Field of Science

Today's work on the RNA-seq samples

The Co-op tech has pressed half the samples through the RNeasy kit spin columns, and will probably get the rest done today.

But she also is still working on the PCR checks (problems getting amplification from the colony-DNA material) and redoing some of the transformation assays (using the frozen cells I'd saved for each sample) because two of her original tests gave surprisingly low transformation frequencies.

I'm hoping we can also get started on the next steps today - checking the RNA concentrations using the Nanodrop and running aliquots in a gel to check that the rRNA bands are intact.  First step for this is to clean up a gel box and comb and see if we have any RNA-loading dye made up.

Other tasks for the soon-to-be-departing Co-op tech

The Co-op tech will be leaving us at the end of the month.  Last week she gave an excellent bab meeting presentation, and this revealed a couple of loose ends and interesting possibilities that should be cleared up before she leaves.

First, she and the sabbatical visitor (now back home in Regina) isolated a new mutation in rpoD that causes hypercompetence.  But this strain hasn't yet been added to our formal collection of frozen strains and the associated 'Strain List' database.  This is essential and urgent.

Second, the mutant hunt turned up a couple of other 'possibly hypercompetent' mutants whose phenotypes haven't been checked out.  These need to be checked in competence time courses, and added to the Strain List collection if they turn out to be genuinely hypercompetent.

A weird result that I'd forgotten about is the finding that cells transformed with a mixture of NovR and KanR DNA fragments (generated by PCR) showed a much LOWER cotransformation frequency than expected.  We suspect this reflects some chromosome-level interaction between these linked segments, but we'd first need to reproduce the result.

RNA-seq progress

I've collected and frozen all the 24 samples for our make-up RNA-seq run. (not the Trizol-prep ones - they've been deep-sixed).  And this morning the co-op tech learned to prep RNA from each sample. She's done the first 4, and will do the rest over the next couple of days.

The next steps are:

  1. Complete the PCR tests of strain genotypes and the analysis of transformation frequency data. She's still working on the PCR tests, but so far everything looks OK.
  2. Check the RNA concentration using the Nanodrop
  3. Run aliquots of the samples in a gel to check integrity of the rRNA bands (surrogate for integrity of the mRNA).
  4. Treat 5 µg of each sample with DNA-free.  We found our stock from last year, and there's still enough to treat all our samples.
  5. The former RA says we can take the DNA-free-treated samples directly to the RiboXero ste; we don't need to first do a 'clean-up' step with the RNeasy Minelute kit spin-columns.
  6. Treat an aliquot of each sample with RiboZero.  We will use only half as much RNA as recommended, and only 1/4 as much of the other reagents (in 1/4 of the recommended volume, of course). This will let us treat 24 samples with a 6-treatment RiboZero kit.
  7. Give the samples to the former RA in her new lab for library preparation and sequencing.

What can I recover from an old failed experiment

About 18 months ago I did a big mutagenesis experiment, intending to isolate new hypercompetent mutations.  I made several mistakes and the experiment was a failure, but I did freeze stocks of intermediate cultures.   At the time I thought that some of these could be used in a future attempt, because they came from stages before the mistakes were made.

I still want to repeat this experiment, and I just found the stocks in the -80 °C freezer.  Now I need to decide which are potentially useful, and throw out the rest.  Here's photos of what I found:


The letters A-G refer to different strains, each with a wildtype version of a gene known to give rise to hypercompetence-causing mutations, and to different levels of mutagenesis
  • A, B & C: wildtype cells, incubated in 0 (A), 0.05 (B) and 0.08 (C) M solutions of the mutagen EMS.
  • D & E: strain RR514, which has a Streptomycin-resistance mutation (StrR) close to the wildtype sxy gene, incubated in 0.05 (D) and 0.08 (E) M solutions of EMS.
  • F & G: strain RR805, which has a chloramphenicol cassette (CmR) inserted within a few kb of (= closely linked to) the wildtype murE gene, incubated in 0.05 (F) and 0.08 (G) M solutions of EMS.

