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[
Worm Breeder's Gazette,
1991]
We have devised a very convenient, semi-automated injection setup closely based on the system of Mello and Ambros (wbg 11(4):7). It provides many of the advantages of the digital automated systems available, at a minute fraction of the cost. Our system is very similar to that described by Mello and Ambros, and anyone with a system like theirs can upgrade. The main difference is that the 3-way ball valve, used to control the duration of injection flow, has been replaced with an electronic solenoid-operated valve. This permits foot pedal (or any other way you'd prefer to operate an electric switch) control, and leaves both hands free to deal with micro- manipulation and focusing. In addition, the solenoid operated valve can open and close in a fraction of a second, permitting very precise control of flow. If the needle happens to be in the wrong place, a quick tap will show this without appreciably damaging the worm. at high pressure, quick on/off pulses also appear to aid in unclogging clogged needles, while at low pressure the system is ideal for pulse- filling oocyte nuclei or gonads. By varying the pulse time, it is possible to compensate for the flow characteristics of individual needles. By selecting the right match of components, it is possible to construct a system as drawn below which is extremely compact, attaching directly to the pressure source, requiring no cut hypodermic needles or tubing (except the piece connected to your needle) and occupying no bench space at all. Shown not to scale below, the entire system that attaches to a pressure regulator measures only 3' x 5' x 3/4', and minimizes dead gas space. Fittings such as those described by mello and ambros can be used; however, the key is to replace the 3- way ball valve (
b41x52) with a solenoid valve such as the nupro 110vac model. All metal-to-metal connections should be wrapped in teflon-based tape. Controlled pressure can be supplied to this system with a nitrogen tank via a high-pressure regulator or with a pump or in-house compressed air via a low-pressure regulator. For a complete parts list and any additional information, please contact us. In addition, a pre-assembled and tested complete system including foot pedal controller and the tubing that attaches directly to your needles can be obtained from tritech research (see above address and phone number). As an introductory offer to worm breeders, the system is available for $210, or $360 with high-quality pressure regulator and tank fitting (approximately the retail cost of the unassembled components). [See Figure 1]
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[
Worm Breeder's Gazette,
1997]
Two years ago, I compiled a small database of nematode strains of species both closely and distantly related to C. elegans. I distributed a report from this database (Worm Systematics Resource Net) at the Systematics Workshop at the 1995 C. elegans meeting in Wisconsin. This report listed strains (with relevant information) both by lab and within a taxonomic hierarchy. To make the database, lists of strains were generously provided by various labs willing to make these stocks available to the general worm community (and in some cases to the general scientific or educational community). This year, I would like to distribute an updated version at the 1997 meeting and put out an Internet version as well. The main purpose of the database is to provide a quick way of locating living stocks of particular nematode species for use in such work as systematics (phylogenetics and hybridization tests) and comparative biological studies (e.g., comparative development or molecular comparisons). But there are several other benefits offered by such a database (e.g., it provides a rough survey of rhabditid diversity that could be adapted for use in undergraduate biology courses). If you are willing to share your non-elegans strains, please send me a strain list with the following information for each strain: Species binomen (as well as is known-see footnote) Strain designation (please see footnote!) From whom obtained (if not isolated by someone in your lab) By whom originally isolated (if known) Isolation date (if known) Isolation locality (as accurate as possible) Isolation habitat (as complete as possible, including notes about ecology, temperature, altitude, etc.) Culture conditions (also note cryophilicity and any special protocols required)
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[
Worm Breeder's Gazette,
1982]
C. elegans var. Bergerac worms contain 1.7 kb of DNA adjacent to actin gene III that var. Bristol worms do not. This DNA polymorphism has been used as a phenotypic marker for mapping the cluster of actin genes I, II and III. We were interested to know more about this 1.7 kb of DNA. Other polymorphisms that were also 1.7 kb larger in Bergerac appeared after hybridization to randomly cloned fragments of Bristol DNA (Emmons et al., PNAS 76, 1333 (1979)). A DNA fragment containing the 1.7 kb insert was cloned from a lambda Charon-10 Bergerac recombinant DNA library. This clone hybridized strongly to about 50 bands in a whole genome blot of Bergerac DNA. The clone hybridized less strongly to Bristol DNA and hybridization was to a different set of about 30 bands. Subfragments of the cloned Bergerac DNA were used as hybridization probes against each other and revealed that the 1.7 kb insert is terminally repetitious. The end subfragments did not hybridize to the middle subfragment. The end and middle subfragments gave the same patterns when hybridized to whole genome blots of Bergerac and Bristol DNA's. The cloned Bergerac DNA was denatured, quick-cooled and examined in the EM and it revealed a single-strand 1.5 kb loop with a double-strand stem of about 75 bp. Therefore, the terminally repetitious ends contain inverted sequences. We hybridized the 1.7 kb DNA to blots of DNAs from worms from the Hirsh, Ward, Riddle labs and from the stock center. All had the same patterns. Two different patterns were found after hybridization to the wild strains PA-1 and C12A (from Carl Johnson) but no differences were seen with GA-l (from Carl Johnson's backyard). We don't know if this is a transposon but it seems like one. We also don't know if it's transcribed.
