda water into sample slowly; or c) adding pieces of dry ice.
Cold: refrigerate specimens in refrigerator, than add fixative to sample at same temperature. Cold: refrigerate specimens in refrigerator, than add fixative to sample at same temperature.
MgCL2 · 6H2O (73.2 g/l is isotonic to sea water): add slowly and keep specimens in solution 10-15 minutes before fixing. MgCL2 · 6H2O (73.2 g/l is isotonic to sea water): add slowly and keep specimens in solution 10-15 minutes before fixing. MgCL2 ·
Alcohol: add pure ethanol, methanol, or isopropanol slowly. Alcohol: add pure ethanol, methanol, or isopropanol slowly.
Formaldehyde. Formaldehyde: acts as a narcotizing agent if added slowly.
Propylene phenoxetol: use 1.5% stock solution, mix 1 volume: 10 volumes of seawater and pour over sample. Recommended for meiofauna (Hulings and Gray, 1971).propylene phenoxetol: use 1.5% stock solution, mix 1 volume: 10 volumes of seawater and pour over sample. Recommended for meiofauna (Hulings and Gray, 1971).
Fill sample bottles 3/4 full. Try to fix within 5 minutes after catch.
3-5% formalin buffered with one of several compounds to reduce acidity is the most commonly used fixative.
Borax (sodium tetraborate - Na2B4O7 ) is perhaps the most common buffering agent. To make up a fixative solution, add 2g borax to 100 ml (40% formalin) and invert jar several times during 1 hour. This will raise the pH to about 8 - 8.2.
Other buffers sometimes used are sodium carbonate (NaHCO3) and sodium acetate (C2H3NaO2 · 3H2O).
70% to 95% EtOH (ethanol) is also a good fixative. 70% to 95% EtOH (ethanol) is also a good fixative.
Advantages of EtOH vs. formalin: EtOH yields more relaxed specimens; avoids the necessity of transferring material later to a long-term storage solution; and is less toxic.
Disadvantages: EtOH is a controlled substance; it is more cumbersome than formalin for field use; the vital stain rose bengal is soluble in it and will leach out of stained specimens; and air transport of significant quantities of EtOH, as of all flammable substances, is illegal.
c. Staining for sorting:
Staining samples before or during fixation helps visual separation of specimens from sediment or detritus-filled samples. However, staining is not recommended if the specimens are to be examined by Nomarski Differential Interference Contrast microscopy.
Rose Bengal is an easily water-soluble vital stain, and is perhaps the most commonly used. Make up a 1% solution. Because the stain is adsorbed by clay and organic detritus, overstain especially dirty samples. A few grains of solid stain can be added to samples with a moistened toothpick. Better staining is obtained if the stain is added to samples a few minutes before fixation. Rose bengal will leach out of specimens stored in EtOH. Rose Bengal is an easily water-soluble vital stain, and is perhaps the most commonly used. Make up a 1% solution. Because the stain is adsorbed by clay and organic detritus, overstain especially dirty samples. A few grains of solid stain can be added to samples with a moistened toothpick. Better staining is obtained if the stain is added to samples a few minutes before fixation. Rose bengal will leach out of specimens stored in EtOH. (Beware, it stains internal tissues and makes study of exoskeleton difficult.)
For staining living animals, Neutral Red is possibly the least damaging; Methylene Blue is also used.
Watch glasses were developed by watchmakers to hold tiny parts. They are much better than small petri dishes for sorting copepods. This is because watch glasses are slightly depressed in the center, encouraging movement of the specimens to the middle of the container. The bottoms of some petri dishes are lower near the rim than in the middle, and the specimens will collect around the edge. Another advantage of watch glasses is that they can be stacked. Some have ground glass edges that can be written on.
Within 7-10 days transfer specimens to ethanol or other long-term storage medium; do not leave material in formalin, even buffered, for long periods. This is because specimens become brittle and setae break off easily.
70% EtOH - 1% glycerine is a good long-term storage medium to maintain flexibility of specimens. The glycerine insures against evaporation.
Note: do not use glycerine if the specimens are to be examined by electron microscopy.
Methanol or isopropanol are also suitable long-term storage media. Methanol or isopropanol are also suitable long-term storage media.
A long-term preservative for zooplankton is a solution of propylene phenoxetol (0.5 ml), propylene glycol (4.5 ml), and distilled water or sea water (95 ml) (Steedman, 1976).
Shell vials (without rims) plugged with cotton are simplest for storing tiny specimens. Any glass vial will do, if the mouth is not too narrow.
