Field of Science

Maybe the toxin is toxic!

Recap of the last few posts:  Starting with a plasmid containing the toxin-antitoxin operon of Actinobacillus pleuropneumoniae, I've been trying to create a derivative plasmid whose toxin gene is functional but whose antitoxin gene has been replaced by a specR cassette.  This involves several steps: PCR of the specR cassette and inverse-PCR of the plasmid (to produce a fragment lacking the antitoxin gene), phosphorylation of the specR fragment with T4 kinase, ligation of the two fragments, transformation into E. coli, and selection for SpecR and AmpR cells.  The PCR steps work but the rest fails to produce any resistant colonies

After the first attempt failed I introduced several controls: EcoRI-cut pUC18 as a ligation control, another SpecR AmpR plasmid as a transformation control, and the kinase-treated inverse-PCR fragment as a kinase control.  The ligation and transformation controls worked, so I decided the kinase was at fault.  This hypothesis was supported by finding that I had been using a long-expired stock of kinase rather than the one bought earlier this year.

But two more attempts using the new kinase have also failed.  The first used the supplied kinase buffer and my stock of ATP, and the second used new ligation buffer (recommended by the supplier) which contains its own ATP.  The second time I also preheated and rapid-chilled the substrates, which is recommended to help expose the blunt ends to the kinase.  Both times I got no transformants from either the test or the kinase control, but got lots with my transformation control plasmid. (I didn't bother repeating the ligation control.)

I've been trying to think of what else could be going wrong with the kinase reaction, but it just occurred to me that maybe there's a completely different problem - maybe this toxin is toxic to E. coli.

Although the honours student had mentioned this concern when she handed this project over to me, I had discounted it because the H. influenzae homolog is not toxic in either H. influenzae or E. coli. But both my desired construct and the recircularized inverse-PCR fragment I'm using as a control are expected to express the toxin, possibly at high levels.  So maybe my reactions are all working, but the plasmid they produce is not tolerated by E. coli.

How to test this?  Directly testing for lethality is tricky.  But I can do a different kinase control using a different inverse-PCR fragment, one that won't express the toxin. If the problem is the toxin, this should give AmpR transformants.  I can also use the same fragment with the spec cassette, and I should now get SpecR AmpR transformants.  I have all the honours student's primers and her CR conditions so this should be straightforward.

Think Check Submit - can't we do better than this?

The Scholarly Kitchen blog has a post about a new initiative from a large consortium of scholarly publishing societies and individual publishers, intended to help inexperienced researchers avoid journals from 'predatory publishers'.  This is a very worthwhile goal, but the actual advice provided so far isn't going to exclude most of the bad guys.   

The first step just explains why researchers need to be careful where we publish:

The third step is just reassurance:

The second step is the one that matters; it tells researchers what they should look for:

There's nothing wrong with this advice, but it's certainly not treading on any publishers' toes.

Most importantly, there's no mention of the most valuable resource we have, Beale's List.  This is a frighteningly long list of open-access scholarly publishers whose tactics are potentially, possibly or probably predatory.  It's maintained by Jeffrey Beale, a librarian at the University of Colorado, at his Scholarly Open Access blog.  The last time I checked, a couple of years ago, there were about 300 publishers on this list, but today there are 882!  And this is just the publishers - most of these have multiple journals.

Beale's list isn't just a list.  Beale also provides explicit sets of criteria for evaluating individual publishers and journals.  The absence of Beale's List from the THINK CHECK SUBMIT campaign isn't really surprising, but it reinforces my concern that we can't rely on the publishers to look out for researchers' interests.

New kinase stock found

Along with new stocks of other enzymes.

Somebody apparently thought it was a good idea to stop putting enzyme stocks in their usual place (the 'Special Enzymes' box) in the -20 °C freezer, and instead put them in this new 'coloning' box.

They then put the new box in a bin in the freezer where its label couldn't be seen.

It's the kinase!

