WORMWOOD

A resource for nematode phylogeny

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Extracting, fixing and mounting nematodes

This page explains some of the easier and faster methods of processing microscopic nematodes from soils and sediments. More comprehensive reviews are listed in the reference sections of the different chapters. (30 November 1995)

Contents of this page:

Sampling and extracting

Fixing and dehydrating

Handling single worms

Mounting worms in temporary slides

Mounting worms in permanent slides

A. SAMPLING AND EXTRACTING NEMATODES

1. Where to look for worms

Finding nematodes is no problem, but getting the species and numbers you want may be more tricky. The best places to look for fast-growing, easily cultured bacterial feeders are sites of active and abundant organic decomposition e.g. compost, rotting wood, chicken coops, dung heaps and other select spots. These will usually contain large numbers and several species of Rhabditida and Diplogasterida. Instead of looking for places of festering stench and decay, you can also bait such species by leaving rotting meat or a boiled potato on soil for a week or so. A wide range of other groups and species can be found especially in dark forest soil, which often contains larger predators and omnivores (Mononchida, Dorylaimida) as well as plant parasites (Tylenchida) and slower-growing bacterial feeders (Cephalobidae, Plectidae). The edges and sediment of pools and ponds can contain a variety of adenophorean bacterial feeders (Monhysterida, Chromadorida, Enoplida) as well as predatory diplogasterids, mononchs and dorylaims. Generally speaking, natural and undisturbed habitats will yield greater diversity and more slow-growing nematode species, while temporary and/or disturbed habitats will yield fewer and fast-multiplying species. Check the Methods for culturing freeliving nematodes page for some pointers on very rough identification.

2. Taking samples

Sampling nematodes does not require exceptional skills, but the one thing to take care of is avoiding contamination of samples with nematodes carried over on or in dirty tools. Collect 200-500 ml of soil/sediment/litter or whatever substrate you want to investigate, take it to the lab in a plastic bag or pot. If you do not want to extract straightaway, most nematodes in the samples will survive storage at 4 °C for several weeks.

3. Extracting live nematodes

An endless range of extraction tools and techniques have been developed, mostly because no single technique will efficiently extract all sizes and kinds of nematodes. Click here for some references on extraction techniques. We will only concern ourselves here with the simplest method: the extraction tray. This method is not quantitatively reliable as it selects against slow and non-moving nematodes, but it is extremely cheap, relatively fast and very good at yielding large numbers of live and kicking worms. The three components required are: (1) a large plastic tray, (2) a wide mesh and (3) reasonably strong kitchen paper. The larger the tray, the more material can be extracted in one run. A handy size and type is e.g. a dissection tray or a photographic developing tray. The mesh can be a single layer of thick plastic or rubber netting, or alternatively it can consist of a plastic-coated metal grid covered with thin plastic gauze. It should NOT contain any bare metal, because this may release toxic ions.

Place the mesh in the tray, cover its bottom and sides with a single sheet of kitchen paper (one or two layers) and spread a thin, uniform layer of soil out over the paper. Gently pour clean water (tap water may contain nematodes!) into the tray until the soil is wet but not submerged. Avoid spilling soil particles along the sides or through the filter paper - they will cloud up the extract and make it difficult to see the nematodes. Leave the tray until the next day. Actively moving nematodes will sooner or later crawl down through the kitchen paper and sink to the bottom of the tray. A large proportion of the sampled worms will thus collect on the bottom of the tray overnight, and the next morning you should have a plentiful abundance ready for further treatment.

Concentrate the suspended worms by pouring the water out of the tray over a fine sieve (mesh 25 um or less) and washing the nematodes off in a small beaker with about 20-30 ml water. If you do not have a sieve, pour the extract water into a sufficiently large beaker and allow the nematodes to settle (takes about 30 minutes). Then pour off most of the top water and transfer the remaining water to a smaller beaker. Repeat until you have 20-30 ml suspension left. If you have sampled on a good spot, you should now have hundreds or even thousands of nematodes ready for further processing.

