线虫基本操作方法

线虫实验指南Michael Koelle's C. elegans ProtocolsDOWNLOAD FROM:/mbb/koelle/protocol_list_page.htmlAugust 8, 2009Culturing Wormsby Michael Koelle1. Bacterial strain for feeding worms: OP50, a uracil auxotroph. Streak OP50 out on a 9 cm NGM agar plate, grow overnight at 37°. Can then parafilm the plate and keep it at 4° for months. To make bacteria for seeding plates, use a flamed wire loop to pick a single OP50 colony into a 100 ml bottle of B broth. Set the bottle (no shaking) at 37° overnight. It should be cloudy the next day. This bottle can be stored at 4° and used for ~3 months.2. Seeding plates. NGM plates should be allowed to sit out at room temperature in a closed box for a couple of days after pouring to allow them to dry, and so that plates with bacterial or fungal contaminants become obvious. Using a sterile pipette, drop 1-2 drops OP50 suspension on the middle of each 5 cm NGM plate. It is best if the bacterial lawn does not touch the side wall, as worms then tend to crawl up the wall and die. (Don't move the plates soon after seeding, or the puddle of bacteria will definitely slosh against the wall). Take care not to damage the surface of the agar with the pipette tip; worms will then burrow under the surface. Let the plates sit overnight to form a bacterial lawn before using. Plates can be used for about 7 days after seeding, and are best 2-4 days after seeding.To seed large (9cm) NGM plates: drop 5 drops OP50 in the middle of a plate. Use a sterile pipette at a very shallow angle to spread the OP50 liquid around so that a thin pool of liquid covers the middle 2/3 of the plate. Set the seeded plates at 37°overnight, or at room temp. for 2 days. (Many people try to seed large plates by dropping a huge amount of OP50 on the plates, until the thick pool of liquid covers a large portion of the plate; this results in a wet mess that takes several days to dry into a mealy-looking lawn.)3. Transferring worms. Use a platinum wire pick. To construct this, take a hammer and bang on a piece of ~32 gauge platinum wire to flatten it. Others use pliers to flatten the end of the wire. Then cut the wire so that only a few millimeters of flat wire are left attached to a longer section of round wire. Can use a razor blade to slice off the sharp corners of the flat end. Then bend the flat section so that it is at a shallow angle to the rest of the wire. Finally, place the non-flattened end inside of a short Pasteur pipette, and seal into the glass over a Bunsen burner. About 2 cm of wire should protrude from the glass, but personal preferences vary. Eventually the end of the pick wears out, and the end can be cut off and a new end flattened and bent.To pick up worms, flame the wire to sterilize it, and drag the pick along a bacterial lawn to coat the under side of the pick with bacteria. Goopy, several day old bacterial lawns are the best for this; some keep old plates next to the microscope especially for coating their picks. Gently brush against the worm to be picked up so that it sticks to the bacteria on the pick. To set the worm down, gently brush against the new plate, and allow the worm to crawl off. The worm should be immediately active on the new plate if it is undamaged. It is possible to pick up at least ten worms at a time, however, you don't want the worms to spend too much time on the pick, as they will dry out.4. Worms are typically cultured in a 20° room, with the plates upside down, stored in old scintillation vial boxes. The dividers in the boxes can be reconfigured to fit the 5 cm worm plates. The boxes and dividers are baked at 65° for >1 hour before use to sterilize . Worms can be cultured between 15° and 25°, and will grow slower or faster, respectively, at these temperatures. To keep a stock available for a long time without maintenance, parafilm the plate and set it at 15°. Can then recover the worms for ~5 months afterwards. The limiting factor is that the worms die eventually of dessication when the plate dries.If you're leaving town for a while a good way to keep your worms is to put the plates in a 12.5° C incubator. The worms will be almost in stasis, and will show only barely perceptable development after 1 week. Apparently prolonged incubation at 12.5° kills worm stocks by sterilizing them. One week is certainly safe, and after 2-3 weeks you'll probably still have some fertile worms.5. To transfer a stock to a new plate, typically you "chunk it out" instead of transferring worms with a pick. Keep small capped bottle of EtOH