2) For annealing, mix 3 µl (2-3 µg, ~1 picomole) of M13 template DNA; (or Bluescript ssDNA - see comments below) containing the sequences to be mutated, 3 µl of kinased primers (~70 picomoles) (directly from the above solution), 1 µl of a solution containing 200 mM Tris-HCl pH 7.5, 100 mM MgCl2, 500 mM NaCl and 10 mM DTT and 3 µl of H2O to complete 10 µl final volume. Heat at 55° for 5 min, then at 23° for 5 min.
3) For extension and ligation, mix: 10 µl of the above annealing reaction, 10 µl of a solution containing 20 mM Tris-HCl pH 7.5, 10 mM MgCl2, 10 mM DTT, 1 mM each deoxynucleotides and 0.1 mM ATP, 1 µl of a 1:2 dilution of BRL large fragment E. coli DNA polymerase (10 units/µl) and 1 µl of Biolab's T4 DNA ligase (400 cohesive end ligation units/µl), in a total volume of 22 µl. Incubate at 16° for 18-24 hrs.
4) OPTIONAL: For treatment with nuclease S1; mix: 2 µl of the solution after extension and ligation, 1 µl of 10X S1 buffer, 1 µl of a 1:100 dilution of BRL S1 nuclease diluted in 1X S1 buffer, and 6 µl of H2O to complete 10 µl final volume. Incubate at 37° for 15 minutes.
5) For transformation, mix: 10 µl of the S1 reaction mix and 100 µl of JM-101 competent cells. Let sit on ice for 60 min. For plating, add 200 µl of log phase JM101 in YT medium and 3 ml of 0.8% agarose in YT, preheated to 45°. Mix and plate over preheated (37°C) agar plates. Incubate at 37° overnight.
6) For screening by plaque hybridization: Cool plates in the cold for 60-90 min. Overlay a nitrocellulose filter over the plaques, press slightly and leave for 10 min. Carefully lift filters away from plate and leave them drying at room temperature for 60 min.
7) For lysis and fixing place filters over 3 M filter paper moisted with the following solutions: 5 min over 0.5 N NaOH-1.5 M NaCl, 5 min over 3 M NaCl-1M Tris-HCl (pH 7.5), 5 min again over 3 M NaCl-1 M Tris-HCl (pH 7.5) and 5 min over 2X SSC. Bake filters for 2 hrs at 80° under vacuum.
8) For pre-hybridization, incubate each filter in 5 ml of 0.2% SDS, 10% Denhardt's solution, 6X SSC at room temperature for 60 min.
9) For hybridization wash filters in 6X SSC for 5 min and incubate each filter in 5 ml of 10% Denhardt's, 6X SSC and 2x105 cpm of probe fragment (probe DNA fragment is kinased with -ATP to specific activity of 1-4x107 cpm/µg). Incubate at room temperature for 2 hrs.
10) Wash filters in 6X SSC at room temperature for 5 min. Expose to X-ray film for 2 hr with screen. To differentiate specific from non-specific hybridization wash again at higher temperatures, i.e. 40°, 55° etc. (depending on the length and composition of the probe), and expose to X-ray film.
11) For plaque purification, take the agar region containing the positive plaque and suspend in 5 ml of YT. This should contain a total of 108 to 109 pfu. Dilute this solution 200 times in YT to make a stock of approximately 106 pfu/ml. Dilute this again 100 times in YT to approximately 105 pfu/ml. Plate 10, 50 and 100 µl as described in step 5.
12) Repeat steps 6 to 10 as necessary.
13) Take isolated plaque and suspend in 1.5 ml of a 1:40 dilution of log phase JM101
in YT. Grow for 5 hrs at 37°. Centrifuge in 1.5 ml Eppendorf tubes. Save
supernatant for DNA sequencing and pellet for alkaline-SDS miniscreens.
Comments and Changes on Procedure
Purification of M13 and Bluescript Single Stranded DNA
This is my procedure for preparing M13 single strand DNA, but feel free to use
your normal procedure. One ml of a TG1 overnight is added to 300 ml of 2xYT.
Thirty minutes later add 20 ul of your M13 recombinant phage. Shake for ~ 7 hours,
then spin down bacteria for 10' at 6,000 RPM. Add to supernatent (300 ml) 20 ml of
5 M NaCl and 10g PEG. After PEG goes into solution let it sit for 10 min. on ice. Spin
10 min. at 6,000 RPM, pour off supernatent and let the pellet air dry. Resuspend the
pellet in ~5 ml of TE, phenol 2 times, chloroform 1 time and ethanol precipitate.
Resuspend the pellet in TE (as small a volume as easily possible, usually between 400
and 800 µl).
Many people use this M13-recombinant single strand DNA directly for
mutagenesisis. I however, first clean it up over an alkaline sucrose gradient. The
above single stranded DNA is contaminated with short fragments of DNA and/or
RNA that can prime the single stranded DNA in the Klenow reaction, resulting in
increased background and possibly double mutations. These short fragments are
removed on the alkaline sucrose gradient, which results in DNA that is primed only if
an exogenous oligonucleotide is added.
