I. Preparing worm genomic DNA: requires 1-2 days to seed agarose plates, a few days for the worms to grow, 1-2 days to prep the DNA
1. Seed large agarose plates with HB101. Agarose is preferred over agar since the latter contains more inhibitors of restriction enzymes. HB101 is healthier than OP50, and produces a thicker lawn allowing a higher yield of worms. Expect a yield of ~10-130 µg of genomic DNA per large plate, averaging around 40 µg, so scale up accordingly. For Southerns, you will use 1-5 µg per lane.
2. Harvest the worms before the plates starve. After the bacteria are gone, the worms begin breaking down the agarose and releasing enzyme inhibitors. It is best to harvest when the worms are forming a wave across the plate as they eat the last bacteria.
3. Rinse the worms off the plate in M9, spin down briefly (30") in a clinical centrifuge. May want to wash once with more M9 to remove bacteria.
4. Optional step: this is usually totally unnecessary. To remove agarose contamination, can float the worms in sucrose. Resuspend the worms in 5 ml M9, put on ice. Place 5 ml cold 60% sucrose in a 15 ml polypropelene tube. Shake the worms and carefully layer on top of the sucrose with a pasteur pipette. Spin in a clinical centrifuge at 4° for 5'. Remove the top layer, which contains clean healthy worms, to a new tube, add some more M9 to wash, and pellet the worms.
5. Wash and pellet the worms in 4 ml TEN. Remove the supernatant, resuspend to a total volume of .5 ml (note: use less than ~200 µl worms here, if you have more then split into two tubes. Should get 200 µl worms from 1-3 large plates). Move to a 1.5 ml eppendorf tube. At this point, can throw the worms into the -20° freezer to synchronize preps of different strains. Freezing/thawing here helps the following proteinase K digestions, and so is useful even if you don't want to store the worms.
6. To the worms in 500µl TEN in a 1.5 ml eppendorf tube add:
25 µl 10% SDS (final conc. 0.5%)
2.5 µl 20 mg/ml Proteinase K ( I make this fresh) (final conc. 0.1 mg/ml)
1.0 µl beta-mercaptoethanol
- Incubate in a 50 to 60° water bath; gently resuspend by inversion at 1', 3', 10' to achieve a ~uniform suspension instead of one big glop floating in liquid.
- After one hour add another 2.5 µl proteinase K, and put back in the water bath.
- After the second hour, add another 2.5 µl proteinase K, put back in the water bath.
- At the end of the third hour, you should have a uniform not very viscous milky yellow solution. If there is still gloppy worm goop present, can let the digestion go overnight.
7. Phenol/chloroform extract by adding 0.5 ml 50/50 phenol/chloroform (isoamyl alchohol). The Horvitz lab uses a commercial preparation of this from American Bioanylytical (#AB1605). At this step and all subsequent ones, treat the DNA very gently to avoid shearing. Mix the aqueous and phenol phases by gentle inversion. Then centrifuge for 2 min at room temp. Transfer the upper aqueous phase to a new 2 ml eppendorf tube using a cut off 200 µl tip (be clean, not greedy). To increase the yield, reextract the phenol/CIA phase with 200 µl TEN, and combine the two aqueous sups. If the protein interface was extensive, can repeat the phenol/CIA extraction.
8. Add 1.2 ml EtOH at room temp, and mix by inverting gently, and flicking the tube to produce a gentle swirling that winds the DNA strands. Should get a precipitated clot of DNA almost immediately. Do not spin the tube: instead pick the DNA clot out with a 200 µl tip on a pipetteman (without sucking) and place in a tube of 70% ice cold EtOH to wash. This gives cleaner DNA than pelleting. (If no ppt. is visible, you can spin the tube, but the yield of DNA is probably poor). Spin the 70% tube very briefly (10"), remove the supernatant, and dry the pellet only very briefly; want it to remain moist with 70% EtOH for easier resuspension.
9. Resuspend the pellet in 0.5 ml TEN. This is a difficult resuspension and may require the following sequence:
- Let the pellet rehydrate at 4° a few hours to overnight.
- Dissolve the pellet more by putting at 50-60° for 1-2 hours. At this step you may accelerate the resuspension by breaking the pellet up using a pipetteman and a 200 µl tip to gently suck up and spit out the pellet, breaking it into small pieces. This treatment appears not to shear the DNA enough to show up in the Southerns.
- Be careful, because the pellet often becomes crystal clear after rehydrating, and can only be seen due to its refraction. Again, be very gentle (no vortexing) so as not to shear the DNA.