by your microscope, and a small spatula. Dip the spatula in EtOH, then place the spatula in the bunsen burner flame for ~15 seconds: the EtOH will burn off and then the spatula should get almost red hot. (If you just quickly let the EtOH burn off this won't sterilize the spatula, and you will always end up contaminating your worms). Dip the hot spatula back in the EtOH until the spattering and fizzling stops- this cools it off so the heat won't kill the worms. Then quickly pass the spatula through the flame to ignite the EtOH, which will burn off without heating the spatula much. Use the now sterile spatula to cut a ~1 cm square chunk of the old plate and place it upside down on the new plate. Worms will crawl out onto the fresh bacterial lawn.NGM agar: 3 g NaCl17 g agar2.5 g peptone1 ml cholesterol (5 mg/ml in EtOH)975 ml H20Autoclave, and then sterilly add the following, mixing after each addition:1 ml 1 M CaCl21 ml 1 M MgSO425 ml 1 M potassium phosphate pH 6Typically pour 5 and 9 cm plates. Store in plastic boxes with covers at room temperature for a couple days before use to allow the plates to dry.B broth: 10 gm bactotryptone5 gm NaCl1 liter dH20dissolve over bunsen burner.Then distribute to 100 ml bottles and autoclave.Freezing Wormsby Michael Koelle6/27/94I. The preferred method1. Wash worms off of 1 large plate or 3 small plates in 3 mls S Basal into a sterile 15 ml disposable centrifuge tube. Worms should be harvested off of just starved plates which are predominantly L1 and L2. This should be 1 day after the bacteria have been exhausted. The worm book says dauers don't freeze, but E. Jorgenson says he freezes from really old plates and it works fine. Rumor has it that if the plates aren't completely starved, the freezing won't work - i.e. food in the gut somehow prevents the worms from surviving. To wash the worms off plates, add ~2 mls S Basal to each small plate (w/sterile 5 ml glass pipette and a pasteur pipette bulb), swirl briefly to dislodge worms, and suck off the suspension with the sterile glass pipette and place into a 15 ml sterile plastic centrifuge tube.2. Spin down in a clinical centrifuge for ~30 sec. Remove all but 1.5 ml of the supernatant.3. Add 1.5 ml freezing solution, mix well, and aliquot 1 ml each into 3 sterile freezing vials (Nunc CryoTubes #363401).4. Freeze slowly to -80°. This is accomplished by placing the vials in a styrofoam rack (the kind that 15 ml disposable sterile centrifuge tubes come in), placing another inverted such rack on top of the first, fastening the two racks together with rubber bands, and placing in a -80° freezer.5. The next day, move two vials to a permanent location. Record the strain number, genotype, and comments in a computer database. We use "FileMaker Pro" software. Na An tells me that freezing in a liquid N2 freezer truly is preferable -80°; she gets excellent thaws from 15 year old vials stored in liquid N2, but less good thaws from old -80° vials.6. The third vial should be used for a test thaw. Take the vial out of the freezer, thaw quickly by holding in your hand or in a 37° water bath (leave in the bath only until the ice is gone so as not to heat it up). Can dump the whole vial out in a seeded large plate, or use a sterile 200µl pipetteman tip to withdraw the bottom 200µl (containing the settled worms) and place it on a small plate. The next day, pick live worms to a new plate.II. The agarose method. This gives much lower viability than the above method in most people's hands, but has the advantage that one doesn't need to thaw an entire vial at a time. On balance, I don't think it is worth risking losing strains (especially risky with heterozygous mixtures) to get this minor savings. However, some people seem to have better luck with this method than me. Leon Avery says the agar method works better for him than the non-agar method. He suggests being especially careful when thawing not to break the worms by scraping, but rather to be sure to actually take an unbroken frozen chunk of worm suspension out of the tube.1. Melt freezing solution + agarose in microwave (carefull to fully melt but not boil over).2. Put FS + agar bottle on the bench to cool. Some people place it in 50° water bath for ~15'. The idea is to have it cooled to ~50° by the time you use it, but to use it while it is still liquid.3. Wash worms off of 1 large plate or 3 small plates in 3 mls S Basal into a sterile 15 ml disposable centrifuge tube. Worms should be harvested off of just starved plates which are predominantly