Glass jars with Bakelite® plastic caps fitted with a Teflon® insert are probably the best type of container for holding smaller vials. Metal caps will rust, softer plastic caps will split eventually, and rubber stoppers will deform, especially if exposed to alcohols.
Storage of many small vials is most conveniently done inside a large mason jar (food preservation jar) filled with 70% EtOH or other storage medium. The rubber seal of the mason jar should be checked periodically (every 2-3 years) for cracking or deformation, and replaced when necessary. The advantage of the mason jar is that it is relatively easy to open and reclose. Jar tops can also be sealed with melted paraffin, an excellent long-term sealant. Disadvantages include the flammability of paraffin, and the necessity to reseal opened jars. Because most plastic caps (including all snap-on caps) split or deform after a few years soaking in alcohol, plugging vials with cotton is much preferred. If plastic-capped jars must be stored in alcohol, Bakelite® plastic screw caps are best.
For long-term storage of important specimens, the triple vial system (Fig. 1) is surest. This is because the alcohol must evaporate to the bottom of the outer vial, then out of the next vial before air reaches the specimen.
Label paper should be heavy, with a high rag content. Vellum paper is acceptable, but it is easily split. New plastic 'papers' are becoming available. Lightweight 'notebook' paper is unacceptable. Labels should be placed inside the vial, or in an outer vial if the specimens are delicate. Especially if delicate specimens are to be shipped, never put the label together with the specimens. India ink or other permanent drawing inks, which bind within the paper fibers, are still the best. Typed letters or lettering from laser printers, which adhere to the paper surface only, will rub or lift off if not specially treated. Laser-printed labels can be sprayed with a fixative and then baked. For mass production of labels, a thermal pressure printer is best. External labels should be considered as temporary only.
General storage conditions and maintenance:
Biological material should be stored away from sunlight and protected from extremes of temperature. The 'wet' storage room must be well ventilated to avoid buildup of formalin or alcohol fumes. Earthquake bars on shelves are always advisable, as they help avoid accidentally knocking bottles off the shelves. Alcohol will of course evaporate faster than water, essentially diluting the alcohol content. Therefore, every few years, the alcohol in the jars should be renewed completely, not just 'topped up'.
e. Recovery of dried specimens:
Even under the best conditions, alcoholic samples may dry out. Either lactic acid or a weak solution of potassium hydroxide may be used with some success to reconstitute dried specimens, but these chemicals have disadvantages. The gentlest reconstitution method is to soak the specimens for hours or days in a dilute (0.25-0.5%) aqueous solution of trisodium phosphate or other strong detergent. Major features of the specimens should become visible, and the specimens can then be rinsed and returned to alcohol. The trisodium phosphate method was discussed by Van Cleave and Ross (1947).
Copepod taxonomy is based mainly on external morphology. Therefore one needs to see details of the integument. It may be desirable to use a clearing medium to reduce visual interference from internal structures, and to stain the integument in order to highlight spine patterns, pores, and other features. Sequence of treatment:
- Mediums, temporary or permanent
- Mounting on slides, temporary or permanent
- Making a record of the specimen
a. Optional pre-treatments to remove soft tissues:
Muscle tissues of some genera such as Eucyclops do not clear satisfactorily in glycerine or lactic acid, and for these, a pre-treatment is helpful.
Treatment 1: Warm in solution of 10% KOH at 90°C for 1-2 hours
Treatment 2: Immerse in a solution of 0.1% household bleach for a few minutes.
The time in bleach or KOH must be adjusted carefully so that spines do not drop off, or the specimen completely disarticulate!
If a pre-treatment is used, follow by rinsing briefly in distilled water and then transfer through serial concentrations to the medium of choice.
The specimen may be stained at this stage. Good stains for small crustaceans are Chlorazol Black E, Fast Green, Acid Fuchsin, Lignin Pink. All stains may be prepared as 1% solution by weight in distilled water or 70% EtOH. Chlorazol black E is useful for integumental features and is stable for years in several media. It is a mild carcinogen; avoid ingesting dust or eating in the laboratory, etc. Chlorazol black E requires several days to dissolve in some media, and residues may remain in the vial. Strain the medium or use a glass rod to pick up media from the top of the vial, to avoid stirring up the residues at the bottom.
Stains may be added to the dissecting media or the mounting media. Because a stain may eventually leach out of a specimen, it is a good idea to add stain to the permanent mounting medium.
Note: Staining may interfere with Nomarski Differential Interference Contrast microscopy.
