Hill KL et al. (1996) Worm Breeder's Gazette "Fluorescent Staining of Live Worms Using SYTO Nucleic Acid-Binding Dyes"

L'Hernault SW, Hill KL

[Worm Breeder's Gazette, 1996]

We have been studying spermatogenesis-defective (Spe) worms that make spermatozoa and are defective in some aspect of fertilization. To understand the exact nature of the defect, we needed to develop tools that would allow us to study the behavior of these mutant spermatozoa. One tool we found to be especially useful is the SYTO vital nucleic acid dyes from Molecular Probes. Using these dyes, we have been able to label male sperm in vivo and subsequently observe them within the reproductive tracts of mated hermaphrodites after insemination. This has revealed that certain mutants contain spermatozoa that can migrate to the spermatheca and maintain their position as oocytes pass through to the uterus. This technique maybe of general use, for the following reasons: The dyes penetrate readily (with some variation between dyes) to label entire worms, including internal structures. There appear to be no serious effects on viability. Labelled males are able to mate efficiently, and their sperm are able to fertilize oocytes and produce apparently normal progeny. Labelled hermaphrodites are fertile as well, and their eggs hatch and produce healthy larvae. The dyes come in a range of excitation and emission maxima and therefore should be amenable to a variety of applications. We have had particularly good results with SYTO 17, which achieved the best penetration of the dyes we examined, and gives a brilliant red signal in the rhodamine channel. SYTO 25 behaves similarly, with an intense orange-red signal. Of the green dyes, (visualized in the fluorescein channel) SYTO 11-16 and 20-24, we found #13 to be most effective. To date, there are fourteen dyes, #11-18 and #20-25 from which to choose. Listed below is a protocol for using these dyes that has yielded consistent results. 1) Wash worms off plate with TBS using a cotton-plugged Pasteur pipette. Transfer to a 1.5 ml microcentrifuge tube. (TBS: 10mMTris, 1mM EDTA, 5mM NaCl, pH 7.4) 2) Spin worms to pellet. Remove supernatant rapidly to prevent redistribution of worms. 3) Prepare dye solution: 3 ul of 5 mM stock in 1497 ul TBS makes a 10 uM solution (be sure to check the stock concentration; some are 1 mM). Add dye to worms and mix well. 4) Transfer worms and dye to a clean, empty 3.5cm plate. Cover, seal with parafilm, wrap with aluminum foil and place in the appropriate incubator. 5) Incubate worms in dye for 3 hours. This is a general guideline; different dyes will need more or less time. 6) After incubation, transfer worms to a fresh microcentrifuge tube, spin to pellet and remove dye solution. This should be poured through activated charcoal to inactivate it for disposal. Resuspend in a few drops TBS and spot onto plates with bacteria for recovery. Once the plates are dry the worms can be picked. Because of the rapid photobleaching of these dyes, we have found it best to keep plates containing dyed worms shielded from the light as much as possible. These dyes can also be viewed by SIT camera technology, which allows at least 15 minutes of continuous (dim) illumination. In addition, we recommend double gloves when handling the dyes.

Ward S et al. (1980) Worm Breeder's Gazette "growth of large populations of synchronous worms in small volumes."

Hogan E, Ward S

[Worm Breeder's Gazette, 1980]