A few references on sampling and extraction techniques

Southey, J.F. (1986) Principles of sampling for nematodes. in: Southey, J.F. (ed.) Laboratory methods for work with plant and soil nematodes. MAFF, London: 1-4.

Hooper, D.J. (1986) Extraction of free-living stages from soil. in: Southey, J.F. (ed.) Laboratory methods for work with plant and soil nematodes. MAFF, London: 5-30.

Hooper, D.J. (1990) Extraction and processing of plant and soil nematodes. in:Luc, M.; Sikora, R.A. & Bridge, J. (eds.) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, Wallingford: 45-68.

Bloemers, G.F. & Hodda, M. (1995) A method for extracting nematodes from tropical forest soil. Pedobiologia 39: 331-343.

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B. FIXING AND DEHYDRATING NEMATODES

A number of fixatives are commonly used for preserving terrestrial and aquatic nematodes. Most of these contain formalin and should be handled with due care - rumor has it that a substantial percentage of nematologists died from cancers induced by a lifetime of inhaling formalin vapors.

After fixation, nematodes are usually transferred to glycerin via a dehydration procedure, because this will stop further denaturation and decay, while also resulting in highly transparent specimens ideal for observation through high-powered light microscopes. Stains are not commonly used, because these will not only highlight certain features but also mask other structures. Click here for some references on fixation and processing methods.

Below, two easy and quick methods for fixing and processing worms are presented. Prior to this, however, it is important to stress that you must at all costs avoid contraction of the animals prior to death.

1. Two humane ways to kill worms

NEVER FIX LIVE NEMATODES WITH COLD FIXATIVE. They will not be killed instantaneously, but (apart from suffering a gruesome death) will contract and coil to a degree that often makes them useless for detailed study. To make sure nematodes are nicely stretched upon fixation, you must kill them instantaneously, either by using hot fixative or by heat-killing them prior to adding fixative. Both procedures result in what is known euphemistically as "heat relaxation", relying on a knock-out heat shock to instantly relax the musculature. Another method consists of cooling the nematodes prior on melting ice to adding hot fixative, but this is not fail-safe. Note also that it may sometimes be advisable to starve the nematodes for a few days prior to killing and fixing, because well-fed animals can contain so many intestinal granules that other organs remain obscured even after transfer to glycerin.

Hot fixative often works quite well for direct fixation of an entire soil sample prior to fixation. Heat an equal volume of double-strength fixative to 65-70 C, add it to the soil sample in a sufficiently large plastic pot with screw-cap, close and shake thoroughly. Note that dead worms don't wriggle, so you cannot extract them with an extraction tray. If you fix a soil sample you will therefore need to use more complicated extraction devices such as Cobb sieves, a centrifuge or such. For simplicity's sake, these methods are not discussed here and the remainder of the text applies to nematodes extracted alive prior to fixation.

The best way to kill live nematodes that are collected in a small volume of water (e.g. from an extraction tray or an agar plate) is to transfer them to a glass vial and plunge this in a 70-90 °C water bath. Stir the vial for 20-30 secs and check under the stereo microscope that they are all motionless and stretched out. Make sure they are not boiled - this messes up cellular structure. After heat-killing, it is usually best to fix with hot fixative anyway because this will be more chemically active. Note that this also implies it will be more chemically active on your respiratory tracts, so keep the hot fixative capped as much as possible or work under a flow hood.

2. Fixing with formalin-glycerin and transferring to glycerin through ethanol

THIS PARAGRAPH PREVIOUSLY CONTAINED A SILLY MISTAKE IN THE ORDER OF STEPS TAKEN! It has now been corrected courtesy of Wim Bert. Apologies to anyone who followed the erroneous protocol and up with unusally wrinkled worms...

Prepare double-strength FG fixative containing 8% formalin and 2% glycerin in distilled water. Transfer live nematodes to a small glass vial and allow them to settle to the bottom. Draw off surplus water until they are left in about 2 ml water. Kill the nematodes by stirring the vial 20-30 seconds in a 70-90 °C water bath, check they are all dead and stretched, and then add an equal volume of 65-70 °C fixative. Stir, and then leave the vial alone for a day to allow the fixative to penetrate and act on all tissues.

