WORMWOODWORMWOODCompiled by Paul De Ley -WWW pages maintained by Mark Blaxter and David Giuliano
This page explains some relatively simple methods for culturing a range of bacterial-feeding nematodes from soils and sediments. Click here for references on specific C. elegans culture methods. (30 November 1995)
You can also find a very rough guide to major groups of soil nematodes.
Many kinds of equipment, media and methods can be used to grow nematodes, and the choice of one over the other depends entirely on your research aims, available time and tools, target species and personal fancy. Where applicable, it is often a good idea to use the relatively standardized methods used for Caenorhabditis elegans, because this may allow you more easily to exchange data and cultures with C. elegans researchers, consult C. elegans literature, engage in similar work as the C. elegans community (genetics, neurology, development, etc.) and compare your own findings directly with what is known of C. elegans. However, C. elegans methods often require specific chemicals and tools that may not be available to you, or which you may not really need for your own investigations anyway.
Culturing nematodes is relatively simple, but success is not guaranteed. A wide range of species can be kept in the laboratory, but they may have specific requirements that can only be discovered by extensive experimentation or chance. Also, be aware that starting and maintaining cultures is labour-intensive, especially so if you want to maintain more than just a few species. And finally, all steps of the procedure can be influenced by chance events beyond your control or knowledge. Some groups of species are nearly impossible to stop from growing on agar, while others must be teased along very patiently until a reasonable number is available for subculturing. Whenever you have to open a plate, contaminants may invade, sometimes with fatal effects to the nematodes. On the other hand, maintaining an organism in culture results in artificial selection for its survival on the medium used. Adaptation may occur at any point in time, with a sudden improvement in culture efficiency as a result. And if you have enough specimens to risk some experimentation with different methods, it may be a good idea to try other media/bacteria/handling methods to see if culture efficiency can be improved.
This text outlines an assortment of simple and less simple handling and growing methods for use in a broad range of laboratory conditions and on a broad range of species. In general, you will need to process your cultures in three steps:
In practice, these steps need not be separate from one another, e.g. when a species multiplies well from the very first transfer to a plate. But other species may take a lot of care and patience, and for these it is quite important to be aware of the distinct aims during each stage of the process. Thus, a seed culture is not the same as a mass or stock culture because it may take quite some time before you can harvest enough specimens for research and/or before you are sure your strains can be kept indefinitely in the lab. And a mass culture is usually quite different from a seed or stock culture, because the very factors required to grow large numbers of specimens (optimum abiotic factors and food supply) will typically favour contaminants as well (that may kill off your target species), or lead to a sudden crash after peak density, which is clearly not good for long-term maintenance of your strains.
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Any extraction method that yields live nematodes at the end of the procedure can be used, but it is always best to have lots of healthy specimens of your target species. The most convenient method is usually a tray extraction (see the Extracting, fixing and mounting worms page), because this allows you to handle sufficiently large samples (at least if your tray is big enough) and because this generally results in minimal mortality of the nematodes. Unlike trays, Baermann funnels often need to stand more than one day (resulting in a larger number of asphyxiated worms) and may need to be tapped repeatedly (which requires you to spread out subsequent handling over a longer time and in repetitive steps). Flotation-centrifugation typically yields drastically lower proportions of healthy worms, even with less damaging and more expensive separation solutions. Elutriation does not damage the nematodes nearly as much, but obviously requires an elutriator, which may be quite expensive.
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It is not necessary to add bacteria to cultures if you plan to use natural associates of your nematodes extracted from the same soil sample(s). However, this will limit your scope for improvisation in handling subcultures, because you will have to ensure that you can maintain the bacteria along with the nematodes in the same plates, or else that you can culture these bacteria separately on plates or in flasks. In most cases, this will not save you work or worries, and it is often easier to culture Escherichia coli as a food supply for the nematodes. Many (but not all) bacterivorous species do well on E. coli, and E. coli is very easy to maintain and store, with no risks to yourself or other lifeforms in your lab. Be aware, however, that your bacterial culture is the most likely source of contaminations in your nematode culture!