L1 and L2. This should be 1 day after the bacteria have been exhausted. The worm book says dauers don't freeze, but E. Jorgenson says he freezes from really old plates and it works fine. To wash the worms off plates, add ~2 mls S Basal to each small plate, swirl briefly to dislodge worms, and suck off suspension with a sterile glass pipette and place into a 15 ml sterile plastic centrifuge tube.5. Chill on ice 15' or longer.4. Spin the worms down (30 sec. in a clinical centrifuge), and remove all but 3 mls of supernatant with a sterile pipette.6. Pipette 3 ml FS + agar into a tube of worms and immediately mix by inversion of tube 8-10X.7. Pipette 1.8 ml of the mixture into 3 prepared freezer vials.8. Repeat steps 6 and 7 for each strain.9. Freeze slowly to -80°. This is accomplished by placing the vials in a styrofoam rack (the kind that 15 ml disposable sterile centrifuge tubes come in), placing another inverted such rack on top of the first, taping them tightly together, and placing in a-80° freezer.10. The next day, move two vials to a permanent location. (Or, to a box in the -80°, and when the box is full, move all the vials to a liquid N2 freezer). Record the strain number, genotype, and comments in a computer database.11. The third vial should be used for a test thaw. Can either thaw the whole vial and dump on a large plate. Or, using a flamed spatula, dig a ~0.1 ml chunk of frozen worms onto a small plate, and put the remainder of the vial back in the freezer. Leon Avery says to be sure not to scrape flakes of frozen stuff out of the tube; this will break all the frozen worms. Instead, run the spatula around the edge of the tube and break out an intact chunk of worm suspension.Freezing Solution200 ml 1M NaCl100 ml 1M KPO4, pH 6.0600 ml glycerolBring to 2 liter w/ dH20Distribute to 200 ml bottles; autoclaveAdd 0.06 ml sterile 1M MgSO4 per 200 ml bottle(N.B. in Horvitz lab the stuff Na An supplies does not have MgSO4 added yet!) Freezing solution with agarosefreezing solution with 0.4 g agarose added per 100 ml and reautoclaved for 20 min.M9 buffer (Horvitz lab version)Na2HPO4 5.8 gKH2PO4 3.0 gNaCl 0.5 gNH4Cl 1.0 g dH20 to 1lResponse to food assayby Meng-Qiu Dong 1/1/2000Wild type worms adjust egg-laying behavior in response to food. They lay eggs when they are fed and stop laying eggs when they are starved. If starved wild type worms are put back on food, they resume egg laying almost immediately. I use the following assay to determine how well worms regulate egg laying in response to food quantitatively.Measure the # of eggs laid by 10 worms in 30 min under three conditions:1, starved; 2, starved then re-fed; 3, never starved.Note:∙Transfer of worms in liquid should be done with a glass pasteur pipette, not plastic pipette to which worms stick.∙Always use dry and non-cracked NGM plates. The plates need to quickly absorb a few drops of liquid placed on them.1. Pick large dark-looking late L4 larvae, ~60 L4s/plate x 2 plates. L4s are recognized by a white crescent in the presumptive vulval region, which acquires a black central dot in late L4. Worms need to be staged as precisely as possible, because this seems to reduce fluctuation.2. Incubate worms at 20 ºC for 30 hrs.3. Wash worms off plates with 2.5 ml M9 buffer at a time x 2 times, combine the washes in a 15 ml falcon tube.4. Spin down worms in a clinic centrifuge for 10 sec at maximal speed.5. Remove supernatant by aspiration, leave ~0.5 ml liquid. (so you don't lose worms during washing)6. Rinse worms with ~7 ml M9, spin 10 sec, remove supernatant by aspiration and leave about 0.5 ml liquid.7. Then carefully remove as much sup as possible with a pasteur pipette, usually about 100ul liquid is left in the tube.8. Take a 5 cm NGM plate seeded with a lawn of O.P.50 bacteria, drop on the edge of the agar 2-3 drops of 4M fructose from a pasteur pipette and rotate the plate so that the solution flows along the very edge and makes a fructose circle. Take an unseeded plate and do the same to circle the edge with fructose. You can do step 7 while you are waiting for the centrifuge to stop. The 4M frunctose creates an osmotic barrier that prevents worms from crawling off the plate and dying.9. Under a dissecting scope, use a pasteur pipette to transfer 20-30 worms to the seeded plate from step 7. I count the # of worms transferred because I don't want to waste too many worms on this plate. Put worms down near the bacterial lawn and let the liquid be absorbed into the plate. I come back to this plate