Johnson TE et al. (1986) Worm Breeder's Gazette "developmental effects of loci lengthening life-span."

Bryant M, Johnson TE, Conley WL

[Worm Breeder's Gazette, 1986]

Both recombinant inbred (RI) lines and single gene mutants (see other abstracts in this issue) have been produced which have dramatic changes in life-span. Stocks average a 60% increase in mean and maximal life-span. We have been interested in knowing whether these changes also result in a change in developmental events (length of the embryonic period, timing of cell divisions, rate of larval growth, timing of larval molts) or in the length of the reproductive period. We have assayed the length of the embryonic period by isolating individual eggs at the 2-cell stage and transferring these eggs to incubation chambers at 16 C (chosen for convenience of monitoring). About 20 hours later the eggs hatch. We find no change in the length of this period in either the RI lines or in age-1(hx542) or age-1( hx546). Surprisingly, N2 seems to take about 1 hour longer to complete this phase than any other stock analyzed, including Bergerac BO. We see variation among the RI lines in the rate of larval growth, in the timing of larval molts, and in the time from hatch to 50% adult fecundity. These variations do not seem to correlate with the adult life-span but limited numbers of stocks have been assayed as of yet and we remain unclear about our level of statistical significance. It is clear that both(?) age-1 alleles are without effect on any aspect of larval development rate. There is marginal variation in the length of the reproductive period and in the total fecundity, among the RI lines. The quantitative variations in total fecundity showed statistically significant correlations with the length of life in one repeat which has not yet been confirmed. There is no correlation between the length of the fecund period and life-extension. For the age-1 locus, there is a significant reduction in total brood size but no significant change in the temporal distribution of progeny production, after normalizing to total brood size. As always, we welcome helpful and provocative suggestions of additional questions to ask and directions to proceed.

Albertson DG et al. (1982) Worm Breeder's Gazette "antibody staining of C elegans."

Hedgecock EM, Sulston JE, Albertson DG

[Worm Breeder's Gazette, 1982]

We have used two procedures for staining C. elegans with florescent antibodies. In the first procedure, which is especially useful for eggs: 1) Young gravid adults (about six) are picked and placed into a 5-10 l drop of M9 buffer on a microscope slide subbed with bovine serum albumen. (Slides dipped in a 0.1% BSA solution, drained and dried in a vertical jar beforehand.) The animals are rinsed, if necessary, with M9 buffer then cut at the vulva with a razor blade. 2) A 12 x 12 mm coverslip is placed gently on top of the drop so that the eggs are not crushed. The slide is immediately placed on dry ice for at least 10 min. 3) The coverslips are pried off with a razor blade and the slide placed in methanol at -20 C for 2 minutes followed by acetone at - 20 C for 4 minutes. Slides are then rehydrated through a series of alcohols (90, 70, 50, 30% ethanol; 2 minutes in each) and then into two changes of PBS (phosphate buffered saline). They may be stored in the final jar of PBS. 4) Just prior to use, the excess PBS is wiped off the slide leaving a drop over the specimen and 5-10 l of antibody are pipetted onto that spot. The slide is placed in a humidified chamber for 45-60 minutes, rinsed briefly by gentle dipping into a jar of PBS, and then for five minutes in a second jar of PBS. A second antibody may be applied to the damp slide. It is then incubated and rinsed as before. This method gives reproducible staining of eggs using antitubulin and useful, though less consistent, penetration of the cut adults. Penetration of antibodies into whole mounts of eggs appears to be possible because the freezing and removal of the coverslips crack the eggshell and sometimes remove it from the egg. The thickness of the liquid layer between the coverslip and the slide is probably most crucial and the correct amount will probably be found by practice. To overcome penetration problems in larger animals, we have used a squashing procedure, followed by freezing and fracturing which yields flattened half-animals attached to each slide with their internal surfaces facing outward. This procedure can be modified for antigens which do not survive the organic solvents used in step 3 above. Washed, densely settled, worms are pipetted onto BSA subbed slides in 1 l drops of M9 buffer (up to 2 x 4 = 8 spots per slide). A second subbed slide is lowered over and the two slides are pressed together with the thumbs without displacing the top slide laterally. The pair of squashed slides is frozen on dry ice and then pried apart with a razor blade. At this stage, the slides can be fixed in one of three ways, according to the antigen: 1) sequential immersion in cold methanol and acetone as described above (slides are air dried and stored after acetone) 2) heating for 2 minutes on a hot plate at 100 C or 3) air drying at room temperature. The last procedure is the mildest but the fractured wormhalves are poorly fixed and may wash away during staining. For humidified chambers, we use either a large petri dish containing water and the lid of a small petri plate to elevate the slide or, more simply, an inverted agar plate (containing 0.1% sodium azide to maintain sterility).

van Swinderen B et al. (1997) Worm Breeder's Gazette "Halothane, A General Anesthetic, Confers Resistance To Aldicarb."

