What else are my experiments telling me?

The previous post discussed what I can call Problem area #1: the evidence that my plasmid prep results have been unreliable - that the absence of plasmid in the prep didn't mean that the cells I started with didn't contain plasmid. So now I need to go back through the other experiments, to check if my conclusions are still solid.

Problem area #2:  Can the specR PCR fragment be ligated into a blunt-cut plasmid, when phosphorylation isn't needed?  Answer: NO.

My first experiment said 'No', but it was flawed by using too high a ratio of plasmid to insert.  So I repeated it using much more specR fragment than plasmid.  This time the results were a cleaner 'No'. Religation of the cut plasmid gave 437 AmpR colonies for 510 µl transformation mix.  Ligation of the same amount of plasmid in the presence of the specR fragment gave only 29 colonies from 150 µl of the transformation mix, and no SpecR colonies (from 150 µl) or AmpR SpecR colonies (from 150 µl).  A positive control transformation using a plasmid that carries both AmpR and SpecR gave several hundred colonies of each from the same volumes.

So I conclude that the specR PCR fragment cannot be ligated, and that it also interferes with ligation of the blunt-end plasmid.  One way to interpret this is that one end of the specR fragment is OK and the other is unligatable.  I'm discouraged but not surprised by this result, because the specR PCR product always behaves oddly in agarose gels: a bit blurry before cleanup and worse after cleanup.

Solutions: (1) I could design new specR primers and try again.  We might even have other specR primers for this cassette. (2) I could try amplifying with the original primers from a different template.  The undergrad used a plasmid I think.  I should have a plasmid with specR in a long-enough segment to encompass my primer sites..  (3) I could cut my existing specR PCR-product with NheI to generate sticky ends.  This is what the undergrad had originally planned.  I would need to design new primers for the inverse-PCR step that generates the rest of the desired plasmid.  Or blunt-cut specR fragment out of a plasmid and ligate this to the inverse-PCR product.


Problem area #3:  Are the kinase (phosphorylation) reactions working?  Answer: YES.

I originally tried to test this by phosphorylating the inverse-PCR product and self-ligating it, transforming E. coli and selection for AmpR.  I got no transformants, so either the kinase reaction failed or there was another problem.  The ligation control, transformation control and plate-selection controls all worked fine.  This was when I discovered I'd been using very old kinase, but repeating the experiment with new kinase gave the same result.  Was the ATP stock bad?  No, repeating the kinase reactions using ligase buffer (contains its own ATP) gave the same result.

A better test of the kinase function comes from the grad student, who has been using it to label chromosomal DNA with 32P from 32P-ATP.  He's getting modestly successful incorporation, suggesting that this reaction is working OK.


Problem area #4:  Is the inverse-PCR product's intact toxin gene toxic to E. coli?  Answer: Weak No (no evidence that it is).

I tested this by making a different inverse-PCR product using the undergrad's old primers.  These cut off the last 5 amino acids of the toxin gene.  She was able to get successful ligation of this product to her specR PCR product, after kinasing a mixture of both fragments.  I ligated this with my kinased SpecR fragment.  This produced one AmpR SpecS colony and one AmpS SpecR colony.  The negative control self-ligation of the inverse-PCR fragment alone (not kinased) gave a few AmpR colonies.  If these do result from self-ligation (I didn't do a plasmid prep on them).

I had only kinased the specR fragment, because I didn't want the inverse-PCR fragment to be able to self-ligate.  In retrospect, especially now that I know that the specR fragment is unligatable, I should have also kinased the inverse-PCR product as a better control to show that the kinase was working.  


THE BIG QUESTION: SHOULD I KEEP TRYING?

This project shouldn't have been such a big deal.  But it's the bottleneck in getting the toxin-antitoxin work finished and published.

One possible plan:  Buy some NheI, cut the (blurry) specR PCR fragment and run it in a gel.  Do I get a nice sharp fragment of the right size?  If yes, design and order new inverse-PCR primers with NheI sites.  (Why did the undergrad choose NheI?  I have no idea?)  Cut the new inverse-PCR product with NheI and ligate it with the specR fragment.  Promers are cheap, so I oculd instead jsut design new specR and inverse-PCR primers with matching sites.

Another plan:  I recently found a note I made at last summer's Gordon Conference, reminding me that a favourite colleague had offered to make this damned mutation for me.  But I don't like to ask this of him...