Take a clean jam jar with airtight-sealable cap. Prepare a "weck pot" by adding ethanol to the jam jar until there is a layer of about 5-10 mm on the bottom. Place a small platform (e.g. a small inverted petri dish or cavity block) on the bottom of the jam jar so that its top surface is raised above the ethanol. Take the vial with FG-fixed worms; if this has a narrow opening transfer the worms to a cavity block. Draw off as much fixative as possible without losing nematodes, and then place the block or vial on the platform inside the jam jar and seal this. Leave the "weck pot" overnight in an incubator at 35-40 C. This will allow all water in the suspension with the nematodes to be replaced with ethanol.

The next day, take the vial or block out of the weck pot and fill the vial or block with 5% glycerin-95% ethanol solution (to the brim if in a cavity block, to about 5 mm high if in a vial). Leave it open in the 35-40 °C incubator for 2-3 hours, to evaporate about half of the ethanol (if necessary cover partly to prevent complete evaporation). Refill with 5% glycerin-95% ethanol solution, leave for another 2-3 hours, and refill one last time before leaving the vial or block overnight in the incubator at 35-40 C. By the next day, the nematodes will impregnated in pure glycerin and ready for mounting in slides, or for stocking without fear of desiccation.

The entire FG/ethanol procedure takes only three days and usually results in well-fixed worms that will not decay for decades. Transferring through ethanol dissolves cuticular lipids, however, and may result in a finely wrinkled cuticle that will show up as such under the scanning electron microscope. If you want to avoid this, try the slightly slower method below.

3. Fixing with TAF and transferring to glycerin through evaporation

Prepare double-strength TAF fixative containing 8% formalin and 2% triethanolamine in distilled water. Transfer live nematodes to a small glass vial and allow them to settle to the bottom. Draw off surplus water until they are left in about 2 ml water. Kill the nematodes by stirring the vial 20-30 secs in a 70-90 °C water bath, check they are all dead and stretched, and then add an equal volume of 65-70 °C fixative. Stir, and then leave the vial alone for a day to allow the fixative to penetrate and act on all tissues.

Take the vial with TAF-fixed worms; transfer the worms to a cavity block if this is easier for subsequent manipulation under a stereo microscope. Draw off as much fixative as possible without losing nematodes, and then fill the vial or block with a solution of 5% glycerin in distilled water (to the brim if in a cavity block, to about 5 mm high if in a vial). Place the block or vial in an incubator at 35-40 °C and cover it nearly completely, leaving a narrow slit for for slow evaporation. Leave until a substantial amount of water has evaporated. Refill with 5% glycerin, and then leave at least two more days in the incubator until all water has evaporated. Check the degree of dehydration by transferring two or three specimens to a drop of pure glycerin. If their cuticle collapses, they are not yet completely dehydrated - return everything to the incubator for another day or two.

The entire TAF/evaporation procedure takes four to six days and often results in perfectly fixed worms. Triethanolamine affects the cuticle of some groups of nematodes, however, and can e.g. result in strong swelling of the cuticle of Rhabditida after several years. In comparison with FG, this method is recommended for better instantaneous preservation and for SEM material, but less suitable for long-term preservation.

A few references on fixation and dehydration techniques

Hooper, D.J. (1986) Handling, fixing, staining and mounting nematodes. in: Southey, J.F. (ed.) Laboratory methods for work with plant and soil nematodes. MAFF, London: 59-80.

Sulston, J. & Hodgkin, J. (1988) Methods. in: Wood, W.B. (ed.) The nematode Caenorhabditis elegans. Cold Spring Harbor Laboratory, Cold Spring Harbor: 587-606.

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C. HANDLING SINGLE WORMS

During various procedures, you will often need to transfer individual nematodes from one dish or vial to another. There are basically two ways of doing this. The traditional method relies on a picking device such as a handle equipped with a fine needle, eyelash, nylon fibre, platinum wire, etc.; it usually requires a good deal of exercise before you will get the hang of it. The alternative is pipetting, which is usually much easier but requires that the nematodes are more or less suspended in liquid.