A good light stain for use with lactic acid is a few drops aqueous solution of methylene blue or lignin pink, added either to the undiluted acid or to 50% aqueous solution. An alternate stain is 1 part chlorazol black E (1% wt. in 70% alcohol/distilled water) to 19 parts lactic acid or glycerol. Or, lactic acid may simply be added to the medium until it becomes about the intensity of weak tea.
c. Mounting media:
The choice of mounting medium depends on the use to be made of the mounted specimens, the type of microscopy employed, and the need for long-term preservation. Certain media, particularly polyvinyl lactophenol and lactophenol, are more appropriate than others for use with Nomarski Differential Interference Contrast Microscopy. Some clearing, staining, and mounting agents were evaluated for this purpose by Koomen and Vaupel Klein (1995).
The use of clearing vs. non-clearing media will depend on the purpose of the mount. If it is desired to keep a dissected specimen for a long time, remember that a clearing medium will eventually make dissected appendages difficult to find and/or to see. Addition of a stain to the medium will help with this problem. Cleared appendages are usually easily seen by means of phase-contrast or Nomarski optics.
Lactophenol was recommended by Huys and Boxshall (1991) for type collections.
Inhalation of phenol or chloral hydrate fumes from some media is inevitable unless the microscope is fitted with a vacuum hood, or a small table fan is used to blow the fumes away. Long exposure to these chemicals is undesirable. Common symptoms of overexposure include headaches, dizziness, and sinus problems. If these symptoms occur, improve the ventilation or use another medium.
- provide good optical contrast, i.e., a suitable index of refraction
- be somewhat viscous for specimen stability
- be makeable with easily available materials
- be stable over a period of time, depending on the purpose
- not react with stains or sealants
- be relatively non-toxic
Common problems with media:
- excessive clearing of specimens
- formation of precipitates or crystals
- excessive drying and formation of bubbles
i. Temporary mounting media:
Glycerine is included in this section because it is liquid, but it is sometimes used for permanent mounts.
water: cheap but evaporates; index of refraction (contrast) poor for crustaceans.water: cheap but evaporates; index of refraction (contrast) poor for crustaceans.
glycerine: specimens must be brought gradually to isotonic state by transfer or evaporation. Gives good contrast for microscopic examination; clears specimens somewhat; non-toxic; stable almost indefinitely, i. e. years. glycerine: specimens must be brought gradually to isotonic state by transfer or evaporation. Gives good contrast for microscopic examination; clears specimens somewhat; non-toxic; stable almost indefinitely, i. e. years.
Problem: will support growth of bacteria and mold after a few months. Add a bacteria/mold retardant such as 1% formalin if necessary. Also attracts ants and other insects.
See comments on permanent mounting under glycerine jelly.
Lactic acid: clears and gives excellent contrast; specimens should be placed in 50% aqueous solution before being placed in full-strength acid. Low toxicity, pleasant to work with.lactic acid: clears and gives excellent contrast; specimens should be placed in 50% aqueous solution before being placed in full-strength acid. Low toxicity, pleasant to work with.
Lactic acid also tends to extend specimens, sometimes bringing legs into better view and making dissection unnecessary. Because of this extending effect, when reporting measurements, it is best to note the medium in which measurements were made. Problems: lactic acid keeps on clearing and softening specimens. After some time, specimens will become as clear as glass and very delicate, and may disarticulate. For permanent storage, specimens must be transferred to another medium. Also, lactic acid clears some internal structures such as seminal receptacles that are used in the taxonomy of some genera. Examine the specimen in glycerine before transferring to lactic acid.
ii. 'Permanent' mounting media:
Media with a gum arabic base - all clearing media:
50 g gum arabic (use lump form, not powder)
50 g pure cane sugar
50 ml distilled water
0.05 g thymol / 1ml formalin
Warm water and gum until dissolved; then add sugar, then thymol.
10 g chloral hydrate
10 ml distilled water
2.5 ml glycerine
6 g gum arabic
Dissolve chloral hydrate in water; add glycerine and mix with glass rod; add gum arabic and mix cautiously, avoiding bubbles; wait 1 week before using. The solid gum arabic and water may be substituted by 12 ml liquid gum arabic. Protect the medium from light. Reyne's medium may develop tiny crystals, thus is not 'permanent' beyond a year or so.
50 g distilled water
50 g gum arabic (clear crystals)
200 g chloral hydrate
20 g glycerin
Mix ingredients as with Reyne's. For a more viscous medium, the amount of distilled water may be reduced to 25 g. This medium may remain stable and provide good viewing conditions for 10 years or more (F. Stoch, personal communication).
10 ml water
3 ml glacial acetic acid
5 ml dextrose syrup
8 g gum arabic
75 g chloral hydrate
Mix water with acid and syrup, dissolve in gum (needs 1 week or more). Stir at intervals; when solution is complete, add chloral hydrate. Similar to Reyne's medium in properties.