Yesterday's experiment worked very well, in that the thorough controls clearly tell me where the problem is.  But the actual experiment produced only three candidate colonies.

Control E: No DNA. No SpecR or AmpR colonies.  GOOD-selective plates kill non-resistant cells

Control F: AmpR SpecR Plasmid.  p∆TA::Spec: ~350 AmpR and ~350 SpecR transformants   GOOD-selective plates select for cells carrying the resistance genes on a plasmid, and the competent cells transformed efficiently.

Controls D and G: EcoRI-cut pUC18 ± ligase.  ~12,500 AmpR colonies after ligation, only about 250 without ligase.  GOOD- ligation worked.

Control C: No-ligation control. Kinased spec PCR product plus not-kinased inverse-PCR product, ligation reaction with no ligase.  No SpecR or AmpR colonies.  GOOD-The fragments do not spontaneously circularize and transform cells, and the fragment mixtures do not contain any unwanted intact plasmid.

Control B: Kinase control.  Ligation of kinased inverse-PCR fragment.  Should have given AmpR colonies, but none.  BAD- Kinase failure.

Experiment: Ligation of kinased spec PCR product plus not-kinased inverse-PCR product.  Gave only 1 AmpR colony and two SpecR colonies.

Next steps:  

I've streaked the three candidate colonies on both Spec and Amp plates.  The desired plasmid should confer resistance to both.

And I looked at the expiration date on the tube of BioLabs T4 polynucleotide kinase I've been using. 03/09!!!!  AAARRRGGGHH!!!!

Have I been using the wrong tube of kinase?  Is this not the kinase that the former undergrad and sabbatical visitor used successfully last year?  Searching the 'Special Enzymes' freezer box turned up another tube of T4 polynucleotide kinase, but this one looks even older.  So I've just emailed the undergrad and sabbatical visitor to ask what they used.

Positive control problem solved

I did the test experiment described in the previous post, and then spent the past few days figuring out why my positive control transformation didn't work any more.

The test experiment was to kinase, ligate and transform into DH5alpha the product of the inverse-PCR reaction.  If the T4 polynucleotide kinase reaction worked, its blunt ends would acquire 5' phosphates that would allow it to be circularized by T4 DNA ligase, and to then transform DH5alpha to ampicillin resistance.  The negative control was no DNA and the positive control was the same p∆TA:spec plasmid that had given thousands of AmpR and SpecR transformants in the previous experiment.

Sounds great, but this time the positive control didn't give any transformants at all!  Background small colonies were frequent, possibly because the plates were a bit old and the ampicillin had lost its potency, so I didn't trust the few larger colonies on the inverse-PCR reactions plates.  I streaked a few of the large colonies to check if they were genuinely AmpR - one was.

I repeated the control transformation and negative control with new Amp plates; the no-DNA control plates were clean but so were the p∆TA:spec plates.

I thought the problem might be the plasmid, but I wasn't sure I have another reliable positive control. So I did a miniprep from the one genuine AmpR colony I had streaked and transformed the cells with that DNA.  I also had the usual no-DNA control, the undergrad's p∆TA:spec, another plasmid made by the undergrad (used successfully by me as the inverse-PCR template, and some pUC18 left by a sabbatical visitor.

Success all around.  The miniprep DNA, the other undergrad plasmid and the pUC18 all gave lots of transformants (the photo shows part of a pUC18 plate), and the undergrad's p∆TA:spec and the no-DNA control gave none.  I don't know why the p∆TA:spec plasmid worked well in my first experiment - maybe I had grabbed a 'wrong' (i.e. good DNA)tube.

Next step, repeating my original experiment (the one in the previous post), this time with better controls.

