Clean ligation controls but no insert - (Oct/23/2014 )

Good news.  I think the noise on the vector + ligase plate was actually a mistake on my end - it looks like some insert found its way into this reaction.  Too many late nights apparently took their toll on my focus.

In any case, the cloning appears to have worked like a charm.  Most clones have the insert.  Very efficient.  Only one allele appears to have failed out of the last batch I tested, and it was likely due to low DNA recovery prior to ligation.

Thanks for the tips and troubleshooting advice, Phage434.  I think I'm in good shape now.

Cheers

Good news. If you have insight about what changed, it would be useful to explain it, so others could learn.

Hi Phage434 and community,

I've been running through my notes to compile and check the factors that have made a difference.  I agree, this website is a great resource so I hope somebody else can benefit from my experience in some way:

First, I ran into another problem with my results shortly after the last post - I was PCR positive but RE digest negative on many of my colonies.  This told me I had a transformation and contamination problem with uncut vector persisting.  Several days of troubleshooting later I finally fixed.  Here's what I've learned:

1.  My background was largely due to a bad DpnI digestion.  Check your enzymes if something doesn't make sense.  My DpnI couldn't cut butter, so even tiny amounts of template were still getting through.  Coupled with less-than-ideal PCR backbone product (see #3) this was giving me headaches and represented a major limitation in my strategy.

2.  I've started using a "killer cut" method immediately prior to transformation/after ligation.  I use an enzyme that only cuts undigested vector (in this case, I use a site in a different insert than the one I'm cloning).  This virtually eliminates any carry-over when using plasmid vector as the backbone.  Also helpful as an alternative to DpnI when using a PCR backbone as I have - particularly when DpnI fails and one must wait for the delivery of fresh enzyme.  This works pretty well with .25uL of enzyme added to the ligation mixture for ~10 minutes followed by a brief heat kill (which might be unnecessary).

3.  The purity and homogeneity of the vector backbone acquired from either gel purification or PCR amplification was the single greatest factor in my success and failure.  I scrutinized my PCR product for days, eliminating any trace of nonspecific bands and mispriming.  It turns out that my latest batch of polymerase (Q5) was over-performing, yielding high MW product of an unknown composition.  I dropped the extension time to 20s/kb, increased annealing temp and toyed around with cycle # some more.  Only when I had a post-purification single band on gel that failed to produce colonies when transformed into competent cells did I proceed to ligation.  I compared my PCR backbone to traditionally gel-purified results side-by-side.  Both worked, but the PCR protocol was slightly higher efficiency and much easier.  Oddly, I did still get some background on the vector + ligase plate with the PCR product, but the insert plates were loaded with inserts.  Go figure.

4.  I found spiking my T4 buffer with fresh ATP to be quite helpful.  This increased my transformation efficiency of ligation products on its own.

5.  Wash and wash again when purifying on a spin column.  I don't think 3-4 PE/ethanol washes are excessive.

6.  Less is more.  My ligations worked best and had the highest efficiency when I used 15ng of vector and had almost no difference between 1:1, 2:1 or 3:1 ratios of insert to vector.  Volume of 10uL final using 10x T4 ligase buffer and 1uL ligase (400,000U/mL).  I could still see the ligation products on a gel if I ran 3uL to double check for mobility shifts at various points in the ligation.

When it works, it works well.  I screened some more clones yesterday and had 9/10 screen positive by restriction digest from one transformation (4/4, 6/8, 0/4 in others checked).  Can't complain with those results, some variation is expected.

Cheers and thanks for the feedback.

@miST32 hey your point #4 how exactly does one spike T4 buffer with ATP, I want to try it out

Hi KhaleesiDany,

(Also a fan of ASOIAF, I see!)

I am not very surgical about this, but here's what worked for me when I suspected my old-ish ligation buffer:

- The ligation buffer is supposed to have 1mM ATP, so I rationalize that supplementing to between ~1mM (assuming complete degredation) or ~2mM (assuming no degradation) will be beneficial.  So far I haven't found any problems.

- I prepare 100mM rATP in water, and dilute ~1:100 into an aliquot of ligation buffer (I like to prepare 100 aliquots now to avoid freeze/thaw of the whole tube)

Hope you find it helpful!