After much frustration trying to grow enough males for sperm biochemistry, we spent several weeks experimenting with liquid growth conditions and have improved our yields by a factor of ten. More than 10 ml of healthy adult worms can be obtained from a 130 ml culture. The key parameters appear to be adequate aeration and the absence of contamination. The first is achieved by using deep baffled shake flasks and the second by careful sterile technique. Our present protocol adapts previous procedures and methods published in the Newsletter (Hecht, Vol . 1, 2; Klass, Vol . 3, 1; Sulston and Brenner, Genetics 77: 95-104). 1. Growth of E. coli for worm food As noted previously (Newsletter 3, 2, p. 14) commercial bacteria are unsatisfactory in our hands. We grow our own in a Lab Line Model 29500, High-density Fermentor from which we obtain 180g wet weight of E. coli from 3L of culture. The baffled flasks described below would presumably give less yield per volume, but might work better than bubbling. E. coli strain NA22 is grown in the following medium: [See Figure 1] Add 3 ml trace metal solution (Sulston and Brenner, Genetics 77: 95- 104, 1974), 25 ml 1M MgS04. Grow overnight at 37 C. Harvest cells aseptically by centrifugation and resuspend in S medium at 2-3 X 10+E11 cells/ml (OD550 ~200). 2. Growth of Synchronized Worm Population and Isolation of Males General Comments Contamination of cultures by foreign bacteria or yeasts must be avoided, Tissue culture handling procedures are essential. These should include the following: 1. All glassware should be autoclaved. Growth flasks should be well sealed and dry. 2. Wash your hands with prep-o-dyne before handling cultures, especially for filtration. 3. Open flasks and cultures only in the laminar flow hood- Wipe hood surface with 70% ETOH before use. Flame all openings before transfers. 4. Use only newly opened sterile pipettes for adding to cultures or sampling. 5. Plug all flasks with sterile gauze-wrapped cotton plugs. Handle from outside only. Use sterile silicone rubber coverings for 1L and 2L flasks. Equipment and Solutions Needed Glassware - all sterile 1L and 2L baffled flasks (Bellco 2542 series) This is important 200ml screw cap centrifuge bottles (Bellco 3045 series) sterile cotton or silicone plugs for flasks sterile 25 and 10 ml pipettes. [See Figure 2] Distribute WG medium, 250 ml/bottle Bacteria - NA22 in WG or S medium at 2-3x10+E11 cells/ml. Uncontaminated. Solutions for washing and Isolating Worms 1.25 M-9 salts, autoclaved 30% Ficoll in 1.25 M-9 salts, autoclaved. Worms survive Ficoll floatation better than sucrose flotation. Filters Nitex nylon filters should be stretched tightly over 13 cm embroidery hoops with stainless steel screws. Label size in pencil on the surface. One 35 M and one 20 M will be needed for each 300 ml culture. Wash them out after use. Soak in 70X ETOH then set in sterile glass petri dish lids with support rods in UV hood. Keep them sterile. Starting Cultures For growth of males E1490, him-5(e1490) is the mutant of choice. Doubles with e1490 are used for most experiments with fer mutants. 1. Start from a clean, uncontaminated master plate. Transfer a chunk to a large growth plate. As this plate reaches starvation, rinse the larvae off with several changes of M-9 salts. Work in the laminar flow hood to minimize contamination. Transfer a plateful of worms to 100 ml of WG medium in a baffled flask. Add 10 ml bacteria. Grow two generations. isolate hermaphrodites and eggs as described in the following. If culture is not synchronous, isolate eggs and grow another generation. Preparing Eggs to Start Synchronous Culture 1. Concentrate adults by letting them settle 10-15 minutes or spin at 800 RPM at room temperature for 5 minutes. (About 400 x G). 