The big tubes turn out to be useless, since they contain cells that were incubated with the wrong DNA after the mutagenesis.  Most of the small tubes also are from stages that have been incubated with DNA, (e.g. label 'F DNA'), but others (the ones labeled '90') were frozen after 90 min of post-mutagenesis growth, before the DNA addition step.  These ones I can use.

The first step now is to do a test I didn't do in the original experiment, to check that the EMS mutagenesis did indeed cause mutations by plate some of the cells on low-concentration novobiocin. I'll do this test on the wildtype cells (B & C), so not to unnecessarily use up the more valuable cells in the marked strains (D-F).  I don't have tubes of the control A culture, so I'll just use normal wildtype cells.

If this test shows that the mutagenesis worked, I have two alternatives.  1.  I could isolate DNA from the mutagenized marked cultures and use it to transform wildtype cells to StrR (D & E, to enrich for cells with sxy mutations) or CmR (F & G, to enrich for cells with murE mutations).  Then I'd enrich these transformants for hypercompetent mutants by transforming them with the PCR'd NovR fragment (after first testing that this works well).  2.  I could do the hypercompetence-selection transformation first, and then isolate DNA and transform wildtype cells with selection for the linked marker.  The advantage of 1 is that I can pool many thousands (millions?) of transformants, maintaining whatever genetic diversity my mutagenesis has created in the gene of interest.

New RNA-seq work

(OK, I just checked, RNA-seq should be hyphenated.)

I've made a couple of posts about plans for new RNA-seq work on the Sense Strand blog: and

Now it's time to get down to work.

Here's the planned samples.  We have 26 on the list, but a standard run will only be 24, so two need to be dropped.  Conveniently, the two KW20-in-Trizol samples might not be needed, depending on the available small-RNA data for H. influenzae, so I won't consider those right now.

For the rest, we have 6 mutant strains.  Given the mixups that have occurred so far, it would be prudent to check these every way we can.

We are checking antibiotic resistances and will transform each strain when we grow its culture and collect the samples for the RNA preps. One of the Honours Zoology undergrads has already checked the toxin and antitoxin strains by PCR (she's the one who suffered most from the mixup), and we'll use the same primers to check the cultures we're sampling from.  We need the former RA's help to find the primers for the ∆hfq mutant, but luckily its phenotype is quite distinctive - in our whole collection I can only think of one other mutant that has its competence down 10-fold.  RR753's phenotype is also distinctive, hypercompetent, but not very.  The crp and sxy mutants have the same phenotypes (same drug resistance, same complete lack of competence.  We have lots of sxy PCR primers, but we'll have to check to see which ones will work with this insertion mutant.  And, for both crp and sxy there's the additional problem that miniTn10 insertions do not amplify well because of their end repeats.  I wonder if we have an internal primer for miniTn10kan.

The co-op tech has checked antibiotic resistances and frozen fresh stocks of all the strains.  She's inoculated the 4 strains for the MIV-competence preps, and tomorrow we'll toy to collect all those samples.  (No, I haven't done anything in preparation yet.)

Planning the DNA sequencing part of the PhD student's project

The former post-doc (I'll call him the FPD) visited yesterday afternoon, and we had intense discussions of how to proceed with both the RNAseq work (summarized here on our Sense Strand blog) and with the PhD student's planned DNA uptake experiments.

His planned experiments take advantage of the phenotype of a rec2 knockout mutation.  These cells take up DNA normally across the outer membrane, into the periplasmic space, but they cannot transport it across the inner cell membrane.  This allows him to recover intact DNA that has been taken up, and to use DNA sequencing to compare it to the input DNA the ∆rec2 cells were given.

Some of the experiments will use genomic DNA of the species being tested, fragmented to appropriate length distributions, and some will use synthetic DNA fragments (~200 bp) containing a 30-50 bp stretch of random sequence (see figure).

The FPD, who developed the synthetic fragment protocol, pointed out that his experiments had used full lanes of Illumina sequencing only because it was not then possible for us to 'barcode' our different DNA samples and mix them for sequencing as a single lane.  The sequencing depth he obtained was useful, but it will be extreme overkill for the experiments the PhD student plans.  So we need to design barcoding into our analyses, so we can mix up to 24 samples in one lane for sequencing, and then separate the resulting sets of sequence reads by their different barcodes.  We'll still need to use two lanes, because each 'recovered' sample will need to have a corresponding identical 'input' sample.  Because these samples will have the same barcode they could not be distinguished if they were sequenced in the same lane.

