[
Worm Breeder's Gazette,
1992]
Quantitative traits such as life span and fertility are specified by the interaction of many genes (quantitative trait loci: QTLs). Mapping these QTLs involves assessment of genetic markers distributed throughout the genome as well as taking measurements of the quantitative traits of interest on the same individuals and/or strains. We have begun the process of QTL mapping for genes determining life span and self-fertility using the methodology described by Lander and Botstein (1989: In Genetics, 121:185-199) and .he primers for Tc1 polymorphisms provided by Ben Williams. We analyzed the segregation pattern of five Tc1 polymorphisms among 84 recombinant-inbred (RI) strains generated from crosses between Bristol (N2) and Bergerac Boulder (BergBO). We used the "single worm PCR" technique developed and described by Ben Williams (WBG, 11:4) to ascertain the presence or absence of Tc1 polymorphisms in each RI strain. N2 contains none of the Tc1 polymorphisms and BergBO carries 211 five of the Tc1 polymorphisms that distinguish each autosome. Since the RIs are homozygous, the presence of a Tc1 polymorphism specifies the chromosomal parent of origin as BergBO while the absence of the Tc1 polymorphism indicates N2 as the parent. A priori, we expected a random segregation of these markers. We observed random segregation for the polymorphisms found on LGIII and LGV but not for the polymorphisms found on LGI, LGII and LGIV [See Figure 1]. Apparently the N2 linkage groups I, II and IV are selected for. Next we examined the pattern of association of each polymorphism, again expecting a random assortment. This time we did observe random assortment [See Figure 2] RI strains are particularly useful for QTL mapping because of the various combinations of parental chromosomal regions which, through the use of genetic markers, can be assayed along with the phenotypic traits that are associated with particular regions. We have previously analyzed some of these RI strains on the phenotypic level in terms of their life span and self-fertility in an attempt to estimate the relative magnitude of genetic and environmental effects on life span and self-fertility, to estimate the number of genes involved in specifying these same traits, and to look at genetic and phenotypic correlations among life span and age-specific self-fertilities (Brooks and Johnson (1991). Heredity, 67:19-28). Now we are trying to assign chromosomal regions responsible for these quantitative traits. First we determine chromosomal parent of origin on the basis of presence or absence of each marker. Then, depending upon from which parent the linkage group came, we estimate the mean of the quantitative trait of interest. Using the formula provided by DeFries et al (1989 : In Alcoholism: Clin . Exp. Res . 3 : 196-200), we present our preliminary estimates of the amount of genetic variance in life span and self-fertility each polymorphic region on an individual linkage group specifies in Table 3. We want to emphasize that as we use more markers our estimates of the explained genetic variance per region will become more precise.
[
Worm Breeder's Gazette,
1999]
Fecundity assays are useful for categorizing and analyzing strains of C. elegans. Unfortunately they are laborious and time consuming. Conventionally, individual worms are placed on NGM plates and picked to new plates daily and their eggs/offspring are tallied (1). Picking, however, can be damaging to worms and is a potential source of variation. Our tests have shown that N2 worms picked at the L3 stage had significantly fewer offspring than those picked as eggs (T 0.05 [14] = 4.71 p < 0.001) or even those picked at the L2 stage (T 0.05[14] = 2.55 p = 0.023). To avoid this problem, individual worms were picked as eggs onto NGM plates and the eggs/offspring were picked daily and tallied. This eliminated the physical damage of picking, but it also greatly increased the time required to score plates as experienced "pickers" could spend up to an hour per plate. To speed the process, I have devised a novel, inexpensive piece of equipment called the "Twister Apparatus" which reduces the scoring time by 60%-75%. In addition to timesavings, even novice "pickers" are capable of impressive speed while "Twistering" and there is little adjustment time required. Twistering was chosen as the name for this method (think of a tornado carrying off the worms) when I got tired of saying that I was going to go "suck worms." It must also be noted that there is another way to avoid physical damage to worms that involves mouth pipetting individual worms from plate-to-plate using a drawn out melting point capillary (2). However, this is also time-consuming. At its most basic, the Twister apparatus is simply a worm vacuum. It is a length of flexible tubing connected to a suction source at one end and a nozzle at the other. Fifty microliter micro-pipets are ideal for Twister nozzles. They can be pulled out either by a pipette puller or by hand, although the latter takes considerable practice. The terminal end of the nozzle should be about 0.5 mm in diameter. The nozzle is then placed into a holder fashioned from two disposable pipete tips (Figure 1); yellow 200ml tips work the best (we use Fisherbrand). The tip of one is cut back so that the shaft of the nozzle will just fit inside. The wide end of the other is cut to allow it to fit inside the wide end of the other tip, held in place by friction. This assembly creates a stable holder for the fragile glass rod. This holder is then placed into the length of tubing connected to the suction source. Nozzles can be cleaned by submersion in NaOH to dissolve any worms clinging to the inside and then sterilized by submersion in ethanol prior to use and as needed during Twistering. Do not flame these nozzles, as the heat will melt the thin tips. Twistering plates has no significant effect on fecundity (T 0.05[10] = -1.36 p = 0.204) and does not damage the surface of the plate; only small spots are seen where the E. coli lawn is vacuumed up with the egg/worm and these grow closed overnight. (1) Brooks, A and Johnson, T. 1991. Heredity 67:19-28. (2) Wood, W. B. (ed.). 1988. The Nematode Caenorhabditis elegans. Cold Spring Harbor Laboratory.