Gene Knockout with Conventional Mutagens

Gene Knockout with Conventional Mutagens

Leon Avery

ENG (enriched nematode growth medium): 
1.  in a 6 L flask add:
 3800 ml DH2O
 20 g. bactopeptone
 4 g yeast extract
 12 g. NaCl
 4 ml 5mg/ml cholesterol in EtOH
stir with a large stir bar until dissolved.

2. Add 120 g. agar and stir well.

3. Cover w/ Al foil, autoclave liq. cycle 30'
   (turn on the 70C water bath before the agar is autoclaved)

4. Equilibrate temp. in water bath.

5. Just before pouring the plates add (sterile solutions) with
   stirring:
 4 ml 1M CaCl2
 4 ml 1M MgSO4
 100 ml 1M KHPO4 (pH6)
        4 ml 200 mg/ml streptomycin
 1 ml 40 mg/ml nystatin in DMF (add dropwise)

AENG (agarose ENG):
Same recipe as ENG, except replace 3% agar with 2% agarose, and you
probably won't want to make so much.

Making a grid:

1. Inoculate one or more 10 cm ENG plates with 10 N2 L4
   hermaphrodites.  Four or five days later, when there are many
   gravid adults, prepare eggs by the alkaline hypochlorite method.
   Leave the eggs in M9 after the final wash.

2. Put the tube of eggs in M9 on a rocker (or something else that will
   aerate them) overnight.

3. Count the healthy L1s in a 10 ul aliquot of the suspension.  (I use
   a 10 ul microcap to spread them on an unseeded 6 cm plate.)  Based
   on the count, put 5000 healthy L1s on each of 10 10 cm ENG plates.
   These will be your P0s.

4. When the P0s have reached L4, harvest them in 3 ml M9.  (You'll
   probably have to spin them down and transfer them to new M9 to get
   the volume that low.)

5. Dissolve 20 ul EMS in 1 ml M9.  Add this solution to your P0s.

6. Aerate for 4 hours.

7. Spin down, transfer to 4 ml fresh M9.

8. Put 5000 P0s on each of 10 10 cm ENG plates.  Allow them to become
   gravid adults with lots of eggs.  (I do this at 15C, since the L4s
   will become gravid adults inconveniently soon at 20C.)

9. Harvest all the plates and prepare eggs by alkaline
   hypochlorite.  Aerate overnight.

10. Count the healthy F1 L1s in a 10 ul aliquot.  There will be a lot
   of sick or dead ones (their parents were mutagenized) -- don't
   count these.
   Determining numbers of F1s


11. Put 1250 F1 L1s on each of 110 6 cm AENG plates.
   F1s per plate


12. After 3 days at 20C most of the F1s should be gravid adults.
   Count the gravid adults on one or two of your plates.  There should
   be about 1000 (less than 1250, because many of the F1s didn't grow
   up or were sterile).
   F1s per plate


13. After another 2 days (5 days total) the plates have starved, and
   almost all the worms on them are L1s.  Harvest 96 of the plates
   with 5 ml M9 each.  Discard any extra plates -- they're just in
   case some of your plates got moldy, or otherwise didn't work.
   Harvesting the plates


14. Count a 10 ul aliquot from a few of the tubes.  You should have
   recovered between 100,000 and 200,000 F2 L1s from each plate.
   F2s per plate


15. Let the tubes sit on your bench until the worms have settled to
   the bottom.  with a Pasteur pipet attached to vacuum, suck off
   liquid so that you have about 1.5 ml left in each tube.  (This is
   the step I worry about -- I'm afraid if the worms sit at the bottom
   of the tube too long they may go anaerobic and die.  I do plates in
   batches, so that they don't have to sit too long, and mix the tubes
   from the earlier batches occasionally while working on the later
   ones.)

16. For this step, I use sterile 1.2 ml tubes in an 8x12
   microtiter-spaced rack.  You want three racks of 96, two for worms
   to be recovered later, and one for DNA preps.  Put 0.5 ml of 2x
   freezing solution in each of the tubes of the two worm racks.
   Then, for the first of your 96 tubes, put 0.5 ml into the A1
   position of each of the three racks.  For the second, put 0.5 ml
   into the A2 position, etc.

17. Put the two worm racks into a styrofoam box in a -80C freezer and
   leave overnight.  You can freeze the worms for the DNA preps now,
   or go ahead and do the DNA preps immediately.

18. Let the worms settle to the bottoms of the tubes, and suck off all
   the liquid you can.
   DNA preps


19. Add 250 ul fresh lysis solution (200 mM NaCl, 100 mM Tris-HCl pH
   8.5, 50 mM EDTA, 0.5% SDS, 100 ug/ml proteinase K) to each tube.

20. 50C, 1h.  (This is shorter and cooler than the typical incubations
   used for DNA preps.  It works fine, and long incubations at high T
   are mutagenic.)

21. Prepare DNA from each tube by phenol:sevag extraction, sevag
   extraction, and EtOH precipitation.  Redissolve in 150 ul TER (10
   mM Tris-HCl pH 8, 1 mM EDTA, 1 ug/ml RNAase A.  Store at -80C.