10. Add 3 µl 10 mg/ml boiled RNase A (store this stock frozen) (final conc. 40 µg/ml). Incubate 37° for 1-2 hour. Phenol/CIA extract as before. Chloroform extract (I often don't use isoamyl alchohol here). Add 2 volumes (~0.8 ml) 100% EtOH at room temp. Invert and flick as before, and again pick the pellet out with a pipette tip to a tube of ice cold 70% EtOH. The nucleic acid precipitate will be substantially smaller this time due to the absence of the RNA. Let the 70% tube sit at room temp. for 1 hour to remove salts. Spin 10", remove the sup, dry the pellet, and resuspend in TE (~100 µl per large plate of worms). Again, this resuspension is difficult and may require rehydrating overnight at 4° followed by a couple hours at 50-60°. If the pellet still won't resuspend, try adding more TE.
TEN: 20 mM Tris pH 7.5, 50 mM EDTA, 100 mM NaCl
II. Restriction digestion of worm genomic DNA: takes overnight
Achieving complete digests of genomic DNA is problematic in some people's hands. To ensure complete digests, try using 10 units of restriction enzyme per µg of DNA and digesting overnight (note: this is only useful for stable restriction enzymes- the Biolabs catalog has a table showing the stability of various enzymes). EcoRI is excellent at cutting genomic DNA, HindIII is good, others vary. After digestion, run an aliquot on an analytical gel to check the digestion: should see a smear with a bell-shaped distribution of intensity. The average size varies according to the length of the recognition sequence, and also the GC content of the recognition site; AT rich sites occur more frequently. You don't want to see DNA hanging up at the mobility limit of the gel; you do want to see a few tight bands within the smear (these are repetitive sequences). EcoRV and XbaI are other 6-cutters with AT-rich recognition sites that work okay.
Other tricks for achieving complete digests: some people routinely add 5 mM spermidine to their digests, which reputedly soaks up inhibitors of restriction endonucleases. Another for sure help is to dilute the digest into a larger volume (thus diluting the enzyme inhibitors) and then precipitate and resuspend the cut DNA in the desired volume. You should be able to digest 5 µg DNA in a 20 µl digest with clean DNA, but diluting 10-fold will help if this doesn't work. For genomic DNA prepared as above, these tricks aren't necessary for EcoRI or HindIII digests.
III. Running the gel; takes overnight
Typically, worm DNA is cut with a six-cutter enzyme and a whole cosmid or Tc1 probe is used. In these cases, many bands in the ~1-10 kb range will be detected, and thus it is important to achieve good separation of fragments in this range.
Typical gel conditions: use a 35 cm long x 15 cm wide gel box. The gel poured inside is 23 cm long. Pour a 0.8% agarose gel in 1X TAE, with EtBR in the gel and running buffer. Don't pour the gel any thicker than necessary; this will just interfere with the transfer. Use a 20 well comb (each well is 4 mm wide). Run the gel overnight (11-13 hours) at 125 volts, recirculating the buffer slowly with a peristaltic pump (we use "masterflex" pumps from Cole Parmer Instruments). Run the pump about as slowly as it goes; may want to run the DNA in for an hour or so before turning the pump on so as not to disturb the DNA in the wells. (If you run the pump too fast there is a danger that the gel will be floated off to the side of the gel box by the flow of the buffer!) Load 1-5 µg DNA per lane. Using the large Owl Scientific boxes I pour a 300 ml gel, and it takes 13 hours to run the bromophenol blue to the bottom of the gel.
To achieve a significant increase in separation in the 1-10 kb range, run the gel using a field inverter box between the power supply and the gel box. (Note that the cheap blue "Blair Craft Scientific" power supplies don't really supply steady direct current and shouldn't be used with the field inverter; use the Biorad or Hoeffer DC power supplies). We use the MJ Research "Programmable Power Inverter" model PPI-200 with the following parameters:
A= 0.1 B= 0.01 C= 0.3 D= 0.03 E= 11 F=0 G=0
Plug the leads from the power inverter into the power supply, and connect leads from the gel box to the female sockets in the inverter box. Can run two gels in parallel (not series!) off of one inverter box/power inverter. After setting and checking the parameters, turn the power supply to 125 volts, and then remember to press enter on the inverter to begin the inversion program.
IV. Southern transfer; takes a few hours to process the gel, overnight transfer, and a couple hours the next day to bake the blot.
After the gel is run, take a picture on the UV light box with a ruler laid next to the gel. Later, will use this to measure out the size of the bands on the Southern blot. I cut off the lower left corner of the gel to help remember the orientation. While at the light box, leave the gel on the 250 nm light box with the light on for 1 min. This nicks the DNA and allows more efficient transfer of high molecular weight fragments out of the gel. (Note: don't leave it on the UV box too long: 5 minutes nicks the DNA so much that it will be in tiny pieces that will diffuse in the gel giving fuzzy bands on the blot.)
In subsequent steps, handle the gel very gently to avoid breaking it. Soak the gel in several volumes of denaturing solution for 2X15' with gentle agitation. Rinse briefly with neutralizing solution, and soak 2X30' in several volumes neutralizing solution.
