Overview
Our protocol for generating deletion mutations is based on the procedures of Bob Barstead and Gary Moulder at the Oklahome Medical Research Foundation in Oklahoma City. The procedure is to mutagenize a large population of worms with trimethylpsoralen and UV irradiation, set up 1152 subpopulations, screen DNA made from this library for deletions in specific genes by nested PCR, and then to recover single worms carrying the deletions through a sib-selection process. We have made various modifications to the original protocol, specifically with regard to library complexity (100,000 genomes versus 500,000 to 1,000,000) and screening strategy (the use of poison primers to increase sensitivity of detection for small deletions).
This procedure is not terribly complicated, but it does require great care at every step to ensure that detected mutations can be recovered. Our libraries are not frozen, as we have more genes on our request list than we can search for in a single library?s DNA, and our throughput is such that we can screen until the DNA is used up in about three weeks. DNA is made by a simple proteinase K lysis, and pooled so that the 1152 populations can be screened in 96 reactions. We do a lot of our PCR work in 384-well plates to save time and materials, but everything can be done in 96-well plates if you don?t have access to 384-well equipment. 10 ul reactions work fine in 96-well plates, and don?t require nearly as much Taq. To date we have done most steps with worms grown on solid media, but we have shifted some steps to growth in liquid and continue to explore making the whole library in liquid culture.
The following sections lay out the protocol in fair detail, and other sections of this site contain our PCR protocols, recipes and materials lists. If you have questions about anything that we have not covered, please be sure to contact us.
Library Construction
We use our own subculture of the wild type (N2) to make mutagenized libraries. To minimize accumulation of spontaneous mutations in the stock, it is re-established from a single animal and grown to a density suitable for library inoculation before each library is built.
Worm growth and synchronization for library construction follows a schedule intended to make major events fall on regular work days. The timing of some steps is important, but that of certain others (indicated in the text) may be adjusted to fit your schedule without detrimental effects.
Initial setup
Day 1 (Wednesday) Wash two or three 60 mm plates of growing mixed-stage worms with sterile M9 buffer, pellet by low-speed centrifugation in a sterile 15 ml conical centrifuge tube (2000 RPM/1 minute), and distribute with a sterile Pasteur pipette to three 150 mm RNGM plates spread with a lawn of E. coli OP50 (three spots of worm suspension per plate).
Synchronize
Day 3 (Friday) Synchronize population by egg prep, following the procedure in Barstead, R.J. 1999. Reverse Genetics in ?C. elegans. A Practical Approach.? I.A. Hope, editor, Oxford University Press, NY. 1: 97-118. Distribute harvested eggs (or killed and washed gravid adults) to one or two unseeded 150mm plates. The eggs will hatch, and the L1 larvae arrest development until put onto food.
Transfer to food
Day 5 (Sunday) Wash L1 larvae off plates with M9 buffer, pellet in a 15 ml centrifuge tube, remove supernatant by centrifugation and distribute with a sterile Pasteur pipette to 10 seeded 150 mm plates. Place at 15oC and allow to develop to young adulthood (two days).
Mutagenesis
Day 7 (Tuesday) Wash worms off all plates with M9 buffer, pellet in 15 ml centrifuge tubes, and remove supernatant by aspiration. Wash with fresh buffer several times to remove most bacteria. Resuspend worms in one or two 1-ml aliquots of M9 buffer and transfer each aliquot to a single well of a 12-well untreated tissue culture plate (Evergreen #222-8046-01F).
Add 6.7 ul TMP working stock (0.3 mg TMP/ml acetone) to each 1 ml suspension of worms (final concentration 2 ug/ml). It?s best to do this in a fume hood, probably under dim light. It?s a little hard to see, but if you get the right angle you can see when the TMP goes in because the acetone makes the surface of the worm suspension sheet from one side to the other in a peculiar way. Incubate the worms in the dark for one hour at room temperature, with occasional gentle shaking. We do this in the fume hood, with the plates in a lightproof box.
While the worms are incubating in the TMP, set up the irradiation apparatus. We irradiate with a dissecting microscope fitted with a Kramer fluorescence illumination unit and a UV filter set (360/40 nm). Turn the lamp on at least 15 minutes prior to irradiation to give it time to warm up while you are adjusting the light intensity and field of illumination. Remove the objective and adjust the height, zoom and focus so that the field of illumination is slightly larger than the well the worms are in (use an empty 12-well plate, with the filter set turned to one of the GFP settings so you can see the circle of light). Elevate the plate to the same height as the window of the UV dosimeter on a stable support. We use a styrofoam tube rack for this - it helps to draw an outline of the tissue culture plate on the rack and a circle exactly where the well with worms will be, so you can put the worm well directly in the right place at irradiation time without a lot of fussing. Once the size of the field is set, replace the support and plate with the UV dosimeter, center the circle of illumination on the window with the filter set still on the GFP setting, then change the filter to the UV setting and adjust the height of the head until the intensity is 340 microwatts per square centimeter. The adjustments required are usually small and won?t significantly change the size of the illuminated circle. It?s good to check the UV intensity a few times while the worms are incubating, as the lamp intensity tends to drift over time. Once you?re happy with everything, change the filter set back to GFP, replace the dosimeter with the support and plate and realign it for even illumination of the correct well. Carefully remove the empty plate without disturbing the support, put the lamp blocking slide in place, and turn the filter set back to UV.
After the worms have soaked in TMP for one hour, remove the plate lid, carefully place the plate with worms in position on the support, and irradiate for 90 seconds at 340 microwatts per square centimeter (use the lamp blocking slide to control irradiation). You can leave the room light on for this.