The alkaline sucrose gradient procedure is as follows: one half ml of a solution
that is 50% sucrose, 1 M NaCl, 0.2 N NaOH 1mM EDTA is added to a 1/2 inch
diameter x 2 inch polyallomer tube. A 5% to 20% gradient is then poured ontop of the
50% sucrose solution. This is done by putting 2.3 ml of a 20% sucrose solution (that is
1M NaCl, 0.2 N NaOH, 1 mM EDTA) into one slot of a gradient former (the slot
closest to the spout) and 2.3 ml of a 5% sucrose solution (1 M NaCl, 0.2 N NaOH, 1
mM EDTA) into the other. The gradient is then poured, don't worry if it isn't perfect.
50 to 200 µl of the single stranded DNA solution (100 to 200 µg) in TE is pipetted
onto the top of the gradient. The tubes are then spun for 5 hours at 45,000 RPM at
16° C in a pre-cooled SW 50.1. The gradient is then dripped from the bottom in 200 to
250 µl fractions and the fractions assayed by O.D. The peak 2 to 4 fractions are
usually at fraction 9 or 10. The purified DNA is neutralized by adding 1/10 volume
of 1M tris pH 8.0 and 1/10 volume of 2 M HCl, and thenn 2.5 volumes of ethanol
added and the DNA precipitated. The DNA is resuspended in TE and brought to a
concentration of 1 mg/ml. Final recovery is usually between 30-50% of input DNA.
This mutagenesis protocol also works well for single-stranded templates in the
Bluescript vector system (Stratagene, Inc.). Single-stranded Bluescript DNA is
prepared as for DNA sequencing according to the Bluescript protocol booklet. The
yield from a small-scale ssDNA prep (1.5 ml culture) suffices for 2-3 mutagenesis
reactions.
Comment on Kinasing Oligonucleotide
My procedure is as follows: The oligonucleotide is diluted into the 30 µl
reaction so that the O.D. 260 of the oligonucleotide in the kinase reaction is 4.0. As an
example, if the O.D. 260 of a 1/100 dilution of the oligonucleotide is 0.295, the the
O.D. 260 of the oligonuleotide is 29.5 and I want to dilute it by a factor of 0.135
(4.0/29.5) and therefore will add 4.05 µl (0.135 x 30µl) to the 30 µl reaction. I will of
course round the 4.05 µl to 4 µl. Everything is then done as above except that I only
do one half hour reaction and I don't bother to kill the enzyme at 65°C.
Comment on Annealing
I do exactly as they suggest except that I heat the sample at 65° instead of 55° (I
don't have a 55° water bath) and I allow hybridization at room temperature instead of
23° (I don't have a 23° water bath).
Comment on Extension and Ligation
I add 1/2 µl of both Klenow and ligase without diluting it.
Comment on S1 Treatment
Instead of just treating all the DNA with S1 nuclease for 15 min., I always
perform a titration, removing 2.5 µl aliquots of the reaction at 0, 1, 5 and 15 min. I
find that for some reactions, a 15' incubation is too long and you get no plaques. The
titration allows you to knock down the background but still retain enough plaques to
work with. I should also add that you can skip the S1 treatment altogether, although
this will usually decrease the number percentage of phage carrying the mutation.
However, when using an oligonucleotide that will create an insertion or deletion I
sometimes find that S1 treatment will decrease the percentage of plaques that contain
the desired mutation, presumably because the S1 directly digests the extended and
ligated DNA at the loopout caused by the oligonucleotide.
Comment on Transformation and Plating
I follow my normal transformation procedure: mix 2.5 µl of the S1 reaction mix
and 100 µl of TG1 competent cells. Let sit in ice for 10 min. Heat at 37° for 5 minutes.
Add 1 ml of 2xYT at room temp. For plating, mix 200 µl of the M13-transformed TG1
cells with 100 µl of TG1 overnight and 3 ml of 0.8% agarose in YT, preheated to 45°.
Mix and plate over agar plates. Incubate at 37° overnight. Once the plaques are
visible, plate out all or some of the remaining M13-transformed TG1 cells (which have
been stored at 4°C) so that the plaques will be well separated. I find that between 40
to ~150 plaques per plate are ideal.
For Bluescript templates, you should do a standard bacterial transformation of
the S1 reaction mix and plate out on Amp plates to obtain colonies rather than
plaques. Properly mutagenized templates are then detected by colony hybridizations
rather than plaque hybridizations, using the same conditions as described in steps
6-10 above.
Comment on Making Probe
I mix 1 µl of oligonucleotide (diluted to O.D. 260 = 1) with 1 µl of 10x kinase, 2
µl of 32P-ATP (10mCi in ~50µl, carrier free from ICN), 6µl of water and 1/2 µl of
kinase. Incubate for 30 min. at 37°C. I use 1/2 of this mixture directly for screening,
you don't have to put it over a column.
Comment on Plaque Purification
Even if it is obvious which plaque contains the transformant I always plaque
purify it because some of the plaques contain a mixture of mutated and wild type
genomes. Instead of diluting to plaque purify I streak out the plaque in the same
manner you would a bacterial colony and then very gently add 3.5 ml of top agar
containing 100 µl of a TG1 overnight.
Comment on Repeating
This is usually not needed.
Comment on Sequencing
I usually prep 4 plaques for sequencing.