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[
Worm Breeder's Gazette,
1988]
As a part of study on neuro-muscular function, we tried to isolate mutants having abnormal response against some anesthetics. Sixteen strains of three independent mutations were isolated with EMS mutagenesis. Those had specific response and convulsive movement in 3OmM Ketamine-Hydrochloride(C13H16ClNO.HCl). All mutants had similar response to another drugs; serotonine, octopamine, as N2. In Ketamine, N2 worm had two phased kinetics between time and paralytic states, but responded no obvious convulsion to ketamine. Mutants basically show N2 like two phased kinetics, but were accompanied by clear convulsion during almost all stages. Modes of these convulsion was classified into two classes; quick and vibration-like convulsions in 15 strains. But nine of those show twitcher in the absence of ketamine and other strains had similar as N2. Another one shows wave- like convulsive movement in 30mM ketamine and had cold-sensitive uncoordinated movement in the absence of ketamine. This strain had almost paralytic phenotype at 16 C and recovered perfect motility after 40 min at 30 C. Genetic analysis shows that 16 strains divided into three independent mutations. Three of nine strains having twitcher in the absence of ketamine were mapped on LG-IV and could not complement to
unc-22(
e66). Strain J030 having wave-like convulsion was mapped on LGV. Double mutant from trans configuration to
dpy-11(
e224) was not obtained. This means that mutation site closed to
dpy-11(
e224), or might be the same cluster. Other strains showing vibration-like convulsions in 30mM ketamine but normal behavior without ketamine was in progress. One of them might be on LGII. These results indicate that ketamine had multiple function to neuro-muscular mechanisms in the worm. Further investigation of defectivity of these mutants and molecular characterization of defective genes allows us to know new aspects about mechanisms of receptor-effecter circuit and cold- sensitive uncoordinated movement of C. elegans.
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[
Worm Breeder's Gazette,
1993]
Acedb has been written in collaboration with Richard Durbin to provide better access and a denser meshing of all the information about C.elegans. You are all concerned, because the authors of the data are the genuine authors of the database. Up to now, we only had time to parse the data readily available from the CGC, the EMBL and the Sulston-Waterston mapping and sequencing project. Acedb will of course continue to include these sources and you should keep sending there the relevant data. But much more is known about our dear nematode. It would clearly be too much work for one person to extract the information from the available literature. However, we believe that if each of you was willing to share this burden and contribute short descriptions of what he knows, everybody would greatly benefit from these efforts. We want to include phenotypes, descriptions, cloning, transformations, sequences, expression patterns and also your toolboxes like balancers, expression vectors or protocols. The contributions will be split in short notes, signed by their authors. They will extend the present acedb descriptions and be distributed in the following acedb release. Hopefully every other month or so. To achieve this result in a quick and simple way, we will send some personal questionnaires about those genes or clones we think you may have touched, along the lines of the existing acedb the data into a precanned format. This will at the same time be more accurate and simplify the clerical work. So please feel free to send your own suggestions. We will check technically the format of the data and may suggest some style changes or spelling rationalisations, for the sake of uniformity and cross-referencing, but the general philosophy is that it is more informative and more accurate to let people contribute directly their personal data and sign them. We hope to prototype these ideas before the next worm meeting and think that they should prove extremely rewarding to all of us.