  1. DNA clean-up: I did a new inverse-PCR reaction because the old one got used up in the tests.  I need to start by doing a spin-column cleanup of it.
  2. Two kinase reactions: (i) the 'blurry' spec PCR product and (ii) the not-blurry inverse-PCR product.  Heat-inactivate the kinase before step 3 (65°C 20 min).  This time I'll use a newer stock of ATP, and the official kinase buffer.
  3. Four ligase reactions: A. The kinased spec fragment plus not-kinased inverse-PCR fragment. B.  (kinase control) The kinased inverse-PCR fragment. C. (negative control) The not-kinased spec fragment plus the not-kinased inverse-PCR fragment . D. (positive control) pUC18 cut with EcoRI and heat-inactivated (65°C 20 min).
  4. Six transformations: Ligations A, B, C and D, plus 1 ┬Ál pUC18 as positive control and no DNA as negative control.
Preparations:  We have enough kinase, and I've just sent the grad student to buy more ligase.  Luckily I have lots of frozen competent cells for the transformations.  I'll need to digest the pUC18 and check it in a gel, and pour lots of Amp plates and some Spec plates.


Yesterday I kinased my SpecR PCR fragment, ligated it to the inverse-PCR fragment and transformed this into E. coli DH5alpha.   But my transformations gave only the same tiny colonies as the negative controls (no-DNA and no-ligation).  One of the positive controls (another AmpR SpecR plasmid made by the undergrad) gave thousands of AmpR and SpecR transformants, so I know that the competent cells (RbCl2-competent, frozen many years ago) and the antibiotic plates are fine.

(The other positive control was her pGEM-Spec construct; this gave no transformants for either Amp or Spec.  Since this prep didn't work as a PCR template either I should now throw it out.)

I wasn't totally surprised, because I already suspected that there was something wrong with the PCR amplification of the spec cassette.  The PCR product looked right when I ran an aliquot of the PCR reaction in a gel, with a sharp band just smaller than 2 kb.  But there was also a faint smear of smaller DNAs below this band, and after I had used a spin column to clean up the pCR reaction (should remove primers, enzyme and salts) the sharp band was smaller and blurry, and the DNA smear had become as second blurry band. The column cleanup of the inverse-PCR fragment gave a nice sharp band of the right size, so I don't think the problem was with the column treatment.

This was cause for concern, so I repeated the spec PCR, thinking that maybe I had screwed up (why would a cleaner band be smaller and blurrier?).  But I got the same result.  I again used the student's primers and template chromosomal DNA, but I lowered the annealing temperature for the first two cycles because the 5' 12 bp of each primer are not complementary to the template DNA. Before cleanup the amplification product looked the same as before (red arrow in left gel) - a sharp band with a faint smear below it (the smear doesn't show up in the photo).  After cleanup it again turned into a slightly smaller blurry band with a second blurry band below it (two red arrows in the right gel).  The larger sharp band indicated by the white arrow is the inverse-PCR fragment after identical cleanup.

I have no idea what could cause the SpecR PCR fragment to behave like this.  Before I dig into it I should check that the cloning failure was not instead caused by a failure of the kinase or ligation reactions.

Test for kinase and ligation:  I'm taking the inverse-PCR fragment, and kinasing and ligating it.   This should produce a simple plasmid with the antitoxin gene deleted.  I'll transform this into DH5alpha, selecting for AmpR. (Again no-DNA control and p∆AT:spec control.)  If I get lots of transformants then the problem is the Spec PCR fragment.  If I get none then the problem is the kinase or ligase reaction.  I was hoping to use singly-cut p∆AT:spec as a ligation control, but neither SacII nor HincII appear to cut where they should, so I'm doing without this control.

Just doing it

My inverse PCR reaction using the Q5 High Fidelity polymerase (creates blunt ends) worked on the first try!  I now have the antitoxin-deletion fragment I can ligate to a SpecR cassette to create the antitoxin knockout plasmid. Running 1/10 of the reaction in a gel gave a reasonable band, so I think I have enough DNA for the next steps.

I was initially planning to cut the SpecR cassette out of a plasmid the Honours student made, but we didn't have any of the blunt-cutter (BstZ1) this would need.  Since money is tight, I decided to instead do what the Honours student had done, using her primers to amplify the fragment with the Q5 High Fidelity polymerase, adding 5' phosphates with T4 polynucleotide kinase and ATP, and then blunt-end ligating this with the inverse PCR fragment.