a few minutes later and move any worms that are off food back on food using a worm picking coated with bacteria. Transfer the remaining ~90 worms to the unseeded plate from step 7.10. Incubate the worms at 20 ºC for 2 hrs. Near the end of the 2 hrs, prepare 6 fructose circled plates (4 seeded + 2 unseeded) in the same way as step 7.11. Transfer 20 starved worms to 2 unseeded plates from step 9 with a bare platinum worm pick (no bacteria!), 10 worms/plate. (Here is my way doing it: Use a short flat-ended worm pick, press against the agar beside a worm so that some liquid gets squeezed out to lubricate. Shove the worm pick underneath the worm and pick it up. On a new plate put down the worm pick [with a worm on top of it] against the agar and squeeze out some liquid. The worm will float up and swim off. Don't tear the agar surface when you put down a worm, just make a dent.)12. Transfer 20 starved worms to 2 seeded plates with a worm pick coated with bacteria, 10 worms/plate.13. Transfer 20 nonstarved worms to 2 seeded plates with a worm pick coated with bacteria, 10 worms/plate.14. Wait for 30 min at room temperate (should be close to 20 ºC). Then count the # of eggs laid on each plate. You can just remove the worms and count eggs later if you don't have enough time. Maximum of 3 strains can be assayed at a time.15. Repeat the procedure 4 times (8 repetitions total) and average the results. For unknown reasons, there is some fluctuation in this assay that requires this number of repetitions to average out.M9 (without glucose or MgSO4):MRC recipe; is basically M9 for bacteria without 20% glucose. used as M9 in Horvitz lab, e.g. for egg-laying assays, everything else. Not same as M9 buffer in worm book.Na2HPO4 5.8 gKH2PO4 3.0 gNaCl 0.5 gNH4Cl 1.0 gdH20 to 1lM9 buffer (Worm book recipe):also MRC recipe; Is basically M9 buffer with high NaCl to make up osmolarity for no glucose. Not used in Horvitz lab.KH2PO4 3.0 gNa2HPO4 6.0 gNaCl 5.0 g1M MgSO4 1.0 mldH20 to 1lN2 development times at different temperaturesby Michael KoelleDetermined empirically; times in hours are given from the first division of the zygote.L4's can be easily recognizedand staged to within a few hoursof development. By picking L4'sand then aging them, a fairlysynchronous population of adultsof known age can be obtained;this is especially useful forbehavioural assays, the results ofwhich may depend on age.L4 hermaphordites are recognized by a white crescent shape at the location of thedeveloping vulva, about 2/3 of the way towards the posterior of the animal, on it's"side" as it lies on the plate. This stands out in contrast to the darker yellow gray ofthe surrounding body. Over time, the L4's get fatter relative to their length, a dark spotappears in the middle of the white crescent, and finally the white crescent collapsesprior to the L4 molt. Thus substages of L4 are recognizable.L4 males are recognized by a tail that is just beginning to acquire a spade shape,but has not achieved the well defined sharply cornered spade-tail of the adult. Sinceyoung males are better at mating, it is best to set up matings with L4 males.Liquid culture of wormsBy Michael Koelle and Tory Herman, adapted from Mir Hengartner4/6/94Media1) superbroth (5 X 2 1/2 liters autoclaved in 6 liter flasks)5 X 30 g Bactotryptone5 X 60 g yeast extract5 X 20 mL 50% glycerol stock solution5 X 2.25 liters DDWHint: use a funnel to pour measured ingredients into the flasks.Autoclave 30 minutes. Cool to < 60 °C.Then add 250 mL of sterile 0.17M KH2PO4, 0.72M K2HPO4 to each flask.To prepare 0.17M KH2PO4, 0.72M K2HPO4 :46.2g KH2PO4250.8g K2HPO4 per 2 liters; make 250 mL aliquots and autoclave.2) S- medium (4 X 500mls autoclaved in 2 liter flask)0.5 L 2 L2.9g 11.6 g NaCl25 mL 100 mL 1M potassium phosphate, pH 6.0136.1 g KH2PO4 per 1000 mL; start with 800 mL and adjust to pH 6.0 with solid KOH (approx. 15g) before bringing up to volume.475 mL 1.9 L DDW0.5 mL 2 mL 5 mg/mL cholesterol in 95% EtOHWarm to 37°C O/N to dissolve. When this is added to an aqueous solution some cholesterol will precipitate; don't worry about this.Aliquot to four 2L flasks; Swirl to disperse; autoclave.Sterilely supplement EACH 500 mls with:1.5 mL 1M MgSO43 mL 0.5M CaCl25 mL 100X trace metals solution0.346 g FeSO4.7H200.930 g Na2EDTA0.098 g MnCL2.4H200.144 g ZnS04.7H200.012 g CuSO4.5H20 per 500 mLAutoclave. Keep in dark (wrap in aluminum foil).5 mL 1M KCitrate, pH 6.021.01 g citric acid, monohydrate per 100 mL; start with 80 mL and adjust to pH 6.0 with solid KOH (approx. 