So rather than doing one very-deeply sequenced experiment, he'll be able to do multiple replicates, each sequenced at a moderate but entirely adequate depth.  If he uses a HiSeq machine for the sequencing, he'll be able to get 1.6 x 10^8 reads for each of 12 samples; with a NextSeq this would give 4 x 10^8 reads per sample. (Is that right, per sample, not per lane?).

One issue to keep in mind is that it would be foolish to save all the sequencing for one big batch at the end of the thesis work.  Instead the work needs to be designed with an initial set of samples to be sequenced, so he can (1) tell whether everything is working as it should, and (2) begin analyzing sequence data from one part of the project while generating additional samples for other parts.  For a preliminary batch of sequencing, it might be better to use a MiSeq machine, whose smaller capacity would let us sequence a few samples more economically.

We also talked about how long the random-sequence segments should be in the 200 bp fragments, and about where to locate the barcode segments.  These consist of an independent sequencing primer followed by 8 bp that identify the source experiment.  Putting these to the right of the random segment will let him efficiently create the double-stranded 200 bp fragments, using the same long left-side oligo (containing the random segment) with many different right-side oligos, each containing a different barcode.

Sensitivity of the PhD student's planned analysis

The PhD student is proposing to use Illumina sequencing of input and recovered-after-uptake DNAs to detect possible biases in uptake of DNA by bacteria other than H. influenzae.  (This is a simplified version of the analysis proposed in our funded NSERC proposal.) We're discussing the factors that will affect the sensitivity of this analysis, so he can say how strong a bias would have to be in order for his experiment to detect it.

The factors we've thought of are:

A. Nature of the preferred sequence pattern: 
  1. How long is it (3 bp? 10 bp?)?  How specific is it (e.g. is each base specified, or just 'purine' or 'pyrimidine'?  Together these determine how often this pattern will occur in the input DNA (by chance or due to uptake bias-drive).
  2. How strong is the bias favouring uptake of fragments containing this pattern?  How strict is the preference (are variants of the specified pattern also taken up, but less strongly)?  Are fragments with more than one occurrence of the pattern more likely to be taken up?
 B. Properties of the input DNA:
  1. If this is genomic DNA, what is the size range of the fragments?   The sensitivity of the experiment will be low if the fragments are so large that each has at least one occurrence of the preferred pattern.
  2. If this is a synthetic fragment containing a fully degenerate segment, how long is the degenerate segment?
C. Sequencing coverage:
  1. How high is the sequencing coverage?  Is it the same for the control input DNA and for the recovered DNA?  This will determine the noise due to random factors.  
  2. Does the error rate of the sequencing matter?
  3. For genomic input DNA, are there position-specific differences in coverage across the genome?
  4. For degenerate-fragment DNA, are there non-random factors in the input DNA or in its sequence-ability?
He's going to start by working through the values for a very-strong-bias case, detecting the H. influenzae uptake sequence in genomic DNA (figure below), and then relaxing the inputs.

Mutagenesis plans

(I'll add some explanations later.)

1.  Mutagenize more RR805 DNA, using a range of high EMS doses (10, 15, 20, 25, 30, 40 min in 50 mM).  Transform this DNA directly into competent KW20 (without EMS inactivation or DNA purification) and select for CmR and maybe for NovR.

2.  Mutagenize RR805 cells, using a range of high EMS doses (from expt. #180, 80 mM for 1 hr gives ~10^-2 survival).  The cells don't need to survive, because I'll just grow the culture for a couple of hours and then extract all the DNA and use that DNA to transform KW20 to CmR.

For both 1 and 2, then pool CmR transformants and transform at low cell density to StrR with RR514 DNA.  Test individual StrR colonies for hypercompetence by colony transformation with MAP7 DNA.