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[
Worm Breeder's Gazette,
1997]
In the process of genetic mapping we learned that the left arm of LG IV contains restriction fragment length polymorphisms between the strains N2 and DP13 (a Bergerac RW7000 subclone). These were detectable by Southern hybridization with cosmid probes (G. Beitel, personal communication). This region happens to be the target of current sequencing efforts. Visual inspection of cosmid sequences revealed the frequent occurrence of repetitive DNA, leading us to wonder whether the RFLPs arise from differences in lengths of these repetitive sequences in the two strains. The repetitive sequences are similar to satellite repeats in that they consist of tandem repeats of short units flanked by nonrepetitive sequence. We analyzed 12 such repetitive regions, with repeat unit lengths varying between 6 and 30 bp, and between 300 and 1800 bp in total length. PCR was performed on single N2 and DP13 worms as described by Williams (Epstein & Shakes, Methods in Cell Biology volume 48) using primers based in nonrepetitive flanking DNA. Analysis of PCR products by 1% agarose gel electrophoresis showed length differences between N2 and DP13 for 7 of the 12 regions. In each case the length of the fragment amplified from N2 was consistent with sequence data. DP13 PCR products were longer or shorter than the corresponding N2 PCR products by a few hundred base pairs. It seems likely that the length differences are due to differences in the number of unit repetitions. We plan to test other strains for these polymorphisms. PCR analysis of polymorphisms is a simple and quick way to determine the genotype of a single worm at a physically defined location. This method can be useful for gene mapping through analysis of F2 progeny from a cross between a worm homozygous for the gene of interest and a wild-type worm of a different strain (Korswagen et al., PNAS 93, 14680-14685, 1996). The distance between a repeat region and the gene of interest should be inversely proportional to the frequency of coincidence of the mutant phenotype and homozygosity for the repeat length of the strain of the mutant parent.
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[
Worm Breeder's Gazette,
1988]
We have been characterizing the C. elegans small nuclear RNAs ( snRNAs) and their genes (see Thomas, Zucker-Aprison and Blumenthal, this issue). snRNAs are the RNA component of small nuclear ribonucleoproteins (snRNPs) which catalyze various nuclear RNA processing events. The snRNPs which we have been studying, U1, U2, U4, U5 and U6, have been shown to catalyze cis-splicing of mRNA introns in human cells and in yeast. As probes we have used DNA oligomers designed to hybridize to highly conserved sequences in the snRNAs. In other organisms the snRNAs U1, U2 U4, and U5 have been shown to be bound to the Sm protein and have a special 2,2,7-methyl-guanosine ( trimethyl-guanosine, TMG) cap. The U6 snRNA lacks these features, however it is generally present in anti-Sm or anti-TMG cap immunoprecipitates via its ability to associate with the U4 snRNA by intermolecular base-pairing. We immunoprecipitated C. elegans nuclear extracts with anti-Sm and anti-TMG cap antibodies, extracted the RNAs, separated them on denaturing polyacrylamide gels, and blotted them onto nylon. When these blots were probed with [32P]- labeled oligonucleotides, we found U1, U2, U4, U5 and U6 snRNAs present in the immunoprecipitates, as expected. We also immunoprecipitated snRNAs from deproteinized total C. elegans RNA with anti-TMG cap antibody and analyzed them in the same way. We found U1, U2, U4, U5 and a small amount of U6 snRNAs in these immunoprecipitates. However, if we heated and quick-cooled