Commonly used E. coli strains are e.g. 9001 or OP50, of which the latter is a slightly slower-growing strain, less likely to overwhelm the nematodes in rich media. These strains can be obtained free from any C. elegans lab as well as many other places. The main requirement in culturing this (or any other) species of bacteria is a maximal effort to avoid contamination. This means all labware should be sterilized, and if possible all work should be done on a flowbench.
E. coli is typically cultured on various simple liquid media, and stored in the fridge or deep-freezer. One common medium is "Luria Broth" (LB) which can be bought as a dehydrated powder from biosciences suppliers in various standard formulations. LB medium is prepared by adding the recommended amount of powder to distilled water (usually about 16g/liter), stirring well until all powder is suspended, and autoclaving for 10-20 minutes. The resulting amber solution is sterile and (after cooling) ready for inoculation with E. coli. According to anticipated requirements, fill a number of sterile culture tubes or flasks with respectively 5-10 or 20-50 ml of LB, and inoculate E. coli with a flamed platinum loop or another sterile inoculation tool. If you have a shaker-incubator, place the tubes or flasks in it at 37°C and leave them overnight at a moderately fast rotation speed (e.g. 100-150 rpm). It is best for aeration to leave flasks capped but unsealed, at least if you are confident there is little risk of contamination inside the incubator and you are inoculating a large volume of medium. For tubes, the volume of air in each tube should suffice for bacterial growth and it is safest to seal the cap.
The bacterial culture will be ready for use next day; if it is not used straightaway it should be stored at 4°C. Liquid cultures can be kept refrigerated for weeks, but it is best to make fresh cultures at least once a month. Also, try to finish each tube or flask in one go; repeated opening and pipetting out will inevitably lead to contamination. For long-term storage, you can pipette drops of the E. coli liquid culture straight into liquid nitrogen and deep-freeze the resulting frozen globules infintely at -80°C or in liquid nitrogen.
To check and maintain purity of your stock of E. coli, smear out small drops from liquid culture with a platinum loop on LB agar 1% plates, and leave these for two days at 37 C. Pure E. coli should give milky, homogenous culture smears or plaques with smooth edges. Any other colour or appearance means trouble, betraying e.g. toxic bacteria that can first stop nematode reproduction and eventually kill all remaining adults, or fungal hyphae that will outcompete the nematodes and bacteria for nutrients and modify the chemical conditions of your cultures. If your stock E. coli plates are clean, store them at 4°C for up to one month and use only these for future liquid cultures. If not, get new stock from someone who has a clean culture. If you have recurring trouble with contamination of your E. coli stock, try using one of the many available antibiotic-resistant strains and culture them in the presence of the appropriate antibiotic.
One strange alternative for E. coli as food source is the use of an agar-lysing bacterial species (e.g. an unidentified natural associate of Rhabditis oxycerca). Such bacteria will digest the agar itself, resulting in completely liquid nematode cultures in a few weeks time. At least some of these strains appear to be quite nutritious food for many nematodes, and their use dispenses with any need for separate culture media and vials for the bacteria. If your seed culture plates liquefy spontaneously, you have extracted one of these bacteria along with the nematodes, and can consider maintaining it for further use as food organism - at least if you don't mind starting with solid and ending with liquid cultures in each plate.
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It is best to prepare agar plates one day before freshly extracted nematodes will be available from soil. In the case of a tray extraction, this means you can start up the extraction and pour plates on the same day. Store excess plates at 4°C for later use, but note that seed cultures are always best started on freshly made plates.