3. Mutagenize NovR and NovS PCR fragments (made by the sabbatical visitor), using the same EMS concentrations as in experiment 1.  Then test the effects of the EMS mutagenesis by transforming each DNA into KW20, looking for gain of NovR in cells transformed with the NovS DNA, and loss of transforming ability of the NovR DNA.

I can do experiments 1 and 3 today (if I first pour lots of plates).  I can then do Experiment 2 tomorrow or on the weekend, once the cells have grown up.


1.  I must have put too little chloramphenicol in the Cm plates for this experiment, because all the cells grew on the Cm plates.  I need to repeat this experiment.

3.  Increasing exposure to EMS caused decreased transformation by the NovR fragment, as it should, but the corresponding exposures of the NovS fragment gave no NovR transformants, indicating no detectable mutagenesis.  So the decrease seen with the NovR fragment may just be due to damage, not mutation.

2.  My streak of RR805 cells has grown nice little colonies.


I've inoculated one of the RR805 colonies for an overnight culture, so I will be able to do the experiment 3 cell mutagenesis tomorrow.  And tomorrow I'll make lots and lots of Cm plates, with the right amount of chloramphenicol, so I can also repeat experiment 1.

No new candidate mutants (sigh...)

As I planned here, I pooled the CmR colonies resulting from transformation with EMS-mutagenized CmR murE+ DNA, and grew them to log phase (OD600 ~ 0.1).  The murE+ cells in the pool should have been non-competent under these conditions, but any murE* hypercompetenc mutants should have been competent.  To select for these mutants I transformed the cells in each pool with DNA carrying a streptomycin-resistance mutation, and plated on Str plates.  One pool gave several hundred StrR colonies (many more than I would have expected as transformants), but the other pools had very few or none (4 total). I then screened individual StrR colonies by mixing them with dilute NovR DNA and plating on Nov plates.

Unfortunately none of the StrR colonies transformed to NovR at the high frequency seen for the positive control (murE749) colonies.  In fact, none transformed any better than the murE+ negative control colonies.

This is a bit surprising, given that the 2-fold higher level of EMS mutagenesis reduced by 100-fold the ability of the CmR cassette to transform cells, and the 4-fold higher level eliminated it entirely.  I had assumed that this reduction/elimination was due to too-heavy mutagenesis, but perhaps it was a direct consequence of the DNA damage.  One possible explanation I'm considering is that damaged DNA is almost always repaired or destroyed, and rarely gives rise to recombinants.  Another possibility is that, when cells are mutagenized, the mutations arise mainly only when levels of damage are so high as to overwhelm the repair systems, allowing the damaged bases to be used as templates for DNA replication.  Maybe this also requires induction of the error-prone DNA polymerase.

So now the sabbatical visitor and I are designing a control experiment, to test whether this direct DNA mutagenesis is working as we think it should.  We're going to mutagenize two versions of a DNA fragment containing the gyrB locus.  One is wildtype, and the other has the novR allele we usually use in our transformation assays.  We expect the transformation efficincy of the novR allele to decline with high doses of EMS, and we hope that now novR mutations will arise from high doses to the wildtype allele

* Here's some wishful thinking: Ideally we should be selecting for a G->A transition mutation because those are what EMS induces best.  But we're using novR (G->T) because we have the porimers handy and know they work.  The mutation spectrum of EMS is reported to be much broader with the in vitro mutagenesis we're using, so we hope this will work.  But I just checked the numbers and they didn't see ANY of the kind of change we'd need.

Really we should use selection for streptomycin resistance, since its T->C mutation is a type that arose at high frequency with the in vitro EMS treatment.  I wonder if we have the primers for this - I think the post-doc might have gotten them for us.

Mutagenesis results

I don't have any novobiocin-resistant transformant-mutants after 24 hr (though slow-growing colonies might appear later), so I can't use that to tell how effective the mutagenesis was.  But I have tons of chloramphanicol-resistant ones at the low exposures to EMS (2, 5 and 13 minutes), 100-fold less at 30 minutes exposure and none at 60 minutes exposure (the highest dose).  This tells me that the EMS was doing its job, and that the DNA damage caused many potential transformants to have lethal mutations either in the CAT cassette or in nearby genes in the recombination tract.

