Quickchange site directed mutagenesis/PCR problem - Transformation isn't working :( (Feb/28/2007 )
Hi Bioforumers,
I am new to the field of molecular biology and I have been trying to do a QuickChange experiment where I want to create a library of mutants that have been modified at the TGG region. So I ordered the following forward and reverse primers at MWG:
1) Forward primer:
5' CAC CCG CTG AAT TTT AAG GGA AGG NNS CTG CGC GAC CGA CTG 3'
2) Reverse primer:
5' CAG TCG GTC GCG CAG SNN CCT TCC CTT AAA ATT CAG CGG CTG 3'
NOTE: NNS = the point where I want to introduce mutations
OK, so MWG sent me these primers where they said to add 693 ul of dH2O to the forward primer (since it is solid) in order to get a final concentration of 69 pmol/ul. For the reverse primer I added 653 ul of dH2O in order to get a final concentration of 65 pmol/ul.
I was then told by someone to do a 1 in 10 dilution of these primers for the PCR reactions. So I took 10ul of each primer and added 90ul of dH2O. (Maybe this is wrong)
I then followed the Quickchange protocol and made the following PCR reaction:
5ul 10x reaction buffer
1ul DNA template
1ul forward primer
1ul reverse primer
1ul dNTP mix
40ul dH2O
1ul Pfu turbo polymerase
I then put the reaction into the PCR machine set at these conditions
95 C, 30 secs
95 C, 30 secs
55 C, 1 min
68 C, 7 min (for a 6.4 kb plasmid)
Cycled 16 times.
I then added 1ul Dpn1 to remove the parental DNA and transformed into XL1 Blue cells as instructed in the manual.
RESULT: NO COLONIES!!! AAARGH!!! 
I have been repeating this for the past 2 weeks and getting nothing. It's obvious my PCR isn't working!!!
But I can't seem to find the problem....I have tried it using 0.5ul and 2ul of my DNA template but I don't get anything.
Please could anyone see where the problem is....I think it's my primers....maybe I shouldn't do a dilution and just use the stock of 693 and 653ul?
Thanks for any suggestions!
Sara
What about your bacteria? Have you tested the transformation efficiency? With which method do you transform?
I had a little trouble with the Quikchange kit to start off with but have got it working nicely now.
1) Have you managed to get the test amplification to work with the quickchange kit? You should be able to see a faint band with a 5ul aliquot of the reaction both before and after Dpn1 digestion at about 2.7kb.
2) Can you get the transformation control (pUC19) to work? I find it works best when the 14ml BD tubes are immersed pretty deep in the waterbath when you heat shock the bugs
3) If the controls are working then it might be something to do with the plasmid/primers. It is really important to use 125ng of each primer in the reaction - too much or too little messes it all up. Check that you are using 125ng. Also - make sure your primers have been purified by HPLC - the desalting that MWG use only gives 60-70% purity, which isn't good enough for the kit.
4) I use 1.5uL of DMSO (3% final) in my 50ul reaction to denature secondary structures in the primers. although they look like they are designed properly (i check mine here - http://depts.washington.edu/bakerpg/primer...rimertemp.html)
4) One thing I found makes a huge differences is doing the whole thing in one day - i used to leave the reaction overnight and transform the next day - that seems to reduce the efficiency of the transformation drastically.
Hope that helps!
Apparantley XL1 blue cells have a good transformation efficiency. Colony number can be as much as 1000. I am sure my transformation is working because I do get colonies with the control. I just use a standard transformation protocol:
50ul of XL1 Blue cell in prechilled falcon tubes
1ul of Dpn1 digested DNA
Leave on ice for 30 mins
Heat shock for 45 secs at 42 degrees C
Leave on ice for 2 mins
Add 500ul SOC medium
Incubate at 37 degrees C for 1 hour
Pipette onto LB-Amp agar plates
Incubate at 37 degrees C overnight
Should see colonies next day
I'm pretty sure there's nothing wrong with my transformation.....I'm sure it's the PCR
Thanks for your reply
Sara
1) Have you managed to get the test amplification to work with the quickchange kit? You should be able to see a faint band with a 5ul aliquot of the reaction both before and after Dpn1 digestion at about 2.7kb.
2) Can you get the transformation control (pUC19) to work? I find it works best when the 14ml BD tubes are immersed pretty deep in the waterbath when you heat shock the bugs
3) If the controls are working then it might be something to do with the plasmid/primers. It is really important to use 125ng of each primer in the reaction - too much or too little messes it all up. Check that you are using 125ng. Also - make sure your primers have been purified by HPLC - the desalting that MWG use only gives 60-70% purity, which isn't good enough for the kit.