the total RNA before the immunoprecipitation, there was no U6 snRNA, consistent with the idea that U6 and U4 interact by base pairing. Surprisingly, when we probed these same blots with a labeled DNA oligomer complementary to the trans-spliced leader RNA, we found the leader RNA present in all of the immunoprecipitates. The leader RNA's presence in the anti-Sm immunoprecipitates suggests strongly that it interacts with the Sm protein. The existence of a consensus Sm- binding site in the leader RNA's 'intron' portion suggests that this interaction is direct. The immunoprecipitability of the leader RNA with anti-TMG cap antibody was insensitive to heating and quick- cooling, hence it must interact directly with this antibody. To prove that this interaction was not due to a cross-specificity of the antibody, we showed that the compound 7-methyl GpppG (the standard mRNA cap) competes with the leader RNA for binding to the antibody as inefficiently as it competes with U1, U2, U4 and U5 snRNAs. We conclude that the leader RNA has properties of an snRNA. If the leader RNA snRNP is indeed a substrate for transsplicing, then one would predict that a 'Y'-structure product (analogous to the lariat of cis-splicing) would be Sm-bound and that the recipient mRNAs would have 2,2,7-methylguanosine caps at their 5'-ends. We have tested the latter prediction by analyzing the supernatant and pellet from an anti-TMG cap immunoprecipitation of total C. elegans RNA on blots of formaldehyde-agarose gels. While the U2 snRNA and the leader RNA were present in the pellet, actin mRNAs were present only in the supernatant. Furthermore, we saw diffuse hybridization to an oligonucleotide complementary to the 'exon' portion of the leader RNA only in the supernatant, which we interpret as representing the sum of the trans-spliced RNAs. These data indicate that the mature RNAs that have been trans-spliced lack the TMG cap structure of the leader RNA. This result suggests the existence of a cap modifying or replacing activity whose substrate is mRNAs that have been trans-spliced. Our findings have important implications for the mechanism of trans- splicing in C. elegans. The leader RNA, discovered as substrate of trans-splicing, shares properties with snRNPs, which were heretofore known as catalysts of RNA splicing. Since the splice sites used in trans-splicing fit the consensus for splice sites used in cis-splicing, it seems likely that the two types of splicing are mechanistically related. That the leader RNA has properties of an snRNP suggests it may have special interactions with other snRNPs that catalyze its joining to recipient RNAs in C. elegans.
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[
Worm Breeder's Gazette,
1988]
Borrowing yet again from yeast molecular biology, the following procedure has been used to isolate clean, intact RNA from small quantities of worms. This saves the time and effort of growing up, and grinding in liquid nitrogen, large liquid cultures. This procedure should make it possible to process easily 10 to 20 stocks in parallel for screening by Northerns. Yields are ~200 g of clean RNA from 4x9 cm 'high yield' plates (see below), or about 2 g of clean RNA from a single, 5 cm streaked plate. Mini-prep. Wash worms off 4 plates (5 cm) that are just clearing. If you want, the worms can be incubated in S-media for 30 min to digest whatever they have in their gut, and then purified by standard sucrose flotation in a 5ml tube. Resuspend the worm pellet in 0.5 ml of liquid. (Use any RNAase-free, solvent resistant, sterile tube that is big enough. I use Falcon 2098 50ml disposable polypropylene tubes.) Add 2ml GuEST buffer and 2ml PCI (see below), and then 6g glass beads. Vortex on high speed, using a regular