What next in the antitoxin knockout endeavour?

OK, time to end my two three? four? weeks of sulking because my experiments won't work...

Where was I?  (... must consult my notebook)

I was trying to resolve two issues.  The first was why ligation of my phosphorylated PCR fragments did not produce a plasmid that could transform E. coli to AmpR SpcR.  The second was why my plasmid preps were not producing any plasmid, from cells that I was quite sure contained a high-copy-number plasmid.

This second issue was compromising my ability to investigate the first issue, so let's deal with it first.

I was doing my plasmid preps using Econo-spin spin columns (from Epoch Life Sciences) and column reagents we had made up ourselves.  This is much cheaper than using spin-column kits from Qiagen or Sigma.  But the columns and reagents were quite old, and I was not even sure that I was using the right volumes of the reagents.

Here's what I was doing:

The basic procedure is to start with a version of the standard alkaline-lysis procedure:  

1. Pellet cells and resuspend in a neutral buffer containing EDTA
2. Add 0.2M NaOH +1% SDS.  The SDS will lyse the cells and the high pH will cause the base-paired DNA strands to separate, 'denaturing' both chromosomal DNA and the plasmids.  Each plasmid's two DNA strands will be looped together because they are interlocked circular molecules.
3. Neutralize the NaOH and make the SDS insoluble by adding a low pH potassium acetate solution.  The two plasmid strands will regain their base pairing returning the plasmid to its normal configuration.
4. Centrifuge the tube to pellet the cell debris, including the chromosomal DNA and most of the SDS.  The plasmid DNA and various soluble components remain in the supernatant

In the old-fashioned procedure the supernatant is extracted with phenol and then chloroform, the DNA (and RNA) is precipitated with ethanol, and the pellet is resuspended in TE buffer (often with RNase A added to degrade all the RNA).

With a spin column the supernatant is instead placed in the top part of the column (see figure above, from Perkin-Elmer), and spun through it.  The DNA sticks to the filter membrane in the base of the column, and all the unwanted soluble material washes through and is discarded. The DNA stuck on the membrane is further cleaned with one or two washes of a special solution containing ethanol, and then the DNA is eluted from the membrane into a clean tube using a small volume of TE buffer or water (usually 50 µl).

The kits typically are expensive (Qiagen: $1.35 per column). They include all the reagents, but the reagent recipes are kept secret.  The same columns work for a number of different DNA-purification procedures, but you need to buy a different kit for each to get the specific reagents used in each procedure.

Epoch and other budget suppliers will sell you just the columns (Epoch, about $0.40 each) and provide recipes so you can make your own solutions for all the procedures.  But as I said, our Epoch columns were old (several years?) and I wasn't confident that my hand-written notes specified the correct volumes of reagents to use.  We had the sheet with the reagent recipes but I couldn't find one with the protocols.

Anyway, I did a test.  With replicate samples I used our homemade reagents to do the alkaline lysis steps 1-4 above, and then finished some sample with a spin-column cleanup and some with phenol extraction and ethanol precipitation.  The column samples had some chromosomal DNA contamination but no plasmid and no RNA.  The phenol-etc samples had no chromosomal DNA and no plasmid but a lot of RNA.  Bummer.

So I did another test, this time adding to the two treatments above samples where I did the alkaline lysis steps using reagents specified by an old non-column protocol, followed by the old-fashioned phenol extraction and ethanol precipitation steps.  This time I did get some plasmid from the old-style prep, but again none from either prep using the column reagents.

So this suggests that the column reagents or volumes are at fault.  I checked online, but found a slightly different set of protocols, with some reagent names we didn't have the recipes for.  I contacted Epoch and they kindly sent me a new recipe sheet.

Epoch also provided this advice about column storage and regeneration:

If you have access to cold room you can store the column in a cold room. Or if you still have room in your refrigerator which does not go through defrost automatically you can also store your column there. We can also ship you some "moisture pack", most conveniently with your next order, which can be sealed in the plastic bag. Finally you can apply 20 ul of 0.2N NaOH to the old column 30 minutes before you do miniprep. This will regenerate the membrane to the maximum bidding capacity.  

So...  I suspect my preps didn't give plasmid because I was using the wrong volumes of the column alkaline-lysis reagents volumes, not because the cells didn't contain plasmids.

How does this affect my thinking about the rest of the work?  Might some of the colonies I discarded have contained a desired plasmid?  I'll put that in the next post.