After irradiation, transfer each worm/TMP mix to a microfuge tube and pellet. A 10-second spin at 10K rpm is usually sufficient. Remove the pellet from under the supernatant with a Pipetman (200 ul), and distribute at three dots of worms per plate to each of five or six seeded 150 mm plates. We use one tube of pelleted worms for each set of five or six plates. (You?ll discard some worms, but five to 10 150mm plates is LOTS of mutagenized worms. Don?t sweat it.) Put the plates in the dark at 20oC to 22o overnight to recover.
Synchronize and set up to screen for mutations in unc-22
Day 8 (Wednesday) unc-22 is a large gene that makes a good test target for mutagenesis. Mutations that knock out unc-22 produce a dominant "twitching" phenotype when the animals are exposed to 1% nicotine. The presence of twitchers among the progeny of a mutagenized population is a good assay to indicate that the mutagensis was successful. After 24 to 36 hours, set up 25 60mm plates with three mutagenized adults each. Typically we screen for twitchers before harvesting for the DNA prep. You should see twitchers on at least half the plates. If we see twitchers on fewer than seven or eight plates, we throw the library away and start again.
Immediately after the P0s are picked for the twitcher screen, synchronize the remaining P0s by egg prep and put the eggs on a fresh set of 150mm plates.
Set up library
Day 10 (Friday) The F1s should starve the plates just as the first F2 eggs are being laid. Collect the worms by standard washing and pelleting, and dilute with fresh M9 buffer to desired concentration in 50 ml centrifuge tubes. Set up library by distributing 50 worms per well to 1152 wells in 12-well untreated tissue culture plates containing low-EEO agarose and lawns of OP50 (96 12-well plates, numbered sequentially). We use Nichiryo Model 8100 repeat pipetters fitted with 3.0 ml syringes for this, delivering 50 ul of worm suspension per well (1 worm/ul). It?s not a terribly accurate method, but it works well enough for library construction and it?s fast: one person can dispense the library in a couple of hours.
Harvest library
Days 15 to 17 (Wednesday to Friday) Accuracy at this stage is essential! Each worm well must be sampled into the correct digestion well or addressing will become impossible and you will waste all your time, effort and materials! The actual time of harvesting depends on the worm plates ? harvest when the wells have just starved. Prepare 12 96-well 200 ul PCR plates, labeled A through L with rows numbered 1-8 and containing 50 ul Proteinase K buffer in each well. Harvest each worm well in order by putting on 200 ul of harvest solution (sterile deionized water containing Nystatin and streptomycin), removing 50 ul and placing into the Proteinase K buffer in a digestion plate well. There are several concepts and tricks that will help you do this properly.
The concepts:
Remember that each 12-well plate corresponds to one row of wells in the digestion plate. Thus, worm plates 1 through 8 occupy rows 1 through 8 of digestion plate A, and so on.
To avoid cross-contamination, each well must be sampled with its own pipette tip (one box of tips per digestion plate).
The tricks:
Add the harvest solution using a repeat pipetter, 12 wells (one plate) at a time.
To get good representation in each population sample, swirl the plate gently after adding the harvest solution and tip the plate up on one edge so the liquid sheets down the well and collects in the corner between the agarose surface and the wall. Sample using a 200 ul Pipetteman, and mix well before taking the actual sample by drawing up and expelling 50 ul several times. It?s best to place the tip against the wall just above the agarose to avoid clogging the tip with stray bits of agarose.
The cleanest way to keep samples straight is to eject the used tip into the well with the worms, so that only wells without tips remain to have worms added. This requires some care and a deft touch, as it?s easy to eject with such force that the tip jumps back out of the well. Also, removing the tips at the end with a 12- or 8-channel pipetter requires care so that you don?t puncture the well bottoms with the tips. If you do it this way, be sure to blow out all the liquid that wicks up into the tip while it?s in the well.
Another way to keep samples straight is to maintain strict one-to-one correspondence between tip positions in the box and well positions in the digestion plate and make a visual check for each sample. This requires much greater concentration and a bit of counting to yourself, but eliminates the possibility of bouncing tips out or puncturing wells.
Once you?ve harvested a digestion plate full (eight 12-well plates), put on a silicone mat lid and place at ?70oC for about a half hour or until all the wells have frozen completely. You may leave them at ?70oC for any length of time without causing any trouble. Transfer the plate to a 96-well PCR machine, incubate at 65oC for two hours, then run the temperature up to 95o for 20 minutes to inactivate the Proteinase K. Store digested plates at -20oC until all plates are finished.
Pool DNA for screening Day 20 Thaw all digestion plates and add 100 ul sterile deionized water to each well with a Robbins Hydra 96 (be sure to bleach and wash syringes between plates; the bleach solution is household bleach diluted 1:50, and the washes are done with sterile distilled water). Spin cell debris to bottom of wells (2000 RPM/2 minutes). Pool by sampling 75 ul from each well of a plate with the Hydra and dispensing 70 ul into a sterile 8-channel pooling block (this looks like a tiny little swimming pool with 8 horizontal lanes and a deep end). Mix the DNA well by rocking the block back and forth several times, then allow the mix to collect in the deep end. There should be 840 ul total. Aliquot this mix into a single column of four new 200 ul polypropylene deep-well PCR plates, using an 8-channel pipetter. The mixed samples from digestion plate A should go into Column 1, B into Column 2, and so on. These four plates are labeled CP-1 through CP-4, sealed with clean sterile silicone mat lids, and stored at ?20 until time to sample them for screening reactions. (?CP? means ?column pool,? which is kind of a misnomer since it is really rows of samples that are pooled. It just means that each plate is sampled into a single column of wells.)
Plate CP-4 and the original individual digestion plates should be sealed with mat lids and stored separately from the other three CP plates at ?20oC, and not touched again until time for row tests (see below).
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