The SpecR PCR didn't work the first time, using the Honours student's plasmid as template.  The plasmid DNA didn't look very good in a gel, so after consulting with the Honours student by email I tried using some chromosomal DNA she had made from another knockout mutant.  This worked very well.  My only concern is that in a gel the SpecR fragment had a short smear of what looked like shorter DNA below it (sorry, forgot to save photo image).

Next step will be to clean up both PCR products.  The Honours student did this by gel purification, using a kit favoured by the sabbatical visitor, but I think I'll just use our usual spin columns.  The grad student warns me that the recovery won't be great, but I think I have DNA to spare.

Next step will be to phosphorylate the 5' ends of the SpecR fragment.  This procedure looks very straightforward.  I could also treat the inverse PCR fragment, but this would allow that fragment to ligate to itself, creating many side reactions that I don't want.

Then maybe another spin-column step, because I think I should remove the kinase so it doesn't phosphorylate the other fragment. But I could instead just heat-inactivate the kinase (20 mi at 65°C, says NEB), since the kinase and ligase enzymes have nearly identical reaction buffers, and both use ATP.  After heat-inactivation I'd just need to add the other fragment, the ligase, and maybe a bit more ATP.

Then transform into competent E. coli (IDH5-alpha? I'm pretty sure I have lots in the freezer) and plate on spectinomycin.  The plasmid I want should give resistance to ampicillin too.

What about controls?  (I'll only do the easy ones this time.  If the transformation fails I'll do more controls.)
  • Negative control: Mix of fragments before addition of ligase.  
  • Negative control for transformation: No DNA.  
  • Positive control for ligation:  I'd need to digest a plasmid.
  • Positive control for kinase:  Do the ligation reaction with non-phosphorylated DNA
  • Positive control for transformation: One of the Honours student's successful plasmids.

Fear of cloning: time to stop stalling and just do it!

For several months I've been stalling on a relatively simple project whose completion would let us submit a nice paper.  This is the missing step in an undergraduate Honours student's project; I wrote about her project and what I need to do here.

  1. Starting with a circular plasmid containing the short 'Toxin' and 'Antitoxin' genes, I'll use inverse PCR to amplify a linear fragment that lacks most of the coding sequences of the Antitoxin gene.
  2. I'll also amplify (or find) a SpcR cassette.
  3. I'll ligate these two molecules together to create a circular plasmid with the SpcR cassette replacing the Antitoxin gene.  I could do this by blunt-end ligation (what the Honours student originally did) or do it as the student originally planned, using conventional ligation of 'sticky ends' generated by digesting both fragments with a restriction enzyme whose site is present at all the ends (she designed it into the primers).  I think she changed her plan because our stock of this enzyme was inactive.
  4. I'll use this plasmid (linearized by cutting somewhere in the vector) to transform the bacterium Actinobacillus pleuropneumoniae to SpcR.
  5. I'll use PCR of chromosomal DNA from the new resistant transformant to check that the original antitoxin gene has been replaced by the SpcR cassette.
  6. Next I'll do a transformation assay on this mutant to see if its competence has changed, with the wildtype and toxin mutant as positive controls.

Step 1 actually requires that I get off my butt and:

  1. Resuspend the primers at the appropriate concentration of TE (or water?).  (Check with the grad student.)
  2. Find the PCR reagents. (Ask the grad student).
  3. Find the template.  (Find info provided by Honours student before she left.)
  4. Learn how to run the PCR machine (Ask the grad student.)

What am I doing? What should I be doing?

Here's an attempt to get better focused and organized:  (Trigger alert: most of this post is not about the real science but about the tiresome details that get in the way.)