Because a seed culture has to allow growth of target nematode species with minimal stimulus to competing contaminants or other nematode species, seed culture plates are best made of plain agar without any added nutrients. Depending on the species you wish to culture, a harder or softer gel may be required, but a 1% gel is often a good compromise. Do not bother about using ultrapure agar and deionised or twice distilled water, because these will reduce the amount of trace elements available to the nematodes. In fact, tap water and cheap plain agar usually do quite well. Also, it is known that C. elegans is unable to build its own steroles, and until proven otherwise you should assume other species are similarly impaired. It is therefore best to add a small amount of cholesterol to the agar (stock solution 5g/l in pure ethanol; add to a final concentration of 5 µg/ml after agar is autoclaved and cooled to 50-60°C).
If you are targeting Rhabditida or Diplogasterida, it is usually sufficient to prepare fresh agar plates with an E. coli plaque: after pouring and setting of the plates, administer 0.3-0.4 ml liquid culture E. coli onto each plate, and leave them overnight at room temperature (sealed or unsealed depending on contamination risk and whether some evaporation is desired or not). In this text, such plates will henceforth be referred to as "dry plates". For later washing of dry plates and handling of the nematodes, prepare 100-500 ml of S-buffer (0.05 M phosphate buffered to pH 6; 0.1 M NaCl) and autoclave.
If you are going for more sensitive and/or slow-growing species, especially those from freshwater sediments or wet soils, it is best to make "wet plates". A wet plate consists of a set agar plate to which you have added buffered water, and is convenient in three respects:
After setting up the extraction tray, it is a good idea to measure the pH of the suspension a few hours later, and prepare a buffered solution with the same pH. Alternatively, you can make up a large stock of slightly alkalic buffer and hope that this will not unduly stress the nematodes. Do not use standard S-buffer because this is actually quite saline and will pickle sensitive species even if they do survive a pH shock. 400 ml of good general-purpose 0.05M P-buffer (phosphate-buffer) is e.g. made by adding the following to 380 ml distilled water:
More alkalic buffer can be made by adding small amounts of a KH2PO4 solution to 0.05M K3PO4 or Na3PO4. Shake, autoclave, cool and pipette onto the plates (4-5 ml buffer on a 9 cm plate, 1-2 ml on a 5-6 cm plate). Administer 3-5 drops of E. coli liquid culture onto each plate, stir until E. coli and P-buffer are spread evenly over the whole plate, and leave overnight at room temperature. Instead of P-buffer, you can also use autoclaved water from the extract, which will not subject the nematodes to an abrupt change in chemical conditions. But extract water may not buffer adequately, so it can be more subject to pH increase over time as metabolites accumulate in the culture. Also, since you do not know the exact composition of the extract water you may have trouble later on when your stock of it runs out.
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250 ml soil on a large extraction tray will often yield a plentiful abundance of nematodes after just one day. Concentrate the extract to a volume of about 20-30 ml with sieves or a sequence of ever smaller settling beakers. Stir this final volume and pour into a counting dish for examination under the stereo microscope.
Now is the time to pick nematodes from the extract and inoculate the plates (see the Extracting, fixing and mounting worms page). Depending on your research aims, you may be happy with just any old bacterial feeder, or you may be looking for a very specific genus or species. In the former case, just transfer a reasonably large number of nematodes with a pasteur pipette, and you can be fairly sure there will be sufficient specimens from easy-growing species to get your culture going. If you are looking for specific nematode taxa, on the other hand, you will need to recognize them by their movement and general body shape under the stereo microscope or inverted microscope. This means you should use optics with fairly high maximum magnification (e.g. 15x oculars and 50x objective), or even transfer specimens to live mounts (see the Extracting, fixing and mounting worms page).
Here are some very rough guidelines to the main groups of soil nematodes. For approximate identification to lower levels, try the key to freshwater nematodes in the Plant and Insect Nematodes WWW site. This key contains some, but not all genera found in soil.