Cut ~4 sheets of Whatmann 3MM or equivalent and a sheet of Nytran (Schleicher and Schuel) to the size of the gel. Always handle the Nytran with clean gloves. Cut off the corner of the Nytran that will be over the lower left corner of the gel to later orient the blot. Nytran is preferred over nitrocellulose because it doesn't tear as readily. Wet the 3MM in a dish of 20X SSPE. Nytran is best wetted first in distilled water, and then transferred to 20X SSPE. Place commercial kitchen sponge(s) in a tray; use several sponges side by side if the gel is bigger than one sponge. Fill the try up to ~1 cm below the top of the sponges with 20X SSPE. Lay two sheets wet 3MM on top; smooth them down to remove air bubbles. Lay the gel upside down (i.e. wells down) on the 3MM, smooth out the air bubbles with a wet gloved finger. Place strips of Parafilm along each side of the gel; these will stop buffer from wicking up except through the gel. Lay the Nytran on the gel with the notched corner of the Nytran matching the notched corner of the gel. Smooth out air bubbles. Do not adjust the Nytran once it is placed on the gel. Lay two sheets of wet 3MM on top; smooth out the bubbles. Lay several inches of dry paper towels on top. Place a glass plate or equivalent on top and ~200 gm weight to ensure even contact within the stack of towels. Wait overnight; do not disturb during the transfer.
The next morning, remove the paper towels and upper 3MM sheets, but do not remove the Nytran! Before removing the Nytran, use a dull soft pencil to mark the position of the wells on the Nytran. Ballpoint pen also works, but the marks may run a little. Then remove the Nytran; place between two sheets of dry 3MM. The gel should be a dry smashed flat rubbery thing at this point. After the blot air dries (~15 min), bake it in a vacuum oven at 80° for 2 hours.
Some people UV cross link the DNA to the Nytran. Crosslink while the blot is still moist in a Stratalinker; rinse the blot in 2X SSPE, dab off excess liquid on 3MM paper, place the moist filter in the machine DNA side up, press the on button, press the auto cross link button, press the start button. The machine counts down as it monitors the UV to give the optimal dose. Then take the blot out, turn the power off with the power button, and bake the blot. Of course, baking isn't actually necessary after crosslinking, but you can do it if you want.
Denaturing solution: 0.5 M NaOH, 1.5 M NaCl
Neutralizing solution: 1M Tris pH 8, 1.5 M NaCl
20X SSPE: 175.3 g NaCl, 27.6 g NaH2PO4.H20, 7.4 g EDTA: dissolve in ~800 ml H20, adjust to pH 7.4 w/ ~6.5 ml 10 N NaOH. Adjust volume to 1 liter.
V. Probing the blot; a few hours to make the probe and prehyb, overnight to incubate blot and probe, a few hours to wash the next day, 1-4 days to expose the autorad.
1. Labelling the probe-obviously, follow all the normal radioactivity safety precautions
- If you store it frozen, take the alpha 32P-dATP out of the freezer to thaw.
- Put 50 ng DNA to be labelled in a total volume of 18.9 µl H20. This is usually 1-4 µl of a typical cosmid prep. It is unnecessary to cut cosmid DNA prior to labelling. For labelling fragments from a restriction digest: I prefer not to do the labelling in the low melt agarose, since the agarose somewhat inhibits the labelling reaction and the specific activity of the resulting probe is only marginally adequate for genomic Southerns. It is easy to purify the fragment either using agarase, or by adsorption to glass as in the commercial Qiex kit, and this will increase the efficiency of labelling considerably.
- Boil the DNA to denature it for a few minutes. Briefly spin the tube and put on ice to cool. Add 6.6 µl 5X OLB, 5 µl hot dATP (50 µCi), and 2 µl Klenow.
- Allow the labelling reaction to proceed at room temp 30 min to overnight. The reaction appears to go to completion by 45 min.
- Some people use the entire reaction in their hybridization. However, anyone intelligent will run a spin column at this point and measure the amount of counts incorporated, since this is the main factor determining the strength of signal on the autorad. Add about 20 µl of 50 mM EDTA to stop the reaction and run a spin column as usual. Expect about 50% of the counts to be incorporated for an average labelling.
2. Prehybridization of the blot- can do this while the probe labels.
Wet the blot in a tray of 6X SSPE. Make some prehybridization buffer. Put the blot in a hybridization tube (or a seal a meal bag), and add the buffer (theoretically ~0.5 ml per square inch of blot, in practice I use ~30 mls for a large Hybaid tube), and incubate at 65° for a couple hours or longer. I highly recommend using the "Hybaid" hybridization oven with the roller in it for glass tubes. It maintains temperature accurately, and minimizes your exposure to radioactivity and chance of having a spill. To get the blots into the tubes, wet them first, fill the tube ~1/3 full of 6X SSPE, and use a glass rod (a thermometer will do if you are very careful) to help shove the blot into the tube and to plaster it against the wall of the tube.When using glass hybridization tubes, don't tighten the caps too much: they tighten up in the oven. Make sure the cap of the tube has a red rubber O ring in it; the other type of cap can leak.