So I think I'll go ahead and make pools of colonies from the 5-min and 13-min treatments and enrich them for hypercompetent mutants by selecting for StrR transformants in log-phase cultures. Then tomorrow I can screen these for hypercompetence by our crude colony-transformation assay.

Why not also the 2-min treatment?  OK, I'll include one pool of those too.

I'll have four five pools (10^4 and 10^5 transformants from each of the two treatments), which will be easy to handle.  What control cultures should I include?  RR805 (murE+) will give negative control colonies, and RR797 (murE749) will give positive control colonies.

* One reason to not use the ~1000 CmR colonies from the 30-min dose is that these are less likely to have recombination tracts extending all the way from the CAT cassette to murE.  That's because this segment contains two essential genes (ftsI & ftsL), and recombination tracts that cover the CAT-murE distance are much more likely to have had a lethal mutation in one of these genes than are tracts that don't reach to murE.

Mutagenesis planning in progress

By midday today I'll have checked my strains and made my DNA. 

The strains are RR805, which has a CAT cassette linked to the murE+ gene (normal competence), and RR797, which has the same cassette linked to the murE749 hypercompetence allele and a StrR point mutation elsewhere in the chromosome. I've checked their antibiotic resistances, done platings that will confirm their competence phenotypes (will count colonies this morning), and made crude DNA preps (I'll complete purification this morning).

Next I should do the mutagenesis dose-response curve, and I've now realized that this experiment can also be used for the first hunt for more hypercompetence mutants. 

Mutagenesis (today?):

Set up one tube containing 12 µg RR805 chromosomal DNA in 120 µl water or TE, at 37 °C.

Take a 20 µl time = 0 sample (see below).

Add EMS to the remaining DNA, to a final concentration of 50 mM. 

Take samples at time = 2, 5, 12, 30 and 60 minutes.  Immediately add each sample (including t = 0) to 100 µl of 5% sodium thiosulfate, which will inactivate the EMS and stop the mutagenesis.

The t = 2  sample will have had about 6-fold less exposure to EMS than used by the Lai et al. paper, and the final sample will have had 5-fold more.

Add NaCl to each sample to 0.15 M and add 2 volumes of ethanol to precipitate the DNA.  Rinse the pellets (probably invisible) with 70% ethanol and air dry.  Resuspend each in 50 µl TE.  (If the invisibility of the pellets is a problem I could add some E. coli DNA as carrier, since this won't interfere with the subsequent transformations.)

Transformations (today):

Thaw out lots of vials of frozen competent KW20 cells (wildtype).  I need one tube for each of the 6 DNA samples, and also one for RR797 DNA (chloramphenicol resistance control) and one for MAP7 DNA (transformation control).

Add 2.5 µl (= 100 ng) of each DNA to a tube containing 1 ml of cells.  Incubate for 15 min at 37 °C.

Add 3 ml sBHI and incubate for 90 min longer, to allow expression of the chloramphenical resistance.

Dilute and plate on plain plates (10^-6, 10^-5), Nov1 plates (for low-level novobiocin resistance, plate undiluted and 10^-1) and Cm1 plates (plate 10^-3, 10^-2, 10^-1 and undiluted).

Freeze the remaining transformed cells in case I want to do more with them later.

Analysis and next steps (Friday):

Use the colony counts to assess the extent of mutagenesis and gene inactivation.  For doses that gave high NovR mutagenesis without reducing the CmR transformation rate, make pools of the CmR colonies from plates that have >1000 colonies (one pool per plate). 

Then I cna grow each pool to early log in sBHI and transform it with StrR DNA to enrich for hypercompetent mutant.

Then I'll screen individual StrR colonies for hypercompetence by mixing them with MAP7 DNA and plating on Nov.

What if I don't get any NovR mutants? 

My previous use of EMS, mutagenizing cells, not DNA, gave NovR mutants at about 10^-6 of the survivors.  If this was the level of NovR mutations in my mutagenized DNA, the transformation assay probably wouldn't detect their presence because only about one cell in 1000 will have recombined the nov-containing DNA fragments, giving a transformation rate of 10^-9, below the detection limit.  But I expect the mutation rate to be much higher for the pure DNA, so I'm hoping that I'll see significant increases in resistant colonies.