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Many Rhabditida and Diplogasterida are quite hardy and have a high multiplication rate due to short generation time and/or large egg production. They can usually be cultured easily on dry plates. If you transferred secernenteans to a dry plate with a Pasteur pipette, you may want to evaporate the transferred water by leaving the plate half-covered near the filter on a blowing flowbench (and make sure you don't leave your plate open for too long). Most adenophorean bacterial feeders (Chromadorida, Araeolaimida, Monhysterida and others) are apparently much more sensitive to environmental changes, and are best kept on wet plates. If you're not too sure about the preferences of your nematodes, it is often best to try keeping them on wet plates until you have enough material to experiment with different conditions. Or you can use a "semi-wet plate", i.e. a plate left overnight with P-buffer and some drops of E. coli from which you pour away the standing water before inoculating nematodes. Thus, a semi-wet plate has a moist surface covered by a thin layer of E. coli, but little or no standing water. Always SEAL YOUR PLATES with parafilm when you have finished handling them.
As mentioned above, wet plates offer better protection against desiccation and pH change than dry plates. They also minimise burrowing (suspended nematodes cannot find a foothold to dig into the agar) and allow you to transfer nematodes simply by pipetting. Main drawbacks are the ease of growing and transferring contaminants (especially protozoans), and the impossibility to use nutrient-rich agars. The latter is precluded by the risk of suffocating your nematodes in a soup of teeming bacteria using up all oxygen in the water layer. In theory (but not yet found in practice) it is also possible that wet plates lower reproduction of bisexual species because they may make it more difficult for males to search and find females. Individual nematodes can be picked from all types of plates with a fine needle or eyelash, the traditional method of handling, which requires practice, exposes the nematodes to air for at least five seconds (enough to kill or maim very sensitive ones), and can make you squish specimens or push them into the agar. A generally faster and safer method of picking single specimens from wet plates consists of using a micropipette with a non-sticky tip or a Pasteur pipette with mouth tube (see the Extracting, fixing and mounting worms page).
Dry plates are obviously forbidden for nematodes very sensitive to desiccation, although these may survive in a very soft gel. If you want to avoid burrowing of the worms into dry plates, you have to use a harder agar (1.5-2%) and avoid damaging its surface when picking worms (e.g. use a needle with flattened instead of sharp tip), because the smallest break in the surface will allow them to burrow down. Dry plates allow for much better bacterial growth, however, and are therefore most useful for growing larger numbers of Rhabditida and Diplogasterida. Single worms have to be picked FROM dry plates with a needle or eyelash, but can be transferred TO dry plates with a pipette (if you pick them from extract or wet plates). The most efficient way of transferring large numbers of nematodes is by washing them off the surface with buffer and pipetting the suspension.
If the nematodes have dug in despite your efforts to prevent this, you can extract them with more patience and less efficiency in a small dish with filter paper: cut the agar up, put it on the filter paper and fill the dish with (sterile) water. After 8-10 hours, a significant proportion of the worms should have collected on the bottom of the dish under the filter paper. You can also cut out blocks of agar with specimens inside and transfer these to new plates with a scalpel.
Semi-wet plates combine features of dry and wet plates; they allow better mobility and better feeding of the worms than wet plates, and still reduce risks of desiccation. They usually make it somewhat difficult to pick specimens with a (micro)pipette, as you need to look for places with a sufficiently thick water film.
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A seed culture is doing well if you can see evidence of growth and reproduction after one week. In order of increasing success, this means seeing: feeding animals with dark intestinal contents, females carrying eggs, nematode eggs lying on the agar, first-stage juveniles actively crawling around. If you have a very succesful seed culture (e.g. large numbers of nematodes after only one week) you should transfer to new plates immediately, because fast-reproducing species also tend to crash very quickly.