bench vortex, for 2 min at room temp. Draw the liquid off the beads with a pipetman into 4x1.5 ml eppendorf tubes, rinse the beads with 0.5ml of PCI, and add this to the microfuge tubes. Microfuge for 2 min. Transfer the aqueous layer to fresh tubes, add 1/10 vol 3M NaAcetate, pH 6.0, and re-extract with PCI. Spin, transfer the aqueous phase (leaving the interface, if any, behind), and precipitate the RNA by the addition of 2 vol of 100% ethanol. -20 C for 20 min, spin in microfuge 10 min, decant, and wash pellet with 80% ethanol. Dry, and dissolve the pellet in 0.3ml of dddH20 (see below). Add 0.9ml of 4M NaAcetate (diethylpyrocarbonate ( depc) treated), and let sit at least 5 hr at 4 C. Spin in microfuge 10 min, discard supernatant. Dissolve the pellet in 0.1ml dddH2O, add 5 l 3M NaAcetate, and 210 l 100% ethanol. Precipitate at -20 C for 20 min, spin, wash with 80%, and dissolve the final pellet in 50 l dddH2O. Dilute 2 l to 1ml with dddH20, read A260 and multiply the absorbance reading by 20 to get the RNA concentration in mg/ml. p(A)+ selection, using poly(U) Sepharose (Jacobson [1987] Meth. Enz. 152:254261), should give 2-4 g of p(A)+ RNA, although using total RNA has given me good signals on a Northern with an actin probe. If you are in a hurry, you can skip the NaAcetate precipitation step, since large MW DNA won't transfer upon Northern Blotting. Micro-prep. Wash the worms off a single 5 cm plate that is just clearing. Repeat the above procedure, using 50 l worms, 100 l GuEST buffer, 100 l PCI and 0.5g glass beads. Rinse beads with 50 l PCI. Do the salt precipitation of RNA in 30 l by adding 90 l of 4M Na Acetate, and dissolve the final pellet in 10 l. Frozen Worms. I have used this procedure, successfully, to prepare poly(A)+ RNA from frozen worms. Washed worms were suspended in 0.1M NaCl, quick-frozen in liquid nitrogen, and kept at -70 C for 1 month. Use 4ml of GuEST, 4ml of PCI and 12g of beads per ml of worms. Thaw the tube just until the frozen plug of worms can be removed, and drop the plug into the GuEST/PCI. Proceed as with the mini-prep, although you may want to vortex a bit longer. Buffers. GuEST (From M. Goedert) Add 245ml sterile distilled water to 200g Guanidine isothiocyanate (BRL Ultra Pure). Add 21ml 1M Tris pH 7.4, and 42ml 100mM EDTA. Heat gently to dissolve. Add 9ml Sarkosyl, and 4.2ml -mercaptoethanol. Bring volume to 420ml with sterile distilled water. Filter through a sterile Nalgene filter, and store at 4 C. PCI. Phenol:Chloroform:Isoamyl alcohol, 25:24:1 Glass Beads. 0.3 to 0.4 mm diam (although I haven't tried other sizes). I borrowed mine from J. Kilmartin, but apparently BDH and Sigma sell them. They can be acid washed, baked and re-used. dddH20. Add depc (0.07% v/v) to sterile, double-distilled water, shake for 10 min, and then autoclave. 'High Yield' plates. (From A. Spence) Add a drop of 20% glucose to a large NGM plate, and then spread the plate with a wild-type bacteria such as NA22. Let sit overnight before adding worms. HINTS: Wear gloves, use only sterile tubes and pipette tips, and generally treat the RNA as you would HIV, and you should have no problems. A quick method for checking the quality of your RNA is on a 0.8% agarose minigel. Denature the RNA for 10 min at 65 C, chill on ice and add sterile glycerol/dye/buffer. Run 2 g RNA/lane in a RNAase free gel box with standard gel buffer, ~7 V/cm for 40 min. EtBr stain 5', photograph. You should see two tight, rRNA bands, with no high MW DNA visible. Avoid using old plates, where the worms have started burrowing, as the softened agar tends to wash off with the worms, and contaminate the RNA. In these cases a small-scale sucrose flotation might be useful to clean up the worms, but I haven't yet tried this.