 If I don't?  I could just go ahead and screen a couple of the high-dose CmR pools for hypercompetent mutants anyway, since if I find some then I can just forget about the Nov test.  If I don't find any hyprecompetent mutants I should repeat the mutagenesis using a NovS DNA fragment as control.

What mutation rate do I want for my experiment?

I need to decide on a desirable mutation rate for my murE mutagenesis experiment (described here).  To do this I need to think about (at least) how big the gene is, how large a region of the gene I want to investigate, what fraction of mutations will interfere with or eliminate gene function, and what fraction of mutations might cause hypercompetence.

How big is the gene?  1467 bp (489 aa).

Are hypercompetence mutations  equally likely to occur anywhere in the gene?  The mutations we have are in domain 3, at amino acids 361 and 435, so maybe other mutations would be nearby.  But maybe not.  Let's first consider the whole gene, and then decide * if focusing on the last third of it would make any difference.

What are the expected frequencies of mutations with different effects?  About 50% of random base changes change an amino acid (surely someone has done this calculation...).  Since all three of our known mutations change an amino acid, let's assume that silent mutations don't affect competence. About 34% of random amino acid changes interfere seriously with protein function (Guo et al. 2004).  Our known mutants appear to have normal MurE catalytic function, and defective mutants will not show up in our screen because murE is an essential gene.  So that leaves about 1/3 of all the mutations as causing well-tolerated amino acid substitutions.

What fraction of well tolerated amino acid substitutions cause hypercompetence?  We know of three that do.  How many different amino acids can each codon mutate to?  Probably about 9 or10 on average.  So let's say we have 500 codons of interest, that's about 5000 different possible amino acid substitutions.  About 2/3 of these will be well-tolerated.  So we know that 3 out of 3,300 amino acid changes cause hypercompetence.  Other mutations may cause hypercompetence too, but since half the mutations will be silent, this lower-bound means that at least 1/2000 colonies with a single murE mutation can be expected to be hypercompetent.  That's pretty good odds, given that our transformation-selection step can enrich 1000-fold for hypercompetence mutations.

So an average of 1-2 mutations per kb should give us easy-to-find hypercompetence mutations. Will higher mutagenesis give us more? Issues to consider:
  1. More mutations means more non-tolerated mutations, which means that some hypercompetence mutations won't be seen because their cells died.  I don't think this is a big deal, unless we made the mutation rate very high.
  2. More  mutations means more irrelevant mutations in each gene we sequence.  This is important.  Inference will be greatly simplified if genes from hypercompetent cells have only one mutation.  So it's probably best to  use the lowest level of mutagenesis that will give us easily-detected mutants.
The Lai et al paper had 5-6 mutations per kb. This is probably too high for us.

Another concern is mutations in the genes between the CAT cassette and murE.  Some of these are essential, and mutations in them will reduce the frequency of recovering viable transformants that contain both the CAT cassette and murE.  This is another reason to go for a low mutation rate.

* Back to a previous point.  Does it matter whether we want to screen only the last third of the gene?  No, because we don't have any way to isolate this from the rest.

murE mutagenesis planning

 (Edited on March 1, after more thinking and planning.)

I want to create a pool of cells with random point mutations in the H. influenzae murE gene, and to select and screen this pool of cells for hypercompetent mutants.  I'm going to do this by mutagenizing the DNA with the chemical mutagen ethyl methanesulfonate (EMS) in vitro and then transforming it into cells, rather than mutagenizing cells.

One unanticipated benefit of the in vitro method is that the mutation spectrum is better.  With in vivo mutagenesis, EMS produces mainly  GC-to-AT transition mutations by alkylating guanines in DNA, creating O-6 ethylguanine which mispairs with T instead of C during DNA replication. (info from Wikipedia). But the in vitro work found a much less biased distribution, with 42% GC-to-AT transitions, 34% AT-to-GC transitions, and 24% GC-toCG transversions.