1. The Noble Art of Picking Worms

While you can cheat and use pipettes or micropipettes for most culture procedures, there are no easy ways out when you need to make slides: you have to learn to pick nematodes with an eyelash, a hair, a fine needle or such. Picking worms off fairly dry agar shouldn't be too difficult (unless you have a very agile species), although you may need to use an inconveniently thick or flattened pick if you do not want to damage the agar surface. Picking worms out of a suspension is a different matter, however.

Half the trick is getting a good picking device, e.g. a fine and rigid insect needle tapped against the table to bend its tip to a minute hook, or a hand-picked hair from the most recalcitrant and brushy moustache available in your lab. Mount one of these on a handle and prepare yourself for a few memorable hours at the stereo microscope.

The other half of the trick is getting the hang of suspended-nematode-dynamics while working at a stereo microscope. Select the specimen of your choice in a petri dish or cavity block with nematodes in liquid (water, glycerin or whatever is appropriate for the procedure at hand). Try to pull this nematode free from the bottom with a short twitch of the pick, and tease it up to the surface of the liquid until it is more or less horizontal, using one hand to move the pick and the other to change focus. Then position the tip of the pick for the final twitch. For curled worms, try to catch them by the crook of the curved part(s) of their body. Straight worms are more fun (or more trouble), because these will often only come out if you position the pick just below them and at a slight angle to their body axis. With luck, the viscosity and surface tension of the liquid will make the worm stick to the pick instead of pulling it back down. Small straight worms are the best (or worst) because they only have a small surface for viscosity to act upon, and will obstinately refuse to cling to your pick. Teasing a straight, 0.3 mm long nematode out of pure glycerin is a particularly nerdy way of spending long winter evenings, or challenging friends and enemies!

2. Pipettes are (usually) easy

A generally faster and safer method of picking single specimens from a suspension consists of using a precision micropipette with a non-sticky tip. This will especially be easier if you are a molecular biologist used to micropipettes and not at all used to worm-fishing. Most nematodes will enthousiatically stick to plastic tips, so you have to avoid this either by siliconizing tips (which may not be very healthy for the worms), or by inserting a fine glass tube and welding it to the plastic tip with paraffin. The latter is done as follows. Take a fine Pasteur pipette at both ends, heat it near the tip above a flame until the glass softens, and pull off the tip smoothly. Break off the drawn-out end of this glass tip until you have an opening of appropriate size, i.e. about as wide as one to one-half the body length of the nematodes you are going to handle (check under stereo microscope). Cut off the narrow end of a yellow Gilson tip or a crystal Eppendorf tip and insert the glass tip halfway (if you use an Eppendorf 1-10 µl pipette, make sure the glass tube is not pushed out when pumping). Heat the tips of a pair of tweezers in a flame, stick them in solid paraffin and transfer a drop of molten paraffin to the joint of the plastic tip and glass tip. Allow it to set and seal the joint by turning the tip slowly between your fingers. Mount the non-sticky tip on a 1-10 µl or 1-20 µl micropipette, and check its sucking and pumping action BEFORE handling nematodes.

Non-sticky tips allow you to pick one or more individuals from a suspension in a single motion, check for their presence inside the tip, and transfer them in a volume of water (usually 2-5 µl is ok) without any imminent danger of desiccation. They are especially efficient for handling very small nematodes. However, a major drawback is the near-certainty that you will carry over any contaminating microorganisms in the suspension. After use, wash the tip with alcohol to avoid contamination between different samples.

Instead of using a micropipette, you can also suck up single nematodes with a Pasteur pipette fitted with a mouth tube, which requires more skill, but can also be used on dry agar. In the latter case, the worms are exposed to desiccation and will stick to the glass, so it is only useful if you want to remove individuals rather then transfer them. See Hooper (1986) for some other devices.