If nematodes survive but multiply very slowly, you may be in for a long process of gently teasing up their numbers. Once large numbers are present, inoculating them to fresh plates will no longer be a problem, because even the longest generation time can be offset by a sufficiently large number of reproducing animals. But until then you will have to proceed very cautiously and patiently, because any hurried attempts to push up their numbers may well have the opposite effect. Try to determine whether they are simply restricted by food supply, e.g. by transferring just a few adults to a plate with more E. coli, or to a plate with bacteria from the soil extract. It is more likely, however, that they will be inhibited by a combination of slow reproduction rate and physiological requirements, and this may leave you with very little freedom of action. NEVER put your whole "seed stock" at risk with a single operation, e.g. by transferring all you have to one fresh plate.
If the seed culture(s) of your target species fail, there is only one solution: try again, using different combinations of media, food source and types of plates.
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The easiest way of growing large numbers of nematodes of a certain species is by (1) making sure you have only one species, (2) starting up a monoxenic, synchronized population and (3) throwing lots of food at them. This usually works quite well with secernenteans, but sensitive species may not survive any of the above steps, and can only be grown in large numbers by applying patience and/or hoping for adaptation to occur.
Pick out gravid females and transfer one each to several fresh plates. A single female of a fast-growing parthenogenetic or hermaphroditic strain should produce ample progeny in just a week and present no problems for further manipulation and culturing. Pick the most succesful resulting culture and use this for all subsequent work and subculturing; mark all derived cultures with a symbol indicating that they contain progeny going back to a single female. Keep some of the other plates as spares, but make sure you only use them in case of emergency.
Fast-growing bisexual species will take somewhat longer to pick up in numbers from a single female, and plates will fail entirely if they contained a virgin female. Slow-growing species may not allow you to take this step at all, because it may take many months to build up reasonable numbers from a single female. In that case, fix as large a number of specimens as you can spare and try to ascertain that they are all conspecific by making slides and examining them under high power.
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This is a gentle way of saying that you need to kill all stages except eggs close to hatching. The standard way of doing this is with chlorox, a strongly alkalic and concentrated hypochlorite solution, which will kill all juveniles, adults and early embryos, before also affecting larvae with cuticle inside eggs. The latter survive a few minutes longer in chlorox because of their double protective layer (egg shell + cuticle), and if correctly timed can be recovered alive after all other stages and organisms have died. This aggressive procedure therefore has the double benefit of killing all contaminants, usually including bacteria in the intestines of juveniles and adults, as well as yielding a more or less synchronously developing population. After this procedure, you can inoculate the hatchlings on monoxenic plates, i.e. plates with just a single food organism (e.g. E. coli). For some applications you may even want to grow your worms in axenic medium, i.e. a mixture of organic molecules and minerals devoid of any other life than your nematodes. Axenic cultures are not dealt with here since they require extremely strict sterility procedures, as well as experimentation with appropriate mixtures of nutrients (see Vanfleteren, 1980).
Various modifications can be tried out, but what follows is a typical chloroxing procedure. Take one or two 9 cm plates with many nematode eggs on the agar and/or in gravid females. Pipette off eggs and nematodes (wet plate) or wash each plate with 5 ml appropriate buffer and then pipette off eggs and nematodes (dry plate). If necessary, use a glass spatula before pipetting to gently scrape off eggs and nematodes sticking to the agar. Transfer the nematodes to a 15 ml centrifuge tube and centrifuge 4 minutes at 3000 rpm and (if cooled centrifuge available) at 4°C. Suck off supernatant buffer with a pipette connected to a water suction pump. Prepare and mix 10 ml chlorox in the tube by adding quickly:
Leave the eggs and nematodes in this mixture for 7-10 minutes, and observe to check whether the nematodes die and dissolve properly. Then centrifuge again for 4 minutes at 3000 rpm and 4°C. Suck off the supernatant chlorox with the water pump and immediately add 10 ml appropriate buffer to wash the eggs. Mix, centrifuge, suck off and then repeat the wash, mix and spin. You now have a thoroughly sterilized suspension (hopefully) containing eggs close to hatching as the only living organisms. Leave the suspension in a dish for a few hours until you have numerous hatched juveniles, and then transfer them to a fresh plate with E. coli.