Step 1. Cut chromosomal DNA of strain RR797 RR805 with the restriction enzymes KpnI and BglII. This strain contains the wild type allele of murE (oops, no, this strain has the murE749 hypercompetence allele! The strain I want is RR805.), and has a chloramphenicol resistance cassette inserted about 5 kb away from the site of the known murE mutations,. This digest creates an 8 kb fragment that contains both the CAT cassette and the wild type murE allele.

Here's the map:

This pre-digestion step could probably be omitted if necessary, because random fragmentation of the DNA will accomplish almost as much. But it shouldn't hurt, and it might double the frequency of cotransformation.   But I just looked at some old cotransformation data, and I see 60-70% linkage (selecting for CmR gives the linked murE allele), which is very good

Step 2. Soak this DNA in an EMS solution for 1 hr.

Step 3. Wash the DNA and transform it into competent wildtype cells.  Use about 100 ng DNA per ml, so that each cell is likely to recombine only a single DNA fragment.  As a control, transform the same cells with DNA from the chloramphenicol-resistant murE749 strain.

Step 4. Select for chloramphenicol resistance, to enrich for cells that have recombined in murE.  This will also confirm that the level of DNA damage was not so high as to limit transformation.  I should be able to get many thousands of independent transformants.

Step 5. Pool chloramphenicol resistant colonies, creating separate pools from independent sets of transformants.  Aim for about 5 pools.  Freeze some of the cells of each pool.  Make a pool for the control transformants too.

How many colonies should be in each pool? I want enough colonies per pool that each is likely to contain at least one hypercompetent mutant - how many colonies will this be?  I know of three mutations that produce hypercompetence, which would let me predict the minimum expected frequency of hypercompetent colonies if I knew the frequency of mutations in the DNA and the degree of linkage in the transformation.  I can measure linkage by doing colony assays on the control transformation.  The enrichment can increase the frequency of hypercompetence by 1000-fold, if all the mutants are as hypercompetent as the ones we have.  So if the frequency of hypercompetence in the chloramphenicol-resistant transformants is 1/1000, I should put at least 1000 colonies in each pool.  If it's less, I should put more. 

Step 6. Grow the pooled cells in sBHI at low density for a few hours, then transform with cloned or PCR'd NovR DNA (or a different marker?).  Plate on nov plates.  Do this with the control murE749 transfornation too.

Step 7. Screen individual NovR colonies for hypercompetence by touching them to nov plates and then resuspending the rest of the cells in sBHI containing MAP7 DNA and plating on Kan (or Nov?) plates.  Do only 10 colonies per pool, or 1/1000 as many colonies as went into the pool?  I expect most of the control colonies to be hypercompetent.

Step 8. For each pool, pick one or two high-transformation colonies from their toothpicked plate, and retest their competence with a simple time course.

Step 9.  PCR and sequence the murE genes from5 or 10 of the confirmed hypercompetent mutants (depending on how many I get, of course).  Are the known mutations present?  New mutations?

First we should test different levels of mutagenesis:  

The protocol we have (Lai et al.) says to use 1 µg DNA in 20 µl 10 mM EMS for 1 hr; this gave 5-6 mutations per kb in the clones they sequenced.  It also reduced the transformation efficiency of the plasmid insert they mutagenized to about 60%.  If they carefully standardized the amounts of DNA, this reduction should have been a direct consequence of DNA damage and repair processes, since they were not selecting for function of their mutagenized insert.

5-6 mutations per kb sounds pretty good for us (but see next post, which suggests we want fewer), since about half of them will be silent, but I think we should first try a wide range of concentrations.  For the cell mutagenesis (many years ago) I used 50 mM for 45 min and 80 mM for 30 min (RR expt # 181), but we want much heavier mutagenesis here.  So here let's try 0, 2, 5, 10, 20, 50, and 100 mM - that's 7 DNA samples to do transformations with.

Two assays for the extent of mutagenesis:  

1. (To identify an optimal concentration) Mutations creating low-level resistance to novobiocin: Mutagenize any novS DNA (e.g. RR805) and transform into KW20 and select for low-level novobiocin resistance (1 µg/ml rather than 2.5), to check the efficacy of the mutagenesis.  There should be an optimal dose of EMS, above which the frequency of nov resistance drops because the DNA is too damaged to recombine or contains too many mutations that block gene function.