8 g gum arabic
10 ml distilled water
75 g chloral hydrate
5 ml glycerine
3 ml glacial acetic acid
Dissolve gum arabic in distilled water. Add chloral hydrate, glycerine, and glacial acetic acid. Strain through clean muslin or glass wool. Protect from light.
Another version of Hoyer's:
30-40 g gum arabic
200 g chloral hydrate
20 cc glycerine
50 cc distilled water
Dissolve gum in water 24 hours, add chloral hydrate, let stand until it dissolves, then add glycerine. If the medium is not clear, filter through glass wool twice. Store in glass stoppered bottle.
Another version of Hoyer's (developed by Tatsunori Itô):
8 g gum arabic
30 g chloral hydrate
10 ml distilled water
1 ml glacial acetic acid
2 ml 100% glycerine
Rub gum arabic and chloral hydrate to powder and mix. Add water, acetic acid and glycerine, then mix slowly, warming slightly if necessary. Filter if the gum arabic is not clean. (Communicated by W. Mielke.) Specimens may be mounted on slides with Hoyer's without dehydrating them first. Also, it is thick enough not to require cover glass supports. Reference: Baker and Wharton (1952).
* Chloral hydrate, a component of most of these media, is a highly toxic and controlled substance. *
Glycerine Jellies - semi-clearing:
Glycerine jellies are excellent for permanent mounts; they clear only slightly and thus interior structures such as the seminal receptacle and muscle bands remain easily visible. Stains added to the jelly will gradually leave the mountant and color the specimen. Good commercial preparations are widely available.
15 g best quality gelatine
100 ml distilled water
100 ml pure glycerine
plus any one of the following, to retard growth of fungus:
1 g phenol
0.01 g merthiolate
1 g thymol
1 ml zephiran
1 ml formalin
Dissolve gelatine in water with gentle heat (not above 75°C), then add pure glycerine and the fungus retardant; stir until thoroughly mixed. Store refrigerated in covered jar.
Second recipe (from Morholt et al., 1966):
10 g gelatine
60 ml distilled water
70 ml glycerine
1 g phenol
Soak the gelatine in the distilled water for about 2 hours. Then add the glycerine and phenol. Heat in a water bath (about 40°C or 104°F), stirring gently until the mixture is blended. Allow to cool. Do not allow the temperature to rise above 40°C, or the colloid will no longer solidify.
To make a glycerine jelly mount:
Add glycerine to the alcohol or formalin in which the specimen is kept, until 10% of the storage fluid is glycerine; then evaporate to 100% glycerine. At that point the specimen can be transferred to glycerine jelly.
Place a drop of the jelly on a slide and warm the slide to about 40°C. If warmed to more than 75°C, the jelly changes to metagelatine and will not set (harden) again at room temperature. There should not be too much medium, i.e., the drop must not extend past the edges of the cover slip. If desired, add 3 or more pieces of broken glass (crushed pieces of cover slip) to support the cover slip. Add the whole specimen or the dissected parts and position them near the center of the drop. Place the cover slip over all, slowly, picking up the edge of the drop of medium. Allow the mount to cool and stabilize for at least a few hours before sealing (mounts made in pure glycerine can be sealed immediately). Appropriate sealants for glycerine or glycerine jelly mounts include Canada balsam, Murrayite®, or waterproof polyurethane. Clear fingernail polish can also be used, but it will dry and separate after a few years. Store slides horizontally.
Problems with glycerine jellies:
Specimens must be in glycerine before being transferred to glycerine jelly, otherwise they may implode.
Dissection must be done in glycerine beforehand and the parts transferred to the warmed and melted glycerine jelly before it cools and hardens. With experience, dissection can be accomplished in the gelatine while it is melted; if it starts to gel (set) too soon, simply re-warm the slide.
Glycerine jelly dries and the mount must be sealed for permanent storage.
Be careful not to heat the finished slides, as near a hot light source, because they will melt. This may also happen when slides are shipped.
Polyvinyl lactophenol (PVL) - clearing:
Recipe 1 (from J.R. Hendricks):
Solution A: 20 ml lactic acid, 20 g phenol crystals or 20 ml liquid phenol
Solution B: 100 ml H2O, 8 g polyvinyl crystals mixed and heated in a double boiler to clear and thicken Mix all of Solutions A and B; blend but do not homogenize. Let stand overnight, mix again. Will clear after several days.
Recipe 2 (from Gray and Weiss, 1950):
2 g polyvinyl alcohol powder; place in
7 ml 70% acetone. Stir, then add
5 ml distilled water,
5 ml glycerine,
5 ml lactic acid. Mix and add
5 ml distilled water. Carefully heat with stirring until mixture clears (do not boil).