17 g) before bringing up to volume. Autoclave to sterilize.5 mL Gibco 100X Pen/Strp/Neo (, buy from Gibco, keep in freezer)5 mL 100X Nystatin (buy from Gibco, keep in freezer).3) worm plates- approximately 5 large plates of N2 (more for sick mutants) are needed to start each liter of liquid culture.Grow until the plates almost starve, or if you want to stage the worms, grow until the plates starve (they'll be mostly L1s).GROWING THE BACTERIA (WORM FOOD)1) Prepare 5 x 2.5 L of Superbroth in 5 x 6 L flasks (see above for recipe)2) Innoculate ~5 mL of HB101 in Superbroth into each of the 5 6-liter flasks; grow O/N @ 37° C with shaking.Warning: Be sure to have enough worm food! It is very bad to run out of food in the middle of a growth.3) Spin down the bacteria in 1-liter jars in the J6B centifuge (15 min / 4000 rpm / 4°C). This will require two spins of 6 bottles each ; just pour off the first sup. and add the rest of the culture on top of the first pellets and spin again. Resuspend the combined pellet in a bit (3 mL per bottle works fine) of S Medium. A convenient method is to put the 1-liter jar back on a platform shaker for ~15 minutes. Transfer to a 50 mL polypropylene tube and store in fridge for up to 2 weeks, or store @ -20 or -80 °C for unlimited time. The yield of bacteria per volume of superbroth is disappointing in these large cultures relative to what you would get in a 2 ml tube, presumably because of aeration problems. We have been getting about 9 mls of this bacterial suspension per liter of culture.GROWING THE WORMSStart with 5-8 almost starved large plates of N2 (more of mutant) containing a mixture of adults and young larvae. Wash the worms off the plates with S medium, and wash the plates again with more S medium to be sure to get all the worms. Add the worms to 1 liter of S-medium (in two 2 liter flasks containing 500 ml each.) Add 12.5 mls of bacteria to each flask. Shake @ 20° C on a platform shaker at ~240 RPM. You may need to start the shaker at slower RPMs and slowly turn it up to 240 in order to get the liquid swirling correctly in the flasks. Take approx. 200 µl aliquot every day and dump it on an unseeded worm plate to check that the worms are growing and whether they need more food. After a couple of days you will see tons of oval browinish pellets in the culture, presumbly worm debris and/or waste. If dauers are forming you need more food. Our regimen has been to add 25 mls more of bacteria to each flask after two days, and to harvest the culture 4-4.5 days after it was started. If there aren't enough worms, it is possible to wait 1-3 extra days to harvest. Some mutants grow more slowly than N2 and will definitely require these extra days of growth.Judging when the culture is ready to harvest takes a bit of experience. In the very best cultures, the bacteria are beginning to clear at the time of harvest (culture turns from milky to a clearer yellowish greenish brownish, and bacteria are less evident under the microscope), the worms are extremely dense, all stages of worms are present, and no dauers visible. If the worms get too dense all the worms will go dauer even if the food hasn't run out due to the accumulation of dauer pheromone. Of couirse, if the food runs out, the worms will also go dauer. Looking at 200 µl of liquid culture dumped onto an unseeded plate, when the culture is ready to harvest you should see about as many worms as you would find on a plate of worms (grown normally, on a seeded plate) that is about 0.5-1 days before starving.AFTER THE CULTURE HAS GROWN0) Prepare some ice cold 0.1 M NaCl (500 ml), and ice cold 60% sucrose (100 ml).1) Spin down worms in 3 500 mL jars 3K for 3 min in a Beckman JA-10 rotor. The speed and timing of this and the subsequent spins are fairly critical; we start the timing when the rotor is up to speed, then turn the machine off to start the deceleration when 3 minutes is up. Pour off the supernatant, being very careful and leaving some liquid in order not to lose any of the soft pellets. Expect a huge pellet with tons of bacteria, containing several different colored layers..2) Resuspend the pellets in 0.1 M NaCl, combine in one jar, and make up to 500 mls with 0.1 M Nacl. Spin again, this time at 2 K for 3 min in the JA-10, timing after the machine is up to speed as above. Pour off the supernatant, again being careful not to lose any, leaving some liquid.3) Swirl the flask to resuspend the pellet. Add more 0.1 M NaCl to make up to a total volume of 50 mls. Distribute 