What I've been doing:

1. Preparing for a big mutagenesis project:

I want to isolate a lot more hypercompetence mutants.  I have a plan, and I've pre-tested the mutagenesis step, but a key component is a way to efficiently transform rare competent cells in a large population of non-competent cells.  The best way to do this is with a cloned or PCR'd DNA fragment containing a selectable point mutation giving antibiotic resistance.  In the past I've used a cloned novobiocin-resistance fragment.  I had thought that a PCR'd fragment would work just as well, but tests show it to be only twice as good as chromosomal DNA (it should be at least 10-20 times better).

So I've been struggling to get good plasmid preps.  I have two plasmids carrying this 9.4 kb novR fragment.  One is the original clone, in an old natural ampicillin-resistant plasmid called pRSF0885.  I have very old plasmid DNA preps, and some frozen cells from 1990 with the plasmid.  The cells are still viable and resistant to ampicillin but my plasmid minipreps give no plasmid DNA.  The old plasmid DNA efficiently transforms cells to AmpR, using a method that should produce plasmid transformants rather than replacement of the chromosomal allele, but again my plasmid preps give no or almost no plasmid DNA.  I also have the same novR insert cloned into the CmR vector pSU2718 (almost identical to pSU20), but the plasmid yield is miniscule.  So I'm beginning to suspect that something may be wrong with my plasmid prep method - I need to streak out something foolproof and prep it.

2. Tracking down the cause of non-reproducible colony counts:

Counting colonies is our most important research technique (yes, in some ways we're a very low tech lab).  Normally the results are nicely reproducible, but lately I and the PhD student have been seeing bizarre discrepancies between replicate plates and between different volumes or dilutions of the same culture.  After eliminating other variables (plates too dry? plates too wet? new brand of plates (nasty VWR plates with rounded edges?)? dirty spreading beads?) I did a test of 8 different combinations of different sources of BHI medium and of agar.  This suggested that the problem may be with the old BHI agar we've been using (we were given two big buckets of it by a colleague).  The good news is that this BHI agar is almost all gone, so the problem will solve itself.  The bad news is that this BHI agar is almost gone, so we're going to have to start paying for more!  (And I'm not confident that the problem is completely solved - there are still discrepancies, and tracking down the cause of irreproducibility is inevitably an exercise in frustration.)

What I should be doing:

3. Finishing up the honours student's Actinobacillus pleuropneumoniae experiment:

One of last year's honours undergrads left us with an almost-ready-to-submit manuscript lacking only some RNA-seq analysis and a remake and retest of one mutant.  The other honours undergrad (now our summer student) has been working on the RNA-seq analysis, but we're still waiting for the final missing sequencing data.  My job is to remake the mutant and test its competence phenotype.

The problem with her original mutant was that misannotation of a gene overlap had resulted in a mutant that deleted the last five amino acids of a gene that needed to be intact.  Last month I spent ages designing primers to delete the correct segment, but the primers ahve jsut been sitting on my bench.  I need to reconstitute them and (tomorrow) get instructions form the PhD student on where the PCR reagents are and how to use the PCR machine.

4. Learning R:

Our Life Sciences Centre R group is holding lots of workshops and help sessions, but by bad luck they've mostly been at times I can't attend.  The summer student has given me some R homework, but it's buried on my desk.

5. Clean my desk:

To find the R homework.

Completing the toxin/antitoxin project

The honours undergraduate has finished her work on the toxin/antitoxin genes, and has submitted and successfully defended an extraordinarily good thesis.  The thesis is written as a polished scientific paper, which is ready to submit except for some gaps in the data.  My job now is to fill these gaps.

She began by confirming the mutant phenotypes. We've already shown (Sinha et al. 2012) that the toxin knockout prevents DNA uptake as well as transformation. Under competence-inducing conditions the antitoxin knockout prevents transformation, and the toxin knockout and double knockout both have normal transformation.  This is consistent with our hypothesis that the Toxin protein does something that prevents competence development, and the Antitoxin protein prevents this.

Her next step was a phylogenetic analysis of the toxin and antitoxin genes.  This work is complete. She identified three groups with the gene pair, and used synteny analysis to show that the pair entered these groups by independent horizontal transfer events.
Because there is no outgroup she was unable to root the tree, so the order of transfer events is not established.

