hi

shifali chatrath via methods%40net.bio.net (by shifalich from rediffmail.com)
Sat Oct 10 09:56:18 EST 2009


Thanks DK!
The following info was quite useful for me too.
We can't find such details anywhere!!!!
Few months back, I was trying to use pfu but my PCRs did n't work, so I gave up after 2-3 trials. Now, I'll using thse tips.

Thanks again
REgards
Shifali

On Sat, 10 Oct 2009 03:29:11 +0530  wrote
>In article , monika singh  wrote:
>hi all,
>
>i am doing site directed mutagenesis for single amino acid change .i have
>used both stratagene as well as phusion kit but i didnot get the mutation .
>i have also found out that the melting temp. of primers which i am using is
>between 60 to 65 degrees. But according to stratagene primer guidelines it
>should be greater than or equals to 78 degrres. So, this could be the
>possibility of not working of my mutation. also,i have used top 10 cells
>ultra comp cells for transformation. i get the colonies also but it was not
>positive. so, suggest me regarding com cells, pcr conditions ,dpn1 digestion
>so that i can get result

Without doing all kind of controls, you can't know where the problem is.
The calculation of primers melting temperature is somewhat of a joke
as it can vary over 10 degrees in either direction depending on the 
program you use. Competent cells have nothing to with it as long as 
you do get colonies. 

Below is a protocol that I wrote for our lab internal use long ago. 
It practically never fails. We've done all kind of mutagenesis with
it, deletions of >1 kbp and insertions of up to 36 bp. 

>>>>
Although the regular QuickChange works, it does not always work 
well enough. There are several advantages to the new protocol: 
- twice less money spent on primers;
- twice less polymerase is used; 
- polymerase with lower error rate is used. 

Here is a generic protocol followed by notes and comments that might 
be helpful. 

    Reaction mix (25 ul final) 
50 ng template 
0.2 uM final single primer 
0.2 mM final dNTP mix
2.5 ul 10X Stratagene's PfuUltra II Fusion ("Pfu Fusion" for brevity) buffer 
1.0 ul DMSO (or no DMSO)
water to 24.5 ml
0.5 ml of Pfu Fusion 

Cycling conditions
95�  3 min, then 30 cycles:
    95�   30 sec
    55�   1 min
    65�   1 min/kbp template
65�  5 min
4�   hold

- Dpn I treatment and transformation:
Use Fermentas "Quick Digest" DpnI - it is simply a more concentrated 
version of the enzyme and it costs the same as regular version. Add 
10 ul of the amplification reaction to 0.5 ul DpnI, incubate for 2 hours 
in PCR machine at 37�. During incubation, at least once remove the 
tube and vortex/spin it again, then continue incubation. 
- Electroporate using 1-2 ul of the DpnI-treated reaction, immediately 
add 0.5-1 ml SOC, let cells recover for 45 min at 37�, plate 200 and 
20 ml (recovery is not absolutely necessary but it does seem to increase 
efficiency somewhat). The typical result is several-fold less colonies 
than in a "standard" QuickChange reaction, and the efficiency in the 
50-90% range. I.e., screen 5-6 clones to be sure to get right one no 
matter what. 

NOTES AND COMMENTS
(in no particular order; I am just mentioning everything that I can think 
of being worth consideration)

1. Template: 
Ideally, it should be a fresh, never frozen miniprep. Freeze/thaw
produces nicks in the plasmid and nicks are killers to the whole
plasmid amplification. It also helps for it to bew "clean" so that
UV measures concentration of the plasmid and not just nucleic 
acids with RNA being a major contaminant. 

2. The Pfu Fusion polymerase is a new product that is slightly less 
expensive than any of the other polymerases we use. It works ~ 4 
times faster, makes a bit less errors than Pfu Ultra and its 
reaction buffer has pH of ~ 10.0. 

3. The method requires only one primer - just write down the 
sequence of the sense strand! It's been found in numerous 
trials by that desalted primers from IDT work as well as purified 
ones. 

4. Primer design. Keep primer length between 40 and 60 bp. 
Try not to exceed 60 bp because above that size IDT forces you 
to order 100 nmol scale with the corresponding price increase. 
Ideally, you'd want 15-25 bp on either side of the mismatch, 
approximately equal melting temperature (TM) for both "arms" of 
the primer (but the "right arm", e.g. 3' part, is more significant because 
that's where synthesis starts; try to end it with a couple of G/Cs). 
The overall primer TM (not counting the mismatched pairs) should 
be in the range of 60-70�C (as calculated by IDT's Oligo Analyzer 
with default settings). As an example, the most recent primer that 
worked 100% to delete 6 bases: Left arm 27 bp/51.1�, right arm 
19 bp/53.4�, overall 46 bp/63.8� and it worked with 55� annealing. 

5. In linear amplification applications (sequencing, QuikChange), 
the larger number of cycles gives better results. 30X has been tried 
and works very well. To be sure, 20X also works, although it was 
not compared to 30X side by side. 

6. Amount of template. The stated 50 ng is simply lifted from what 
we find a near optimum for the standard QuickChanges, 100 ng/50 ul,
and was not tested experimentally. In practice, simply using 0.5-2 ul 
minipreps (depending on culture volume and on plasmid copy number) 
works fine. In theory, advantages of higher template concentration are: 
i) more product with less number of cycles, ii) higher chance of priming 
when annealing is suboptimal. The theoretical disadvantages of having 
too much of the template are: i) introducing more crap into reaction, 
some of which might be inhibitory (minipreprs are not clean!), ii) increased 
background if the Dpn I treatment is incomplete, iii) increase in the 
misprimed side reactions. There is an optimal balance between all 
of these factors somewhere... Other things being equal, the best practice 
is to have as clean template as possible and that means: 
i) don't be greedy with your minipreps; for high copy number plasmids, 
1 ml culture is usually sufficient, 3 ml is a lot and more than 3 ml 
borders on insanity; ii) use PB solution volume so that it covers 
all areas of the column that were in contact with cell lysate; iii) use PE 
solution volume so that it covers all areas of the column that were in 
contact with PB, and use PE wash TWICE. 

7. Annealing temperature. Pfu Fusion seems to be less finicky in terms 
of annealing conditions. 2-10 degrees below calculated primer TM have 
been found to work in various cases. 55� should work in the vast majority
of cases. 

8. DMSO concentration. DMSO is optional as it rarely makes all or none 
difference. Its main effect on the reaction is two-fold: it decreases melting 
temperature of DNA duplexes and it reduces primers' secondary structure. 
The protocol above uses 4% final. You may also try 0 and 6% as a substitute 
to an annealing temperature variance (make sure to mix DMSO well with 
other components before adding polymerase). 

9. Extension time. Pfu Fusion is quite fast. The default is 60 sec per kbp 
at 65�C and it definitely works. Tried alternative is 30 sec at 68�C and, 
for sure, it also works. However, in "QuikChange cloning" trials we found 
that 65� gives more colonies (perhaps because of the some strand 
displacement activity at 68�). 

10. Other things that may sometimes be beneficial. Increased concentration 
of the primer two-fold, increased concentration of dNTPs two-fold, an 
extra 2 mM MgCl2 in the reaction all, in theory, have an effect of increasing 
product yield with concomitant slight increase in the error rate (not a big 
deal given the error rate of Pfu Fusion). 

11. Just like regular QuickChange, the single stranded QuickChange works 
for two mutations simultaneously but with reduced efficiency. Make primers 
so that they have similar TMs and expect 20-30% of the clones to be double
mutants. 


More information about the Methods mailing list