2. Aspirate off bacterial supernatant. Resuspend in 1.25 x M-9 buffer. Transfer to 40 ml conical bottom tubes. 3. Repeat spin at 1/2 speed in clinical centrifuge 5 minutes. 4. Aspirate supernatant. Add ~25 ml of 30% Ficoll in 1.25 x M-9, mix. 5. Spin at 3/4 speed (~1000 RPM) in clinical for 10 minutes. 6. Withdraw worms from top of tube. Dilute with 1.25 M-9. Spin again at 1/2 speed 5 minutes. 7. Chlorox hermaphrodites. Make up fresh: 10 ml Bleach 25 ml 1N NaOH 15 ml H20 Add 20-25 ml to 3-4 ml hermaphrodites. Vortex occasionally for 6-9 minutes, until you see that all worms have dissolved. Spin down at 1/2 speed for 4 min. Aspirate chlorox sterilely. Dilute with WGM. Spin again. Resuspend WGM. Check a sample under the microscope to be sure all carcasses have dissolved. 8 ml of hermaphrodites should give 1.5 ml of eggs. 1 ml of eggs = ~4 x 10+E6 worms. 8. Place ~1 ml of eggs/250 ml of WG medium in 2L baffled flask. Add 40 ml bacteria. 9. Leave on shaking table at 20 C. Eberbach 25 mm diameter rotation. 150 RPM. The diameter of rotation is important, shaking water baths don't work well. Growth and Isolation of Males 1. Count the number of worms hatched 12-24 hrs after starting the culture and check their motility and condition. Smell the flask to detect contamination. If contamination is detected at this early stage it is probably best to clean the worms by centrifugation then flotation, return them to culture and rechlorox the hermaphrodites. 2. Adjust culture concentration from ~7500/ml to 25,000/ml. Add 12 ml bacteria for each 10+E6 worms total. We use 250 ml total volume in 2L flasks and 120ml in 1L. 3. After 48-60 hrs culture, feed again 12 ml bacteria/10+E6 worms. 4. After 80-96 hrs culture, spin down or let settle. Aspirate bacteria. Rinse with M-9 spinor settle again. Float with 30% Ficoll as in steps 2-6 of egg isolation. Keep culture sterile. 5. Have sterile 35 and 20 filters ready. 6. Distribute worms on 35 up to 10 ml worms per 13 cm filter. Add 1.25 x M-9 until liquid is even with bottom of filter. After 1/2 hr transfer filtrate to 20 . Add fresh 1.25 x M-9 and mix up worms on 35 filter. Check after ~1/2 hr to see what comes through. 7. If more males,add to 20 . After ~1 hr hermaphrodites will crawl through the 35 filter so don't add filtrate without checking. 8. Collect males with conical centrifuge tube from top of filter by rinsing with sterile 1.25 x M-9. This is the most difficult to keep sterile. Wash bands with prep-o-dyne before collection and handle worms as carefully as possible. 9. Spin down. Note volume. Resuspend in WG medium at <10,000/ml. At this age 2 ml males (= 6 x 10+E5) can go into 100 ml of culture. Add 10 g/ml Gentamycin to culture to minimize contaminants. Add 20 ml bacteria/10+E6 Worms. 10. Feed every 24 hours, ~20ml/10+E6 worms. 11. Prepare worms for squash 48 or 72 hrs after fractionation = 6 days or 7 days total growth at 20 C. 12. Spin worms and float as in egg isolation steps 2-6. Rinse final pellet in squash medium. Let males settle again before squashing. If there are many larvae or hermaphrodites, fractionate again in 35 and 20 filters.