Actinobacillus and Haemophilus species are all members of the Pasteurellaceae, and A. pleuropneumoniae has a competence system very like that of H. influenzae.  Her next project was to knock out the A. pleuropneumoniae toxin and antitoxin homologs, so see if they affect competence in the same way the H. influenzae homologs do.  She used PCR to amplify and clone the genes with flanking DNA, and then used inverse-PCR to create linearized versions of this plasmid lacking either or both of the genes.  (I'd better diagram this out because I'm going to have to do some similar work.)

She then ligated SpcR cassettes into the linear PCR products, creating circular plasmids with the SpcR cassette replacing each (or both) genes, and transformed these back into the A. pleuropneumoniae chromosome, selecting for the cassette.

Once she'd made her knockout mutants she checked their competence phenotypes, and found that all were normal.  She would have concluded that the A. pleuropneumoniae toxin does not block competence development, but in the interim she had discovered that the GenBank record for the toxin gene was incorrectly annotated, missing the last 5 amino acids because these overlap with the start of the antitoxin gene.  This meant that the antitoxin knockout she had created was also missing the end of the toxin gene, and therefore might have actually been a double knockout (if the terminal toxin amino acids are important for its function).

So one of my jobs is to redo the antitoxin mutagenesis, using a new primer that preserves all of the toxin gene, and check the growth and transformation phenotypes of this new correct mutant.  If I find that it has a transformation defect I'll also check its DNA uptake.

 She then examined the growth properties of the H. influenzae and A. pleuropneumoniae mutants.  The A. pleuropneumoniae mutants all grew normally, but the H. influenzae antitoxin knockout grew significantly slower in log phase than the other H. influenzae strains.  This suggests that the H. influenzae toxin, when unopposed by antitoxin, interferes with cell growth even under conditions where the cells are not competent.  The toxin knockout also grew a bit slower.

She then examined the RNAseq data for the wildtype and knockout strains.  Each culture was sampled and sequenced at four time points: T0 is log phase in rich medium, just before transfer to the competence inducing medium MIV, and T1, T2 and T3 are 10, 30 and 100 minutes after transfer to MIV.  

Here we discovered another problem – two of the ‘antitoxin-knockout’ cultures used for these analyses were incorrect - they were instead knockouts of a different competence gene, comN.  So we have three replicates of the toxin and double knockout strains, but only one correct replicate of the antitoxin knockout strain.  The two missing replicates have been recultured and their RNA preps remade, and they are now being sequenced.  My contribution here will be to help the summer student finish the analysis of this data.  (By 'help' I mean pester him with questions and requests for explanations...)

Analyzing the data she had showed that log phase expression of the toxin mRNA increases dramatically when the antitoxin is missing - this is consistent with characterization of related toxin/antitoxin pairs, where the antitoxin protein represses transcription of the toxin/antitoxin operon.  She found lots of other interesting (confusing/perplexing) effects.  Since most of these are based on the single antitoxin-kockout replicate, we'll examine them more thoroughly once we have all three replicates.

Still planning the mutagenesis project

OK, I'm abandoning the old mutagenized stocks and starting the mutagenesis project from scratch.

I want to take advantage of having selectable alleles linked to each of the genes where mutations produce hypercompetence.  That's 1) the StrR point mutation linked to sxy (about 50 kb; 50% cotransduction), 2) the CatR cassette linked to murE (about 4 kb; 90% cotransduction?), and the CatR cassette tightly linked to rpoD (~100 bp; > 90% cotransduction).

Last time I tried to do this by doing the EMS-mutagenesis in vitro (See blog posts: here and here). Call this strategy C.   I directly mixed EMS with DNA of three strains carrying the above selectable markers and wildtype alleles of the hypercompetence genes, and then transformed the mutagenized DNAs into competent wildtype cells, selecting for each of the marker strR and CatR alleles.  But this failed.  I didn't get any low-level novobiocin resistance mutations that would have indicted that the EMS caused mutations, and I didn't get any hypercompetent mutants.  I suspect that transforming cells with EMS-damaged DNA is a very inefficient way to create mutations.