Roberson R, Crowder CM, van Swinderen B

[Worm Breeder's Gazette, 1997]

Halothane is a volatile general anesthetic that disrupts coordinated movement in C. elegans at low, clinically-relevant concentrations (1) (0.2-0.5 volume % halothane), and abolishes gross movment at higher concentrations (3.5 vol. %). The nematicide drug aldicarb, an inhibitor of cholinesterase, paralyzes worms (>0.5 mM for N2) due to an excess of acetylcholine accumulating at the synapse. Mutations in pre-synaptic machinery can either confer resistance or hypersensitivity to aldicarb (a ric vs hic phenotype). Previous work has implicated several pre-synaptic components in halothane-induced anesthesia. We hypothesized that should halothane disrupt pre-synaptic machinery, then the reduction in acetylcholine release would confer resistance to aldicarb. Indeed, halothane makes worms ric. Halothane-induced resistance to aldicarb was measured by a simple movement assay. Worms (N = 20-30) were exposed to a seeded aldicarb plate for 4 hours, and were then picked into a 0.5 cm circle outlined on the same plate. An hour later, the percentage of animals having managed to crawl completely out of the circle was scored. Each assay was done in triplicate at various halothane concentrations within air-tight chambers,including a no-halothane control. Table 1 summarizes results for N2 and some mutant strains. Low, clinically-relevant, concentrations of halothane (0.2-0.5 vol. %) significantly reversed aldicarb-induced paralysis. Interestingly, higher concentrations of halothane (1-1.5 vol. %) do not show a significant effect in N2; the reversal effect peaks at 0.2-0.3 vol. % halothane and then gradually decreases to near control levels. Similar experiments performed with the acetylcholine agonist levamisole and halothane failed to show any reversal, suggesting that halothan acts pre-synaptically (data not shown). We tested several mutant lines for disruption of halothane-induced resistance to aldicarb, reasoning that by blocking this effect one might highlight clinically relevant molecular pathways targeted by volatile anesthetics. Aldicarb concentrations were optimized according to the ricness or hicness of each strain. Shown in Table 1 are results for some G protein and calcium channel mutant strains. egl-30 (n686) (a.k.a.Gqalpha) does not reverse at all, and neither do calcium channel mutations egl-19 (ad1013)lof and (n2368)gof. On the other hand, goa-1 (sy192) reverses significantly at low halothane concentrations, but continues to show robust reversals at higher concentrations as well. The same striking response was also seen for egl-10 (nIs51), which overexpresses the upstream negative regulator of goa-1 signalling. In conclusion, the anesthetic halothane can result in an organism "waking up." By understanding aldicarb's mechanism of action, some light is shed on the way volatile anesthetics work. The analysis of numerous mutants suggests that halothane has a pre-synaptic effect at clinically relevant concentrations, and that G proteins and calcium channels are involved . Table 1. (Crawling Indeces) ** significant movement compared to control Halothane Concentration Range (vol. %) Strains 0 (control) 0.2 - 0.5 1.0 - 1.5 [aldicarb] mM N2 0.16+/-0.04 0.52+/-0.04* 0.27+/-0.04 0.5 egl-30(n686) 0.14+/-0.02 0.15+/-0.03 0.04+/-0.01 0.8 egl-19(n2368) 0.15+/-0.03 0.20+/-0.03 0.05+/-0.02 0.5 egl-13(ad1013)0.20+/-0.03 0.23+/-0.03 0.11+/-0.02 0.5 goa-1(sy192) 0.10+/-0.02 0.62+/-0.06* 0.65+/-0.02* 0.1 egl-10(nIS51) 0.01+/-0.01 0.49+/-0.07* 0.63+/-0.03* 0.2 1. Crowder, C.M., Shebester, L.D. & Schedl, T (1996) Anesthesiology 85, 901-912.

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.