The method usually works well with Rhabditidae, but various problems may occur. Some dauers can survive the chloroxing (!) and carry over contamination as well as messing up synchronization. Juveniles and adults may die without dissolving, allowing bacteria to be carried over in the intestines of dead animals. Or ALL stages may die, including all eggs, which obviously was not your intention. In the former two cases, try using slightly longer chloroxing times, but note that you may lose the eggs as well if extending for too long. In the latter case, synchronizing is probably impossible using chlorox, and sterilization may also be difficult. Note that you cannot surface-sterilize most free-living nematodes, because they will indiscriminately swallow toxins and die (cephalobids may be an exception). A very dilute mixture of bleach (e.g. 1 part in 1000 or 2000) may sometimes work as a gentle sterilizing mix, but if you leave the nematodes in it for too long they will die, and if too short contaminants will still be alive.
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To force-feed nematodes effectively, it is important to sterilize them, so that no contaminants will outgrow and/or kill them in the enriched medium. It is also useful to have a synchronized population, because this will allow you to determine approximately what their generation time is, and schedule subcultures accordingly. Finally, try to determine the optimum temperature for each species, and keep them at it without any temperature shocks. Ideally, you should get plates covered in a writhing mass of worms after two generations. Various enriched agar media can be used to produce sufficient food bacteria for your mass culture, e.g. "nutrient agar" as supplied by various companies (usually with beef extract and peptones), or LB-agar (=agar made with LB instead of pure water). Dry plates have to be used to prevent asphyxiation of the nematodes due to bacterial growth.
Pour sufficient dry plates with e.g. nutrient agar (typically at 26 g NA per liter water) and cholesterol (final concentration 5(g/ml). After setting, pipette 0.3-0.4 ml of liquid E. coli culture to each plate. Add the synchronized hatchlings and avoiding leaving standing water on the plate (e.g. by allowing it to evaporate on the flowbench for 30 minutes to one hour). Seal the plates (some species will crawl out and explore your lab) and observe regularly over the next days. The best moment to harvest worms is when they are just about to finish the available bacteria, i.e. at the point where E. coli patches are no longer visible. Do not wait longer, because the nematodes will stop reproducing and start dying or transforming into dauer.
If you need to grow larger numbers, wash each plate in 5 ml of the appropriate buffer and pipette 1 ml each onto five fresh NA plates with E. coli patch. You can repeat this once or twice to reach enormous numbers, e.g. starting with one synchronized population inoculate 5 plates, then subculture to 25 and then to 125, which will probably give you more than 100 g of worms. To attain maximal biomass, you can then transfer them to large flasks, add one-fifth volume of LB and leave on a shaker two or three days at 120 rpm and optimal temperature until the suspension becomes milky white. At this point, all LB and E. coli are used up and most nematodes will be adult, i.e. at their maximal body size. Reproduction is often hampered in flasks, so they are only appropriate as a last step. Check Lewis & Fleming (1995) for more intensive large-scale methods.
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Losing a succesful culture can be quite a hangup, especially if it took a lot of work to get going. The best way of preventing such unpleasant surprises is to have live specimens frozen in a way that allows them to revive upon thawing. This requires an ultra-low freezer capable of going to -70/80 (C, because at -20°C ice still crystallizes and disrupts cells, while decomposition is not completely halted. And if you're even better equipped, you can transfer frozen specimens to liquid nitrogen to avoid the disastrous consequences of possible freezer failures. If such equipment is not available to you, you will have to keep stock cultures to preserve live strains. Some species can be kept dehydrated for some months, but their recovery drops exponentially over time, and you will need to revive them on fresh plates from time to time.