25 mls into each of two 50 ml centrifuge tubes, and place on ice for several minutes to chill. Add 25 mls ice cold 60% sucrose to each tube, mix, and spin 3.5 K for 5 minutes in a table top centrifuge. The sucrose solution damages and eventually kills the worms; work fast here.4) After the spin you should see a large dark brown pellet at the bottom of the tube (bacteria, debris, dead worms), and a large light brown layer of worms floating at the top. Use a broken off Pasteur pipette to carefully remove the worms, to a new centrifuge tube (it's ok to take about the top 20 mls of liquid here).5) Quickly dilute the worm suspension 4 fold with ice cold 0.1 M NaCl, and spin 3.1 K 3 min. Remove the supernatant. The pellet should contain reasonably clean worms.6) Resuspend the worms in 100 mls (i.e. fill up two 50 ml tubes) of ice cold 0.1 M NaCl, and spin 3.1 K for 3 min. to remove more sucrose. Pour the supernatant off the very soft pellet with great care. At this point the pellet should contain almost entirely healthy worms, with only a very small amount of debris and bacteria. The pellet can now be flash frozen in liquid nitrogen to later prepare RNA, or eggs can be prepped to start synchronized liquid cultures. Note that this last pellet is extremely soft, especially when it contains older worms, so it is impossible to pour off all the supernatant without losing the worms. Just do the best you can do, and freeze orproceed to prep the resulting suspension, which will be about 50% worms, 50% 0.1 M NaCl. If the worms will be used for an RNA prep, it is best to have less than 5 mls of worms in a 50 ml tube, to allow room to add the necessary guanidinium solution.7) The above prep takes two hours.8) Expected yield: about 15-20 mls of purified, packed, mixed stage worm pellet per liter of liquid culture. The contaminating bacteria should be <5% of the mass of the worms.PREPARING EGGS TO START SYNCHRONIZED LIQUID CULTURES1) Start with 20 mls of packed worms prepared as above. Make sure that this contains lots of gravid adults.2) Resuspend the worms in 1.5 N NaOH, 12% NaOCl, to a total volume of 100 ml (in two 50 ml centrifuge tubes). Mix gently at room temperature for 5 minutes. This will kill all the adults and larvae, but not the eggs, which are protected by their shells.3) Spin in a table top centrifuge 3.1 K for 5 min. Pour off the sup. and wash in 100 mls 0.1 M NaCl. Spin again, and wash again twice more in 50 mls 0.1 M NaCl each time, consolidating into one tube.4) Resuspend in 45 mls of S medium. Examining in the microscope at this point, you will see lots of flaccid dead worms, and very few free eggs. Most of the (viable) eggs are inside the dead adults.5) Our regimen has been to set up four 500 ml cultures from the eggs, and to harvest these synchronized cultures over the next 4 days to get variously staged worms. In order to get an equal mass of worms from the various stages we seed the cultures with 30, 9, 3, and 3 mls of the 45 ml egg suspension. These cultures are havested after 1, 2, 3, and 4 days respectively. Each 500 ml culture is initially fed with 25 mls of packed bacteria. Harvesting the first flask after 21 hours we got ~2 mls of packed L1/L2s (mostly L2s). Harvesting the second flask after 48 hours we got ~4 mls of packed L3s and early to mid L4s. Harvesting the 3rd flask after 72 hours we got ~ 4 mls packed adults, a mixture of ~2/3 young gravid adults and ~1/3 pregravid young adults.Lethal phase determination1. Pick a bunch of heterozygote L4 hermaphrodites to a new plate.2. The next day, there will be eggs on the plate; pick exactly 20 of these to each of a few plates. Make sure to use first day eggs like this: wild-type animals eventually run out of sperm and lay unfertilized eggs that don't develop, and you want to avoid confusing these with dead fertilized eggs. Line up the sets of 20 eggs in a row just beyond the edge of the seeded portion of the plate.3. The next day and succeeding days, examine the plates for unhatched eggs, dead worms, and live worms.4. If the worms hatch, but may be dying as young larvae, it can be difficult to distinguish between worms being "gone" because they died and the dead larvae are hard to find, or because live worms just crawled up the side of the dish and disappeared. To help this problem, you can keep the worms from crawling off the bacterial lawn by using a P-20 to ring the lawn with a line of 10 mg/ml palmitic acid in EtOH, which precipitates on the plate and forms a barrier the worms won't cross.。