If too viscous, add more water, a few drops at a time. There is no commercial source of PVL at present. Appropriate sealing materials for PVL include melted paraffin or fingernail polish. Murrayite should not be used, as it forms bubbles. See discussion by Koomen and Vaupel Klein (1995).
CMC® - clearing:
CMC® is the trade name of another family of media containing polyvinyl alcohol, lactic acid, and other ingredients. CMC works best when specimens are mounted directly from water. Low viscosity (CMC-9) and high viscosity (CMC-10) versions, both with several stains added, or without stain, are available.
A recipe for CMC:
6 g 133,000 mw polyvinyl alcohol
40 ml 70% acetone
25 g chloral hydrate
80 ml distilled water
15 ml lactic acid, USP white grade
CMC-P has phenol instead of chloral hydrate, dries more slowly than CMC, and is usually used to relax live specimens and to clear them rapidly. CMC mounts can be sealed with CMC itself, clear nail polish, waterproof polyurethane, or Murrayite®.
Advantages of PVL and CMC:
Specimens can be added from water, formalin or ethanol solutions, glycerine, or lactic acid without serious distortion of all but the most delicate specimens.
Both media are supposedly self-ringing, i.e., slides can be stored horizontally in normal temperature/humidity conditions for long periods (months) without serious drying. For longer storage, storage in unusual conditions, or for slides that are to be shipped: allow PVL slides to dry in a horizontal position, preferably in a desiccation chamber, for 7 days or longer, then seal. CMC slides can be sealed immediately with additional CMC, or other sealant.
PVL/CMC evaporate continuously. Allow to evaporate for 1-15 minutes, depending on the initial viscosity of the medium, before applying a cover slip. This permits evaporation of some phenol, thus avoiding excessive formation of needlelike phenol crystals under the cover slip. The problem of crystal formation is less serious with CMC than with PVL. Allowing the medium to become a bit viscous before applying the cover slip also avoids excessive movement of small body parts to the edge of the mount. One technique to fix parts on the slide is to run a small line of medium on the slide, arrange the dissected parts in this line, allow to dry for several minutes, and then cover with more medium and apply the cover slip. This works best with larger specimens. Dissection must be accomplished within 5-10 minutes before the medium becomes too viscous. If this is a great problem, dissection can be done in lactic acid or glycerine and the parts transferred quickly to the final slide. Also, both PVL and CMC can be diluted with distilled water. Parts mounted in PVL tend to go 'flat,' i.e., lose some contrast, after a few days. This problem can be ameliorated but not completely solved by including a light stain.
Note: Huys and Boxshall (1991) recommended against the use of PVL for type collections because PVL tends to overclear within about 10 years, may form crystals, and dries out if not sealed. Huys and Boxshall recommended lactophenol for type specimens.
Lactophenol - semi-clearing:
30 ml melted phenol crystals
10 ml lactic acid
20 ml glycerin
10 ml distilled water
Lactophenol was recommended by Huys and Boxshall (1991) for type collections. This medium does not clear strongly, and allows re-mounting if necessary. It requires sealing.
Canada Balsam - clearing:
Abstracted from article by Thatcher (1987):
Fix specimen in AFA (85 parts 85% EtOH; 10 parts formalin; 5 parts glacial acetic acid) for at least 10 minutes. Next, pass specimen directly from AFA to stain solution (95% EtOH colored to the intensity of weak tea with equal parts of Eosin and Orange-G stains). Stain in this solution for 3-10 minutes and then transfer specimens to pure phenol (liquefy phenol crystals with a bit of 95% EtOH to make this solution). The phenol simultaneously dehydrates, clears, and destains the material. When the specimen is clear in phenol a few seconds later, it is already dehydrated, but if more destaining is desired it may be left in this solution for a few minutes. After the proper degree of destaining is achieved, pass the copepod to methyl salicylate, which stops the destaining process. After 3 minutes in methyl salicylate, the specimen can be mounted in Canada balsam. The entire process requires 8-10 min.
Material fixed by other means and stored in 70% EtOH can be processed in the same way, but specimens in aqueous solutions (i.e., formalin) must be placed in 70% EtOH for a few minutes before staining.