The main problem with freezing nematodes is that internal ice crystallization destroys their cells. You therefore need to minimize this crystallization, and the traditional method of freezing C. elegans relies on 15% glycerol to act as an anti-freeze. The method tends to be quite erratic in subsequent reviving success, because it depends on the capacity of glycerol to penetrate the nematode cuticle, and requires great speed to prevent the glycerol from actually killing the worms. It works as follows:
Prepare a stock of 30% glycerol in distilled water, get microtubes (eppies will do for -80°C, special plastics are needed for liquid nitrogen storage) and an insulated box allowing cooling at -1°C/min (e.g. a cardboard box with rockwool or a "Mr. Frosty" box with special insulating liquid). Wash nematodes from a plate containing lots of small juveniles (these have the thinnest cuticle and absorb glycerol easiest) with 5 ml S- or P-buffer. Transfer them to a centrifugeable tube and spin for 4 minutes at 3000 rpm and (if possible) 4°C. Remove supernatant buffer, and add 2.5 ml fresh buffer. Set up 5 microtubes and label them. Pipette 2.5 ml 30% glycerol into the tube with worms, mix briefly and use the same pipette to transfer 1 ml to each microtube. Transfer the microtubes to the insulated box and put it in the ultra low freezer. Once the worms are in glycerol, speed is of the utmost importance in this procedure!
The next day, you should remove one microtube, thaw it and empty on a fresh plate to determine the survival. It may take some hours for the nematodes to recover, and you should wait until the next day again to make sure they are really ok. If so, you can then transfer the other tubes to liquid nitrogen (if available) and sleep quietly.
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The first rule to keeping live stocks of your strains is to slow down the inevitable decline of each plate as much as possible. This means sealing plates well to stop desiccation - a good parafilm seal can last for over a year. Also, if you have cooling equipment keep the nematodes at lower temperatures than optimum, but above their minimal requirement for reproduction. This will slow down not only their growth and reproduction, but also their death and decay. Finally, use plain agar or another nutrient-poor medium: rich cultures produce lots of toxic metabolites and can crash quite suddenly. Some popular agar media (e.g. "NGM" used for C. elegans) seem to contain agents that precipitate death and decomposition, resulting in nematode corpses typically looking as if they have been scorched. Plain agar is better because it will keep the nematodes lean and fit for a much longer time, and in many cases they will become inactive and remain in cryptobiosis without dying and decomposing.
The second rule is that you should learn enough about each species to know its specific requirements. Slow-growing and/or inactive species can last quite long, and e.g. many cephalobids will survive in a sealed plate for well over six months. Fast-growing and/or highly active species (especially Rhabditidae) will burn out much more quickly, and may need subculturing every three or four weeks despite all your attempts to slow them down.
The third rule is to keep your eyes open. Maintain at least three separate plates for each strain, to have backups in case one plate fails or gets contaminated. Label plates clearly, including not only the strain name or code but also the date of inoculation and medium used. Check plates regularly (e.g. once a week) and subculture in time. In general, plates with immobile but intact nematodes are often still ok, but plates with large numbers of decomposing worms mean trouble.
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Some groups of nematodes are adapted to dry habitats and survive desiccation if it is gradual. This can be imitated in the lab to lower money and time spent on subculturing, and although this has not been tried yet it might theoretically also allow you to freeze species impermeable to glycerol (e.g. cephalobids). The simplest way of slowly desiccating a culture is by puncturing the sealing ring with a needle. If all goes well, the plate will take weeks to dry out and the worms will first coil up and then slowly lose their body water. It is imperative that they have not burrowed into the agar, because in that case they will be squashed by the gel as it slowly contracts. After drying the gel, you will be left with a paper-thin foil covered with curled-up and crumpled nematode mummies. Try out their recovery by cutting out a small piece of the dry gel and leaving it in water or buffer. If succesful, most of them should have revived by the next day. Dehydrated nematodes can theoretically revive after years, but in the laboratory it may be difficult to provide the optimum conditions for such long preservation, and it is safest to revive them and transfer them to a fresh plate after six months.
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