Skimming J et al. (1984) Worm Breeder's Gazette "growing a lot of (ugh! smelly!) worms."

Skimming J, Lewig JA, McLafferty SA

[Worm Breeder's Gazette, 1984]

In our efforts to make monoclonals against the levamisole receptor, we've needed a passel of worms. Which means growing a helluva alot o' bacteria. 250 liter fermentors do not seem to abound in these parts and so we're unable to emulate the Johnson-Rand gang (1.5 kg of worms from a 250 L fermentor). We've turned to making the best of our Lab- Line S.M.S. fermentor (see an earlier article by S. Ward and E. Hogan, WBG 5 2, p.10). Using pure oxygen and a slightly modified version of the SLBH medium of Sadler, Miwa, Maas and Smith, Laboratory Practices 23, p.642, 1974, we've been able to grow 260 gm of E. coli per 3.3 liter of medium in a 7 hour fermentation run with very good reproducibility. A simplified version of the method described by Sadler et al. works just fine. We use all El Cheapo chemicals from Sigma for our SLBH medium. If you follow this method, be very careful with the oxygen. As you should know, it makes things highly combustible. Apart from the cost of the regulator and a tank rental charge, 244 cubic feet of oxygen cost us only $6.40 and it lasts 4 or 5 runs, giving us an average of 80 gm more a run than the 180 gm yields described by Ward using air. [see Figure 1] Use of the Sigma chemicals leads to a precipitate in the SLBH medium but otherwise things work just fine. To start things off, a 300 ml overnight culture is grown at 37 C in SLBH. We use E. coli strain NA22 whose genotype seems to be lost in the Dark Ages but the genotype does include a ts lethal so careful temperature control is important. The next morning the S.M.S. high density fermentor containing 3 liters of the complete SLBH medium is inoculated with the overnight culture and run at 320 rpm with an initial O2 flow rate of 1-2 liter per minute at 1 or 2 psi. Unlike the method of Sadler et al., all the glycerol needed is present from the start. Also, to set up a run, we autoclave the growth chamber and media separately (partially necessitated by the small size of autoclaves in Missouri). The growth chamber bottle is autoclaved lying on its side in a sling made from Tom Sanford's car seat belts. Another useful invention of Tom's is that the exiting gas from the fermentor is directed down a drain and up the sewer side of the trap. Such an exhaust tube goes a long way to making you more socially acceptable in your building. After 3 hours into the fermentor run, the O2 flow rate is turned up to 4-5 liters per minute and left at this rate for the remainder of the run. Around 4 hours, a slow trickle of cooling water is turned on to cool the hot little growing bugs. One advantage of the S.M.S. fermentor is the bacteria produced are in such a small volume that they can easily be collected in about 10 sterile 500 ml centrifuge bottles. Getting worms from the bacteria. We've found Nalgene transparent polycarbonate carboys to be excellent worm growth chambers. As advertised, they are lightweight and durable but somewhat more expensive than glass (~$40 from VWR). Due to our puny autoclave size, we've had to cut most of the necks off our carboys and create makeshift caps out of the lids of Sigma kilogram bottles. Two holes are cut in a lid as entrance and exit ports for 3/16 inch by 1/16 inch by 5/16 inch Tygon tubing to carry air into and out of a carboy. A third 3/4 inch diameter hole is cut as a sampling port and plugged with a #3 silicone rubber stopper. To keep the water level up, sterile water is also added through this port during worm growth (we don't think pre-humidifying air is good for the 'sterility' of worm cultures). The Tygon tube bringing air into the carboy is run to the bottom of the carboy where it is held weighted by stainless steel nuts loosely held onto the tubing by inserting a flanged plastic tubing adapter into the end of the tubing. The exit tubing begins just inside the makeshift lid and both inlet and exit tubing are glued to the lid by epoxy resin. Air coming in through the inlet tube is filtered through sterile cotton packed in a gas drying tube. Carboys are autoclaved filled with 16 liters of the modified S medium described by Ward (op cit). E. coli are added suspended in S medium to give a final concentration of 28 gm/liter of bugs and a final volume of 18 liters in the carboy. Before inoculating with worms, a carboy is vigorously bubbled for 24 hours. Aeration should be violent enough to form a standing wave 1 to 2 inches high. We have otherwise proven a few times too many that the worms will die of anoxia. The inoculum consists of 600 ml of a clearing culture of worms grown at a similar bacterial concentration in a 2800 ml Fernbach flask by rotary shaking. With such an inoculum, the carboy clears in about 4 days and becomes mostly dauers about a week after that, yielding 100 to 130 gm of dauers from ~500 gm of E. coli. Carboys are sat in an ice-water filled sink for 2 hours and then allowed to continue settling overnight at 4 C. After siphoning off the liquid, settled worms are cleaned by sedimentation through 14% w/w sucrose and flotation on 30% w/w sucrose in 250 ml centrifuge bottles spun at 2000 rpm in a Sorvall GSA head at 4 C. Sucrose is employed because on this scale, Ficoll 400 would be prohibitively expensive. For regular stages of worm, 35% w/w sucrose probably could be substituted in the flotation step with some loss of worms to the bottom of the bottle. A yield of over 200 gm of worms per carboy should be obtainable if the worms are not allowed to go to dauers. As for fermentor runs, directing the exhaust air from worm carboys down a drain helps make the air in the lab breathable. Mixing some air in with the pure oxygen from fermentor runs is probably also a good idea to avoid accumulating an oxygen atmosphere in your local branch of the sewer.