4) I use 1.5uL of DMSO (3% final) in my 50ul reaction to denature secondary structures in the primers. although they look like they are designed properly (i check mine here - http://depts.washington.edu/bakerpg/primer...rimertemp.html)
4) One thing I found makes a huge differences is doing the whole thing in one day - i used to leave the reaction overnight and transform the next day - that seems to reduce the efficiency of the transformation drastically.
Hope that helps!
Hi Adrian,
Thanks for your reply.
1) I do see a band before digestion but nothing after digestion. However the band seems to be at 6kb (looks like it's the whole plasmid)
2) Yes, the control does work and I do use 14ml Falcon tubes.
3) Now this will seem like a daft question - Would 125ng be 1.25ul volume of primer? I always use 1ul. (I'm not familiar with the units in biology as I am a bioinformatician by profession). How would I know I have 125ng?
4) I actually did the whole experiment in one day last week but didn't get anything (as usual)
5) Is my primer dilution wrong do you think?
6) Is it possible for you to post a step by step protocol on how you did your Quickchange (including how you calculated concentrations and volumes)? I feel like I'm going through a crisis on this PhD (and my supervisor doesn't see too happy with me
). I would really appreciate your help on this.Thank you for your suggestions
Sara
Do 40 cycle PCR and put on gel. See if you get the desired result, if not: optimise your PCR. Afterwards, do the number of cycles from the quickchange protocol and proceed.
Hi Sara,
If your controls are working then it sounds like a problem with the template or primers.
1) Primers
a) They must be HPLC purified (i've tried MWGs HPSF primers and it didn't work)
When you get them, spin them in a microfuge at top speed for 1 min and then thoroughly dissolve them in dH2O (pipette up and down lots, especially around the side of the tubes)
c) MWG should give you a data sheet. This will tell you the mass and the molecular weight of the primers you have been sent. Using that you can work out the dilution so that 1ul of primer will equal 125ng. (I tend to dilute my primers to 100pmol/ul stock and normally it's about a 1:7-1:13 dilution to for use in the reaction). At a glance, your dilutions don't look too far off the mark but perhaps they are a little low - might be worth trying 1.5uL of your 6.5pmol/ul dilution - this will be about 125ng (at a glance your primers are of molecular weight around 13000).
d) Your primers are fairly GC rich (60%) so it might be worth adding some 100% DMSO to the reaction - 1.5uL in the final reaction mix to give 3% final volume. This should help prevent any secondary structures.
e) Your primers are slightly unbalanced, 23bp on one side of the mutation, 15bp on the other - I don't know if that will make a difference when the primers anneal (i.e. it will be a bit lop sided) but I've always had my primers symmetrical on either side of the mutant region
2) Template
a) I've always done my reaction with a tube with 50ng and one with 25ng of template. Check your adding the right amount of template to the reaction. You should be able to quantify your template using a spectrophotometer to give a reading and then you can dilute your template down to 50ng/ul and add a 1ul or 1:2 for 25ng. I read somewhere that one of the invitrogen vectors (i think it was pcDNA3.1) causes the kit some problems but stratagene have a protocol to deal with it, so if you're using that vector, it might be worth giving them a call.
c) Check the resistance gene in your plasmid - is it ampicillin - that would be an obvious reason why you dont see any colonies on an amp plate.
Hope that helps
(My protocol for quickchange is pretty much as it says in the manual: two tubes, one with 50ng template, one with 25ng template. Each tube is made up as the manual says, but I make the reaction mixture up so it is 3% DMSO (1.5ul) in the final mix. I run 18 cycles of PCR - an initial denaturation of 2:30 at 95oC to really separate the dsDNA and then cycling for 1min at 95oC, 1min at 55oC and 2min/kB at 68oC. Digest with 1.5uL Dpn1 (just to make sure you get rid of all the template) for 1:15mins at 37oC. Then prechill the BD tubes, 3uL of the digested reaction onto the bugs and swirl them for a bit. ice for 30mins. heat shock at 42oC for 45s immersing the tubes deeply. Then ice for 2mins. Add 500uL of NZY Broth (I notice your using SOC medium - I tried LB once at this stage and it didnt work as well, so maybe NZY might be worth investigating - its a commercially available powder) Then 37oC with 220rpm shaking for 1 hr and then plate them for about 18-20 hours.)
Hi Adrian,
Thanks for getting back to me. I will definitely try your suggestions. I just feel like this experiment is going nowhere though I guess this is a PhD for you! 
Thank you once again.....you've been a great help.... will let you know how it goes (Fingers crossed!)
Sara
Hi Sarah
ever tried longer elongation time ..let's say 14min...with me worked .
NOW my problem is the transformation:)