This time I'm going back to doing the mutagenesis in vivo (call this strategy B), incubating the three marked strains with EMS and allowing them to grow for 1-2 hr to convert the damage into mutations.  Then I'll isolate the DNA from each mutagenized culture, and use this to transform wildtype cells to the three markers.  This will give me pools of cells that have experienced a high mutation rate in the neighbourhood of sxy, murE and rpoD.

But before I do this I need to do the math, to see if this is really any better than just mutagenizing wildtype cells and screening them all for hypercompetence (call this strategy A).  Having the linked markers will certainly be handy later, once I've found hypercompetence mutants and want to find out which gene they're in.  But will using them as described above really let me find more mutations in these specific genes?

OK, I've laid out a situation with realistic numbers, and I've run it by the PhD student and the summer student.  Bottom line:  Strategy B does not enrich for mutations in the desired region.  It's only strength is that it allows use of much higher concentrations of EMS than would be tolerated in Strategy A.

Here's the numbers analysis for strategy A:

  1. Start with 10^9 cells.  (Below I'll consider whether fewer would be OK.)
  2. Treat with EMS (0.08M for 30 min).  Previous work suggests that this creates about 1 mutation per surviving cell.
  3. Grow cells for about 3 hr or more, keeping cell density below OD600= 0.2.  The goal is to allow enough time for the cells to recover from the DNA damage, undergo two rounds of DNA replication to convert some damage into G->A mutations, and express the mutant hypercompetent phenotype.  Assume that the cell numbers increase 10-fiold during this period.
  4. How many mutations will we have?  The genome has about 4 x 10^5 Gs.  Mutations at some of these will be lethal or sub-lethal, so assume about 2 x 10^5 positions where mutants have normal or near-normal growth.  With 10^10 cells, we will have about 5 x 10^4 occurrences of each mutation, on average.
  5. How many hypercompetence mtuations will we have? 10 of these are positions we have already found to be sites of hypercompetence mutations, so in our 10^10 cells we'll have at elast 5 x 10^5 hypercompetent cells.
  6. To select for these hypercompetent cells, transform the OD600=0.2 cell population with the 8 kb NovR DNA fragment from plasmid pRRnov1.  This DNA transforms much better than the equivalent PCR fragment or than chromosomal DNA with the equivalent mutation.
  7. The normal cells will transform at a frequency of about 10^-8 - 10^-7.  (I'm guessing here; with chromosomal DNA it's ~10^-9.) That would give about 100-1000 novR colonies from the 10^10 cells.
  8. The hypercompetent cells will transform at higher frequencies:  rpoD: cells with the known rpoD mutations will probably have a TF of 10^-4 - 10^-3, so the 5 x 10^4 cells of each mutation would give about 5-50 novR colonies.  sxy:  cells with the known sxy mutations will probably have a TF of 10^-3 - 10^-2, so the 5 x 10^4 cells of each mutation would give about 50-500 novR colonies.  murE:  cells with the known murE mutations will probably have a TF of 10^-2 - 10^-1, so the 5 x 10^4 cells of each mutation would give about 500-5000 novR colonies.   
  9. According to this analysis, most of the colonies will be hypercompetent mutantx, and most of the hypercompetence mutants we find will probably have their mutations in murE. Of coulrse this only considers the hypercompetence mutations we already know about.
Things to do first:  
  1. Check that our EMS stock is still good, by mutagenizing wildtype cells and scoring low-level novobiocin resistance.  A resistance frequency of ~5 x 10^-6 is approximately one mutation per viable cell.
  2. Make a big prep of pRRnov1.  The yield of this plasmid is often poor so this may take several attempts and large volumes.  Check the transformation frequencies and efficiencies this DNA gives.