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[
Worm Breeder's Gazette,
1983]
We have developed a method to stain C. elegans embryos for an esterase activity that appears early in embryogenesis and is restricted to the E cell lineage. This provides an additional biochemical marker for early lineage studies. Method: Nuclepore filter circles (~1cm diameter polycarbonate filters, 8 m pore size, Nuclepore Corporation, 7035 Commerce Circle, Pleasanton, California 94565) are rimmed with rubber cement and placed on a siliconized depression slide. Small numbers of embryos ( ~100-200) are pipetted onto the filter and processed in volumes of 100 l. This allows quick exchange of solutions, by drawing off liquid from beneath the filter with a mouth-pipette under the dissecting scope, and minimizes loss of embryos. Pretreatment: Six to eight minutes 1% Triton-X100 in M9 (100 l volume throughout) three washes, 0.1M KH2PO4/Na2HPO4 buffer, pH 7.0, Hypochlorite for 2.5 minutes (1:2 dilution of stock 4-6% NaOCl solution, in phosphate buffer, degassed), three washes in buffer. Chitinase (Serratia marcescens chitinase, US Biochemical Corp., at 20 mg/ml in egg buffer. (Laufer, et. al., Cell 19: 569)) two to six minutes, observing under dissecting scope, until eggs round up and/or pretzels begin to unfold. Fix three minutes at 4 C in 4% formalin in phosphate buffer. Three washes, buffer (total about three minutes). Stain two to six hours at 4 C with freshly made stain (see below). Mount on slides in phosphate buffer, transferring either filter (peel off rubber cement) or individual embryos (better optics, but harder). Seal with nail polish. Staining improves overnight in the refrigerator. Stain recipe: (Lojda, Gossrau and Schiebler, Enzyme Histochemistry, Springer-Verlag, 1979) Mix: 20 l 4% NaNO2 in H2O (make fresh weekly) , 20 l pararosaniline solution*. Add: 750 l 2.8% Na2HPO4 (in H2O), 20 l 0.2M NaOH, 20 l alpha-naphthyl acetate (1% in acetone) *( Dissolve 400 mg pararosaniline HCl in 8 ml H2O, add 2 ml concentrated HCl and stir 15-30 minutes at room temperature. Centrifuge and millipore. Store refrigerated up to two months. Esterase activity (indicated by dark red staining) appears consistently in four to eight contiguous cells of embryos fixed 160 to 180 minutes after first cleavage (20 C) (i.e., with approximately 110 to 180 total cells). It is clearly localized to the E cells at the lima bean and later stages. Eight to 12 cells of partial embryos (50 to 100 total cells) derived from isolated P blastomeres stained for esterase activity, whereas no staining was found in partial embryos derived from isolated AB blastomeres. Thus, expression appears autonomous within the P lineage. We are currently investigating expression following cytochalasin and alpha-amanitin blocking of embryos permeabilized by cracking (Laufer, et. al.). Long-range goals are to clone the gene for this esterase and investigate the regulation of expression of this and additional gut-specific hydrolases.
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[
Worm Breeder's Gazette,
1996]
The reverse genetic analysis of C. elegans has become important as result of the fast amount of sequence data that come out of the genome project. Techniques have been developed to analyse the functions of newly identified genes (e.g. transposon induced gene inactivation). Although forward genetic screens are also facilitated by the genome project, there is no quick route yet from mutant phenotype to gene identification. We have developed a method that greatly reduces the time required to identify transposon insertion mutants from forward genetic screens. The method is based on the disruption of a gene by a known transposon sequence. It starts with a mutator strain of low Tc1 and Tc3 copy number from which transposon insertion mutants are isolated with a specific phenotype. Genomic DNA is digested with a frequently cutting enzyme, and an oligonucleotide-vectorette is ligated to the digested DNA (Riley, J., et al. (1990) Nucleic Acids Research 18:2887-2890). A PCR is performed using a primer corresponding to the transposon end and a primer for the vectorette. This will result in the amplification of only those restriction fragments that contain a transposon end. A second PCR is performed with nested primers, of which one is radiolabeled. The products from this reaction are separated on a denaturing polyacrylamide gel. The autoradiogram shows in one lane the transposon insertions present in the genome of one strain. The novel band (absent in the non-mutant starting strain) is excised from the dried polyacrylamide gel, further amplified, and sequenced. The sequence of the flanking DNA can be compared to the database and the gene that is responsible for the phenotype can be identified. We have used this transposon insertion display method to identify a gene that is involved in the perception of copper. Wild type animals have an aversion for copper; they will not cross a line of 0.25 M CuSO4 even if there is an attractant on the other side (isoamylacohol). We have used the mutator strain NL917 (
mut-7) and isolated a mutant (NL953) that did cross the copper. The mutant was out- crossed with Bristol N2 and a panel of mutant and wild type animals was obtained. Using the display we identified a Tc1 insertion that was present in all mutant animals and absent in wild type animals. The flanking sequence of this insertion was determined and compared to the database. A match was found in the dataset of unfinished sequences released on September 8th 1996. We are further optimizing the protocol and we are analysing other copper aversion mutants as well as other classes of mutants. The current version of the protocol is available on request.