22. Use 2 ul of 1:5 diluted DNA in each detection reaction.

Mutation detection protocols:

I have used three different protocols for detecting mutations.  The
first is for detecting point mutations in a restriction site.  I won't
describe this, since I don't currently think it's the best way to
knock out a gene.  The second method is that commonly used for
detecting deletion of a Tc1 by relying on the much greater efficiency
of standard PCR in amplifying shorter fragments.  I call this method
"primer approximation", since it relies on the deletion approximating
(bringing close together) the primers.  The third method asks for the
deletion of a cluster of restriction enzyme sites.  It has the
theoretical advantage that it may detect small deletions that don't
much change the efficiency of amplification.  Whether this is a
practical advantage is not clear.

I do not use nested primers.  I think this reduces the false positive
rate, but it does mean the primers have to be pretty good, since you
need to be able to amplify from as little as 10 molecules to a good
strong band.  Therefore, before screening with any primer pair, I
first run a series of reactions inoculated with 0, 10, 1000, and
100,000 wild-type genomes, and run under long PCR conditions
(basically the conditions for site deletion below).  I make 25-mer
primers with 40-60% GC, using the Whitehead Institute primer program
to help find them and avoid primer-dimers, etc.  Most of the pairs
work.

Mutation detection by primer approximation:

Solution A:
    0.4 ul 5 U/ul Taq polymerase
    5   ul 10x PCR buffer (100 mM Tris pH 8.3, 500 mM KCl, 20 mM MgCl2)
    0.5 ul 10 mg/ml acetylated BSA (NEB)
    1   ul 10 uM left primer
    1   ul 10 uM right primer
    1   ul 10 mM dNTPs (10 mM each of dATP, dCTP, dGTP, dTTP)
   39.1 ul water
   -------
   48   ul

Put 48 ul A in each of 96 thin-wall 200 ul tubes.  Add 2 ul 1:5
diluted DNA to each tube.  Run PCR program (MJR PTC-200):

    Control method: CALCULATED
    1: 92 degrees forever, beep.
    2: 92 degrees, 55 sec.
    3: 92 degrees, 5 sec.
    4: 65 degrees, 30 sec (adjust for the Tm of your primers).
    5: 68 degrees, 1 min.
    6: go to step 3 for 39 more cycles.
    7: 68 degrees, 4 min.
    8: 4 degrees forever, beep.

Start the PCR machine and wait for the block to heat up.  Now put the
tubes (kept on ice) in the block.  The solution in thin-wall tubes
will heat up very fast when placed in a preheated block: this is
almost as good as a hot start.  Push the proceed button on the PCR
machine once all the tubes are loaded.

Add 10 ul 6X gel loading buffer and run 12 ul on 1% gel.


Mutation detection by site deletion:

In this example, the enzyme BstBI is used.  You need to use a
thermophilic enzyme, since cooling the reaction down to 37C for the
second digestion has disastrous effects on PCR.  You also want an
enzyme that will work pretty well in PCR buffer, and that is pretty
cheap.  There are a whole series of enzymes from Bacillus
stereothermophilus that meet these criteria.

Solution A:
    0.25 ul 20 U/ul BstBI, NEB
    1 ul 10x NEBuffer 4
    0.1 ul 10 mg/ml acetylated BSA, NEB
    6.65 ul water
    ------
    8.0 ul

Solution B:
    0.01 ul 5 U/ul Pwo polymerase
    0.09 ul 5 U/ul Taq polymerase
    3.5 ul 10x PCR-NEB4 (75 mM tris pH 8.6, 500 mM KCl)
    0.35 ul 10 mg/ml AcBSA
    1 ul 10 uM primer 1
    1 ul 10 uM primer 2
    1 ul 10 mM dNTPs
    28.05 ul water
    --------
    35.0 ul

Solution C:
    0.3 ul 5 U/ul Taq polymerase
    0.5 ul 20 U/ul BstBI
    0.5 ul 10x PCR-NEB4
    0.05 ul 10 mg/ml AcBSA
    3.65 ul water
    -------
    5.0 ul

LAMD45 PCR program (CALC mode, MJR PTC-200):
    1 92C forever, beep
    2 92C, 1 min
    3 65C, 30 sec (annealing temp)
    4 68C, 5 min
    5 65C forever, beep (T in steps 5 & 6 is digestion T for enzyme)
    6 65C, 30 min
    7 92C, 5 sec
    8 65C, 30 sec
    9 68C, 2 min
    10 go to step 7 for 8 more cycles
    11 92C, 5 sec
    12 65C, 30 sec (annealing temp)
    13 68C, 2 min + 10 sec/cycle
    14 go to step 11 for 34 more cycles
    15 72C, 10 min
    16 4C forever, beep

1.  Mix 2 ul DNA solution (intended to be about 100,000 genomes) with
    8 ul A.  Incubate 65C, 10 min.

2.  Add 35 ul B to each tube.  Keep everything on ice while you do
    this.

3.  Start the PCR machine.  When it reaches 92C (step 1), transfer the
    tubes from the ice to the block.  Once all the tubes are in the
    block, tell the machine to proceed.

4.  At step 5, the machine will halt again at 65C.  Leaving the tubes
    in the block, add 5 ul C to each.  When finished, tell the machine
    to proceed.

6.  Run on 1% agarose gel.