If a specimen collapses in methyl salicylate (copepods seldom do), it may not have been properly fixed or have been dead too long before fixation. Return the specimen to phenol, where it will return to normal shape in a few minutes. It may be necessary to perforate the animal with a fine needle to permit a more rapid exchange of liquids. It is sometimes helpful to pass such specimens through a solution of 50% phenol/ 50% methyl salicylate before transferring them to pure methyl salicylate. It often happens that a copepod is fixed in an undesirable position with the antennae wrapped around the body or the abdomen curled under. These conditions can be corrected because a specimen in phenol becomes soft and pliable. It can be taken from phenol, placed on a dry slide, and manipulated into a good position with dissecting needles. Arranging the legs at this time may obviate the necessity for dissection. When the animal is in the desired position, place a coverglass on top to hold it and add some methyl salicylate. The latter hardens the specimen in a few seconds, and it will retain the same form when mounted in balsam.
Offers 'HYP' (hundred-year preservation). Canada balsam is one of the few media known to remain stable this long. Any slide made with balsam can be demounted by soaking in methyl salicylate or xylene for a few hours. Slides made with PVA or glycerine jelly, on the other hand, cannot be successfully demounted.
This method will produce good whole-mounts, but is difficult to use for dissected parts. Canada balsam is a thick, viscous medium and will not yield the thin mounts necessary for adequate inspection of the tiniest structures.
a. Manipulation and Dissection:
For initial sorting and manipulating copepods, a blunt probe or a cactus quill mounted in a needle holder can be used. Fine (needle-nosed) forceps, pasteur pipettes fitted with a flexible rubber bulb, or Irwin Loops are useful to transfer individual specimens without transferring unnecessary amounts of medium. Irwin Loops are available commercially (Section 5), or can be made by electrolyzing tungsten wire to a tapered point, then bending the wire in a loop with fine forceps. Pasteur pipettes should be cleaned before use. For manipulating and transferring tiny mouthparts, use a fine needle (minuten pin), or an eyelash mounted on a holder.
Dissection is accomplished most easily in glycerine, lactic acid, PVL, or low-viscosity CMC. Specimens can be dissected in glycerine for temporary mounts, or directly in the permanent medium. It is often convenient to dissect in the eventual mounting medium, rather than to attempt to transfer small parts.
The choice of dissecting medium depends on the eventual mounting medium. Dissect in lactic acid if the specimen is to be mounted in PVL or lactophenol. Dissect in water or glycerine for aqueous media. For dissection, use either fine (0.2 mm thickness) stainless steel entomological pins ('minuten pins,' 'minuten nadeln') mounted in wooden holders; tungsten needles; or a micro-scalpel.
To make tungsten dissecting needles: Sharpen pieces of tungsten wire (about #29 gauge, 0.25 mm, or 400 µm diameter) by electrolyzing one end in a 6-volt circuit. Use a microscope transformer to provide the 6-volts: one lead is bare wire, the other is attached to an alligator clamp for holding a piece of tungsten wire about 2 cm long. Hold the tip of the tungsten wire in a solution of about 20% KOH or NaOH, for several minutes to an hour, dipping the wire in and out repeatedly until the desired shape is obtained. Mount the sharpened wire on the end of a wooden or plastic holder. To make tungsten dissecting needles: Sharpen pieces of tungsten wire (about #29 gauge, 0.25 mm, or 400 µm diameter) by electrolyzing one end in a 6-volt circuit. Use a microscope transformer to provide the 6-volts: one lead is bare wire, the other is attached to an alligator clamp for holding a piece of tungsten wire about 2 cm long. Hold the tip of the tungsten wire in a solution of about 20% KOH or NaOH, for several minutes to an hour, dipping the wire in and out repeatedly until the desired shape is obtained. Mount the sharpened wire on the end of a wooden or plastic holder. To make tungsten dissecting needles: Sharpen pieces of tungsten wire (about #29 gauge, 0.25 mm, or 400 µm diameter) by electrolyzing one end in a 6-volt circuit. Use a microscope transformer to provide the 6-volts: one lead is bare wire, the other is attached to an alligator clamp for holding a piece of tungsten wire about 2 cm long. Hold the tip of the tungsten wire in a solution of about 20% KOH or NaOH, for several minutes to an hour, dipping the wire in and out repeatedly until the desired shape is obtained. Mount the sharpened wire on the end of a wooden or plastic holder. Tungsten is not reactive to acids, alkalis, or ordinary laboratory solvents, and can be cleaned in a flame.
To make a micro-scalpel: Use a standard (non-stainless) insect pin, size 00 or 000. Mount the pin on the end of a thin wood or plastic holder. Sharpen the pin on two sides on a fine stone with a little mineral oil, such that it forms a knife blade. To make a micro-scalpel: Use a standard (non-stainless) insect pin, size 00 or 000. Mount the pin on the end of a thin wood or plastic holder. Sharpen the pin on two sides on a fine stone with a little mineral oil, such that it forms a knife blade.