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D. MOUNTING NEMATODES IN TEMPORARY SLIDES

If you want to check the detailed morphology of live nematodes, you can do this by making a temporary slide with live worms on a thin layer of agar. Administer one or two drops of hot 4-5% agar on a glass slide, and immediately flatten this agar with another glass slide provided with spacer strips of thick plastic tape. Carefully remove the top slide when the agar has set. Add a drop of water on the agar, transfer your nematode to it and drop a coverslip on top. The pressure between the coverslip and the hard agar will slow down the worm sufficiently to observe it with oil immersion magnification. If you want to immobilise it almost completely, try smearing some vaseline on the rims of the coverslip, place it on top of the agar and nematode, and very carefully press down the rims of the coverslip until the worm is trapped but not squashed. A complete vaseline seal will also prevent desiccation. It is usually possible to recover the live nematode from such a slide after studying it. See Sulston & Hodgkin (1988) for more specialized procedures.

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E. MOUNTING NEMATODES IN PERMANENT SLIDES

Once nematodes have been fixed and transferred to glycerin, making permanent slides is easy. Various types of slides and mounting aids exist, and this text only deals in detail with glass slides using paraffin as seal and separator. But it is worth noting that the most luxurious and safe mount for microscopic nematodes is a "Cobb slide" (see e.g.Hooper, 1986), which consists of an aluminium carrier supporting two coverslips between which the specimens are sandwiched and sealed. Unlike glass slides, Cobb slides will not break when dropped, and they can can be turned over and observed from both sides at high power.

1. Preparing a glass slide

Fill a petri dish with paraffin granules, melt them at about 60 °C and allow the paraffin to set into a solid layer. Take a 10 cm long cross-cut metal tube with smooth, thin rim and slightly smaller diameter than the coverslips you use (e.g. a 16 mm diameter tube for 18 mm diameter coverslips) and heat one end in a flame. When the other end of the tube is beginning to get hot in your hand, push the heated end down vertically in the paraffin so that it gets covered by melting paraffin, and then press this end down vertically on the middle of a glass slide. Lift the tube, and you should leave behind a complete 3-4 mm thick ring of setting paraffin. Transfer a small drop of pure glycerin to the center of this wax ring, e.g. with a thin glass rod, leaving a spot of 4-5 mm on the slide. Repeat this for as many slides as you wish to make. Getting the proportions of wax and glycerin right is important: too little paraffin and too much glycerin will result in an incomplete seal, too much wax and too little glycerin will result in nematodes being covered or trapped by paraffin.

2. Transfering worms

Pick out the specimens you want with a needle and transfer them to the glycerin drop in the center of a wax-ringed glass slide. You can usually mount up to ten of them per slide; more will too often result in specimens overlapping or ending up in paraffin. After transferring the required number to a slide, put it under the stereo microscope and push all nematodes to the bottom of the glycerin drop with the pick, making sure none overlap with one another.

3. Sealing and shuffling

Drop a coverslip over the wax ring and glycerin drop, and put the slide on a moderately hot plate, or a mesh or metal plate above a small flame. Make sure one end of the slide sticks out over the rim of the plate, or you'll get fried fingers trying to pick it up again! Allow the paraffin to melt around the glycerin drop, and allow all air to escape from under the coverslip. Then put the slide back under the stereo microscope, and check that no nematodes are overlapping. If so, gently push the coverslip in the required direction to dislodge one of the overlapping nematodes. If the paraffin has set by now, return the slide to the hot plate. You can also re-heat and gently push the coverslip sideways to turn specimens over. Once set, the paraffin will both act as a seal and a separating layer between the coverslip and the glass slide, and ideally your slide will contain just a small circular central area with glycerin and nematodes.

If some specimens are covered by smudges of paraffin under the coverslip, and/or the paraffin is too thick to observe specimens with high power objectives, put the slide back on the hot plate and allow the wax to heat and spread out further so that it forms a thinner layer. If you want to pick out specimens for transfer to another slide or for use in SEM or cross-sections, gently prise the coverslip open with a scalpel or thin needle while keeping track of your specimen(s) under the stereo microscope. This may go better if you first heat the slide gently (e.g. leave it on the lamp housing of the microscope for a few seconds). Make sure you do not allow the glycerin to boil during any of these operations, and never push down on the coverslip while the wax is molten.

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