3. Hybridization: pour the prehyb sol'n out. Add hybridization solution (I use 20 mls for a large Hybaid tube) to the blot. Denature the probe DNA by A) heating to 100° C for 5-10 min, or B) add NaOH to 0.2 M, incubate 5' room temp (the SSPE will buffer this out when you add it to the hyb solution). Add the probe to the blot/hyb. solution, close the tube/bag, and incubate overnight at 65°.
4. Posthybridization: Make 1 liter of post hyb. wash solution, and heat a shaking water bath to 65°. Pour out the hyb solution into a 50 ml plastic disposable centrifuge tube if it is to be reused (probe is good for a couple of weeks, and must be boiled 5 min and chilled on ice before reuse). Rinse the blot (in the tube) briefly with some wash solution. Remove the blot to a tupperware dish. Add a generous amount (a few hundred mls) of wash solution. Wash the blot 3X30' in the 65° water bath. At this point the geiger counter should not detect counts on the blot if you are just probing for single copy genomic sequences. When probing with TC1 or other repetitive sequence probes, the signal may be hot enough to detect with the geiger counter.
Dab the blot on 3MM paper to get rid of excess liquid, wrap the moist blot in Saran wrap, tape to an old autorad, apply a strip of phosphorescent marker next to it (Stratagene), and take an exposure with an intensifying screen at -80°. Single copy genomic Southerns require overnight to 4 day exposures with whole cosmid probes. I've gotten blots that require only 1 hour exposures using labelled restriction fragment probes. Always keep the blot wet and frozen, so that it can be stripped and reused.
5X OLB: (oligonucleotide labelling buffer)
250 mM Tris pH 8.0 250 µl 1M
25 mM MgCl2 25 µl 1M
5 mM ß-mercaptoethanol 0.35 µl 14.4 M stock
2 mM dCTP 20 µl 100 mM stock
2 mM dGTP 20 µl 100 mM stock
2 mM dTTP 20 µl 100 mM stock
1 M HEPES pH 6.6 500 µl 2 M (2.38 g + 4ml H20, adjust to 6.6 w/ NaOH, adjust vol to 5 ml)
1 mg/ml oligonucleotides (random primers: Pharmacia # 27-2166-01)
H20 to 1 ml
Prehybridization solution: note that some leave out the Denhardt's
30 ml 20X SSPE
1 ml boiled 10 mg/ml salmon sperm DNA (store frozen in 1 ml aliquots)
10 ml 50X Denhardt's solutions (see below)
5 ml 10% SDS (add some H20 first to prevent this from ppting)
H20 to 100 mls
Hybridization solution: Same as prehyb sol'n above except without the Denhardt's
50X Denhardt's solution:
5 g Ficoll
5 g polyvinylpyrrolidone
5 g BSA (pentax fraction V)
500 ml H20
Mix overnight. Then filter thru a disposable filter unit and freeze in 10 ml aliquots. Alternatively, add EDTA to 5 mM and store at 4°.
Posthybridization wash solution: 0.2X SSPE, 0.5 % SDS
Recipe: 10 ml 20X SSPE, 940 ml H2O, 50 ml 10% SDS
VI. Stripping the blot for reprobing.
Hybridized radioactive probes can be stripped off the blot and the blot reprobed several times. Alternatively, if the blot has sat in the freezer for several months to the point where the radioactive signal has decayed to an insignificant level, it can be reprobed directly without stripping.
1. Prepare 3 erlenmeyer flasks each with 500 ml of 0.05x SSPE, 0.01 M EDTA
Recipe: Add 1.25 ml 20x SSPE and 10 ml of 0.5 M EDTA (pH8.0) to 500 ml dH20.
2. Heat first flask in microwave oven 6 min to boiling. Immediately add SDS to final concentration of 0.1% (5 ml 10% SDS), and gently place the blot into the hot solution (using a glass rod to push it in helps).
3. Put the flask in a 65° water bath for 15 min.
4. Repeat steps 2 and 3 with the next two flasks of stripping solution.
5. Rinse briefly in 0.01x SSPE at room temp (0.25 ml 20x SSPE in 500 ml dH20).
6. Remove excess liquid by dabbing the blot on 3MM paper. Wrap in Saran wrap, and expose overnight to check for the absence of radioactivity.
7. To reuse the blot, prehyb and hyb. as usual.
Warning: Probes irreversibly bind to the filter if it is ever allowed to dry. Therefore, make sure the blot is always moist at all stages during hybridization, washing, exposure to film, and storage, if you want to reuse it.