Dissecting with this tool works best on a plastic petri dish. With the left hand, impale the copepod in a convenient area and anchor it to the bottom of the dish. With the micro-scalpel in the right hand, slice off the mouthparts and appendages. (Personal communication from J. Cordell.)
Needle holders: Holders should be light and about 6 inches long. The thickness is according to personal preference. Wooden holders can easily be made by cutting lengths of dowel rod, grinding one end to a rough taper, then making a hole in that end with a dressmaker's pin. Next, place a drop of epoxy over the hole, insert a needle into the hole, and store the handle with the needle up until the epoxy hardens completely. Needle holders: Holders should be light and about 6 inches long. The thickness is according to personal preference. Wooden holders can easily be made by cutting lengths of dowel rod, grinding one end to a rough taper, then making a hole in that end with a dressmaker's pin. Next, place a drop of epoxy over the hole, insert a needle into the hole, and store the handle with the needle up until the epoxy hardens completely.
Thick glass capillary tubing is also handy for holding needles, but it is becoming less available. Use the best dissecting microscope available, such as the Wild M5A. This microscope provides excellent optical quality and a range of magnifications (fixed stops of 60x, 120x, 240x and 500x), using 10x eyepieces. A 2x auxiliary lens is available to double these magnifications; however, this lens tends to reduce the available light when used at 500x, which is a critical factor. Zoom optics are adequate, but do not approach the optical quality available in fixed-stop lenses. Some people may prefer armrests, also available on the Wild M5. Substage illumination providing transmitted light is almost essential for the best contrast.
This is the most frustrating part of copepod manipulation and only much practice will produce results.
Hold a pin in each hand. Hold the specimen down with one pin, remove the body part with the other pin. For small specimens, removal of an entire somite with associated legs (P1-4) may be easier; the body somite can then be separated from the paired legs. Mouthparts must be dissected individually. Some people prefer to begin with the cephalic appendages, others with the urosome. Dissection sequences have been discussed by Coull (1973), Hamond (1969), Humes and Gooding (1964), and Huys and Boxshall (1991).
Use standard 3 × 1 inch glass slides, with one end frosted if preferred. Thin slides are best. Clean with 70% alcohol. Cover slips should be No. 0 or 1 thickness for plankton-size specimens. The shape of the cover slip (rectangular or round) and arrangement of specimens is according to personal preference. One option is 15 mm round cover slips, 2 of which fit easily on a slide with room for a label (Fig. 2):
Make a streak of the mounting medium horizontally or vertically in the middle of the slide. Add a small drop of mounting medium at the end of the slide. Place the specimen in this drop and dissect off the appendages, placing each in sequence in the streak of medium. Allow the streak to dry slightly. Then take a clean coverslip in a forceps, make a cross of the mounting medium on the coverslip, and gently lay the coverslip over the streak of medium on the slide, with the arms of the cross at 45° to the partly dried streak of medium. Do this slowly to avoid formation of air bubbles. The cover slip may be supported with glass fragments, threads, or small bits of wax. Do not use synthetic modeling clay, which eventually disintegrates. Slides may be conveniently stored temporarily in glass or plastic petri dishes, covered against dust and insects. In humid climates, a desiccation chamber is useful.
Store slides horizontally 1-2 weeks to stabilize and dry the medium. Ring (seal) with clear nail polish, Murrayite®, balsam, waterproof polyurethane, epoxy, paraffin, etc., or the medium itself if it will harden. (Nail polish tends to dry and separate after a few months, and therefore is not the best choice for 'permanent' slides.) The sealant should be allowed to run under the edges of the cover slip if possible. It is best to label slides permanently with a diamond needle in addition to paper labels, especially in tropical climates.
Procedure for mounting specimens
- Transfer the specimen to a few drops of 70% EtOH - 10% glycerine on a depression slide. The choice of depression slide thickness will depend on the size of the specimen. Use the thinnest possible depression slide, because thin slides provide better optical conditions.
- Wait 5-10 minutes while the solution evaporates to nearly pure glycerine. Watch to be sure that the specimen remains in the liquid drop.
- Place a cover slip (round ones are best) partly over the depression, allowing liquid to flow under the slip. Add a little pure glycerine to the depression if necessary.
- With needles, push the specimen under the cover slip near the edge of the depression. The specimen can be rolled gently beneath the cover slip to any desired orientation.
For calanoids, look for details of thoracic 'wings' and urosome; antennule (A1) details may also be visible at this stage. Next, remove the specimen from beneath the cover slip and dissect off both antennules and leg 5, either returning these beneath the cover slip or transferring them to a temporary or permanent mounting medium on a flat slide if desired.
