Clean ligation controls but no insert - (Oct/23/2014 )
I've been working to clone several mutant alleles of the same gene into a new expression vector but have encountered a new problem (new to me, at least). I have clean ligation controls and negligable background from the vector - However, despite getting between 100-200 colonies on my ligation plates and 4-8 on my vector + ligase plates, only one of my alleles has yielded a successful insert (verified by PCR and digest so far). Pretty frustrating, but also fascinating - not sure what's going wrong.
I've designed the cloning strategy as follows:
Two REs: KpnI (5' of insert) and XbaI (3' end of insert)
1. PCR amplify the vector (i.e. linearize it with primers containing RE sites and extra 5' bases, "in reverse" leaving RE+5'bases on the ends of the PCR product)
- No problem -
2. PCR amplify the inserts with primers containing RE site and extra 5' bases.
- No problem -
3. Check products on gel for size and purity (Vector is ~5100bp, Insert is ~3400, both are clean PCRs)
- No problem -
4. DpnI digest the template away for 1hr. (Might omit this in future attempts, but it has reduced the background).
5. Purify PCR products with GuHCl and silica membrane spin column (QiaQuick system). I do an extra wash with 35% GuHCl after buffer PB to remove potential leftover dimers and elute into warm EB. Usually my concentration is 45-60ng/uL in 25uL elutions.
- No clear problem -
6. Digest insert and vector independently with my two RE's for 2h at 37C in appropriate buffer (NEB 2.1) with 1X BSA.
- No problem, digests are complete in control experiments -
7. Ligate at 1:1 and 1:3 (vector:insert) for 2h at 25C in 10uL. I use 50ng of vector and the aforementioned molar ratios of the insert.
8. Transform full 10uL ligation into DH10b and grow overnight.
So far I've screened nearly 20 colonies from each of my dozen or so transformations across each of my alleles via PCR or digestion (or both). As I mentioned, only one colony has yielded the correct insert!
Usually I find that 10 colonies are more than sufficient to identify a correct insert with this method. This protocol does work as indicated by the one positive clone, but the efficiency is apparently quite low - not good when I have multiple variants to clone.
In my digest screening, I've noticed several uncut plasmid profiles migrating at different supercoiled/nicked sizes. But again, they are not cut. Both of my restriction sites are within the insert for this screen, indicating that nothing looking like the insert has been cloned into these pieces of DNA. My first thought is that perhaps I've cloned my primers into the vector? I am trying to avoid gel-purification since we only have UV in the lab, and I don't want to introduce mutations or further reduce efficiency. Perhaps I should do it anyway?
Mainly, should I just play the numbers game and keep screening, or change the protocol significantly?
Thanks in advance!
You didn't mention a heat kill following your RE digestion. It is essential, to remove enzymes which, if still active, will cut your ligation product.
You can simplify your protocol by adding the DpnI digest to your other restriction enzyme digestions of the vector. 50 ng of vector is probably too much. Optimal for a 10 ul reaction is closer to 20 ng, which reduces concatamer formation relative to closed circular DNA.
You can likely reduce the template amount in your vector PCR to reduce background. You only need vanishingly small amounts as a template.
For your column cleanup, the real problems occur by inadequate washing of the column (do an extra PE wash). Then make very sure all of the ethanol is spun out of the column before eluting. Have you checked your DNA on a gel after cleanup?
Thank you for the quick reply.
I will start the procedure over again today with your suggestions. KpnI is not heat-inactivatable, but XbaI is. I had not inactivated either, but this hasn't caused problems in the past. I will give it a shot. I hadn't thought that 50ng would be too much vector, so thanks for the tip there - I will try it later today.
To answer your question, I checked the purification last night on a 1% gel. The purified DNA runs clean for the insert, though there was a bit of high MW haze in the vector. Surprised by this because there isn't any such haze imediately following PCR. I presume this is some form of nonspecific product that is being enriched.
You could be seeing restriction enzymes bound to the DNA, effectively lengthening the molecules. You can clean up your digestions on a column, even if the enzymes cannot be heat killed. Old school but very effective would be a phenol/chloroform cleanup and precipitation. This is why I choose enzymes that can be heat killed.
I also took a closer look at my PCR inserts before doing the whole cloning operation. Using a higher % gel, I found an odd smaller band I hadn't noticed in several of my reactions. Somewhat faint and very low on the.8% gels after a long run-out, but there.
Perhaps not the main culprit, but it could be a problem in my cloning. I've managed to eliminate it in most of my reactions with a tweaked hot-start and touchdown protocol. I'd consider new primers, perhaps, but this may be about the best I can do (Limited sequence "real estate" to work with due to repetitive sequences).
I read some other posts about PCR-cloning and I'm intrigued by the possibility of simply mixing my vector and insert products together prior to digest as you've suggested elsewhere in this forum. Any specific tips for this approach (amounts of PCR product to mix, primarily)? Seems very simple in principle.
Assuming clean PCR products (no/limited nonspecific products), I will try the following streamlined approach:
1. PCR amplify insert and vector.
2. Check on gel for product size/purity.
3. Purify on column.
4. Mix products and digest (REs + DpnI as suggested)
5. Purify to eliminate REs (Phenol/Chloroform EtOH ppt, or column depending on yield from 3).
6. Ligate and transform.
That all sounds very good. You can test the cutting and ligation efficiency of your PCR products by cutting with each enzyme independently. Then you ligate, and observe the double length fragment. If you don't see it, you know you have a problem. Do this for each part and for each enzyme. It's hard to fail at a ligation if all of these work. Could your T4 ligase buffer be off? You could add a microliter of ATP and see if that made a difference.
Ahh yes - I like the ligation test you've suggested. I'll play around in the lab tonight and see what I can learn about my insert digestions using this technique.
If all looks good, I'll transform some cells tonight with the ligations and know tomorrow. Thanks again for the suggestions, this gives me some confidence.
My ligase and REs check out with the single-restriction/ligation test. One less confound. Looks like I might need to do longer ligations or supplement with ATP though, because the 2X band wasn't very bright. *edit - Or the digest needs to go longer, perhaps?*
Unfortunately, my vector + ligase control plate had lots of colonies this time, indicating failed DpnI digestion. I will repeat tomorrow, doing the DpnI digest independently immediately following vector amplification, and also reduce the template concentration 100-fold. I'm not sure how I'm getting so much carry over when I was already using less than 1ng of template.
Since I'm amplifying my linearized vector segment from a plasmid containing an unrelated insert (insert "A"), I designed a multiplexed colony PCR that can discriminate between the parental vector + Insert A, and a vector containing my insert of interest (insert "B"). Before screening colonies, I ran the PCR on each of the ligation reactions themselves, and found that the dominant product corresponded to uncut vector + Insert A, confirming what I saw on the plates. The vector + insert B product is also there, but it is a minor product, and I don't think screening hundreds of colonies for each of 14 inserts is a good plan - it's what I've been trying to avoid. Seems like I can't win.
Trying again with the cleaner vector product tomorrow. Hopefully almost out of the woods.
Another possibility that I failed to mention is that your vector + insert is toxic. It won't matter how well your ligations work if the product cannot be transformed.
True. Pehaps a lower/slower growth will help in that case. I do notice a size distribution among colonies - might be consistent with toxic plasmids being transformed. Going to note that and see if the small vs. large colonies contain consistently different digest or PCR peoducts. Could be a simple way to screen if it holds up.
Not giving up until the control plate is clean again, in any case.