For cyclopoids, look for the presence of a spine on the medial expansion of the leg 1 (P1) basipodite. Counting the number of articles in the antennule (A1) and the swimming legs (P1-4) is also easy at this stage.
Next, remove the cyclopoid from under the cover slip, catch the 4th legs against the edge of the cover slip, and push the specimen back under the cover slip so that the swimming legs are bent anteriorly (Fig. 3a). This pins the specimen in a stable position with the structures (4th and 5th legs and urosome) usually needed for species-level determination easily viewed (Fig. 3b, c).
For most harpacticoids, it may also be possible to view all necessary structures without dissection by rolling the specimen under the cover slip into various positions. Sometimes most structures can be seen if the specimen is separated between the 4th and 5th legs. If legs 1-4 are preserved in awkward positions, full dissection may be necessary.
For small species, details may not be sufficiently visible in glycerine. Transfer to lactic acid, supporting cover slip with several pieces of glass. Lactic acid usually extends the specimen so that dissection may not be necessary.
- Specimens can be returned from glycerine to 70% EtOH if desired; serial dilutions may be desirable. To make permanent mounts, specimens can be transferred from glycerine directly to glycerine jelly, Hoyer's, CMC, or PVL.
Baker, E. W. and G. W. Wharton. 1952. An Introduction to Acarology. MacMillan Co., New York. 465 pp.
Beckett, D. C. and P. A. Lewis. 1982. An efficient procedure for slde mounting of larval chironomids. Transactions of the American Microscopical Society 101: 96-99.
Coull, B. C. 1977. Marine fauna and flora of the northeastern United States. Copepoda: Harpacticoida. U.S. Dept. of Commerce, National Oceanographic and Atmospheric Administration Technical Report, NMFS Circular 399. 48 pp.
Graeter, E. 1910. Die Copepoden der unterirdischer Gewässer. Inaugural-Dissertation zur Erlangung der Doktorwürde, Universität Basel. E. Schweizerbart'sche Verlagsbuchhandlung, Stuttgart. 87 pp. + Plates I-III.
Gray, P. and G. Weiss. 1950. The use of polyvinyl alcohol and its derivatives as microscopical mounting media. Part I. Water miscible mounting media. Microscopical Society Series III, xx, 3: 287-291.
Hamond, R. 1969. Methods of studying the copepods. Journal of the Quekett Microscopical Club 31: 137-149.
Hulings, N. C. and J. S. Gray. 1971. A Manual for the Study of Meiofauna. Smithsonian Contributions to Zoology 78: 1-83.
Humes, A. G. & R. U. Gooding. 1964. A method for studying the external anatomy of copepods. Crustaceana 6: 238-240.
Huys, R. & G. A. Boxshall. 1991. Copepod Evolution. The Ray Society, London. 468 pp.
Koomen, P. & J. C. Von Vaupel Klein. 1995. The suitability of various mounting media for permanent mounts of small chitinous crustaceans, with special reference to the observation of integumental organs. Crustaceana 68: 428-437.
McKinnon, D. 1991. A Toolbook for the Practical Identification of Pelagic Copepods. 60 pp. (Unpublished pamphlet)
Morholt, E., P. F. Brandwein & A. Joseph. 1966. A Sourcebook for the Biological Sciences. 2nd ed. Harcourt, Brace & World, Inc., New York.
Steedman, H. F. 1976. Zooplankton Fixation and Preservation. UNESCO Press, Paris. 350 pp.
Thatcher, V. E. 1987. Hope for HYP. Monoculus Copepod Newsletter 15: 20-23.
Westheide, W. and G. Purschke. 1988. Organism Processing. Chapter 10, pp. 146-160 in Higgins, R. P. and H. Thiel, eds. Introduction to the Study of Meiofauna. Smithsonian Institution Press, Washington, DC.
Van Cleave, H. J. and J. A. Ross. 1947. A method for reclaiming dried zoological specimens. Science 105(2725): 318.
Note: Some of these references are available from the C. B. Wilson Copepod Library, Department of Invertebrate Zoology, National Museum of Natural History, Smithsonian Institution, Washington, DC 20560-0163 USA. Contact J. W. Reid or T. Chad Walter, firstname.lastname@example.org.
I am grateful for helpful comments and contributions by Dorothy B. Berner, Maria Cristina Bruno, Jeffery R. Cordell, Vicki Medland, Wolfgang Mielke, Carlos E. F. Rocha, Fabio Stoch, Regina Wetzer, and many other colleagues over the years.
Any comments, corrections, additions or
deletions should be directed to:
T. Chad Walter
This page was lasted updated October 2007
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