2. (To identify concentrations that are too high) Mutations that inactivate the CAT cassette:  Mutagenize RR805 DNA and transform KW20 to chloramphenicol resistance. At some EMS dose the transformation frequency will decrease because the DNA is too damaged to recombine or contains too many mutations that block gene function.  (This test could also be done with any point mutation creating antibiotic resistance.)

What we know about the competence-regulon gene comM

The grad student of an upstairs colleague has been doing a lot of excellent work on the Rhodobacter capsulatus homologs of some H. influenzae competence genes, because he has discovered that they are also needed for gene transfer by GTA, the phage-related 'gene transfer agent'.

One of the genes he's looking at is comM.  ComM is predicted to be a cytoplasmic protein, a member of the YifB subfamily of AAA-ATPase proteins.  Here's a review about the AAA+ superfamily.  These proteins have a very diverse range of activities, so it's hard to make any prediction about a likely function for ComM from looking at its relatives.

ComM was originally studied in H. influenzae, by Michelle Gwinn and Jean-Francois Tomb in Ham Smith's lab.  They reported that their comM mutant had normal DNA uptake but reduced transformation (down about 300-fold).  It had normal expression of a lacZ fusion to another competence gene, indicating that it didn't affect regulation of competence.  (It also had reduced phage recombination, but we still don't know what this assay means.)

To find out why transformation was reduced, they followed the fate of end-labelled DNA fragments. The kinetics were like those of both wildtype cells and a rec1 mutant (rec1 is the H. influenzae homolog of E. coli's recA; it's absolutely needed for homologous recombination). So the authors concluded that the comM knockout does not affect the transport of DNA into the cytoplasm.  But their data doesn't distinguish between an effect on DNA degradation (indirectly preventing recombination) and a direct effect on recombination.

We've independently created a comM knockout; its DNA uptake is also normal, and its transformation is also down, but only about 20-fold.  We haven't done anything more to evaluate its phenotype.

*Interestingly, Gwinn et al. commented that "In addition, HI1117 has homology to a magnesium chelatase gene of Rhodobacter capsulatusbchI, involved in bacteriochlorophyll biosynthesis (1) and to related genes from other photosynthetic organisms."

Grant proposal's done! What experiment shall I do?

I clicked 'Submit' on my grant proposal last night; my immediate teaching responsibilities are light, and there's nothing else big on my plate, so now I get to start doing experiments again!

I think the most fun thing to do will be to join the sabbatical visitor and the co-op tech in doing mutant hunts for hypercompetent strains.  They're mutagenizing the rpoD gene and screening for new mutations that cause hypercompetence, and I can use the same methods on the murE gene.

This old post describes what we know about the relationship between murE and competence.  Well, what we used to know, because now we have new RNA-seq data that will tell us how transcription changes.  Basically, we have four independent mutants that all cause very similar extreme-hypercompetence phenotypes.  murE749 is the main one we've studied.  Some lacZ-fusion analyses indicate that it acts by causing overexpression of genes in the competence regulon (we looked at two genes) and one low-quality microarray appeared to confirm this and (maybe?) show some overexpression of sxy (the regulatory protein that controls expression of the competence regulon).

We assume that the other murE mutations act the same way.  But we have absolutely no idea how the mutations cause the phenotype.  MurE is an essential cytoplasmic enzyme in the pathway that synthesizes the cell wall.  The mutants all grow normally (though we haven't done a BioScreen run), and are not unusually sensitive to any simple test of cell-wall function.
One big part of the puzzle is how the mutations change the protein. The diagram above shows that three of the mutations change a poorly conserved amino acid (at position 435); these changes wouldn't be expected to have any serious impact on the enzyme's catalytic function.  So how do they have such a big impact on cometence?

On the other hand, the mutation in murE751 changes the strongly conserved leucine at position 361 to a very different amino acid (serine).  Leucine is hydrophobic but serine is polar, so they make very different interactions with their surroundings.  Because this leucine is highly conserved we think it must play an important role in the enzyme's catalytic function.  This would explain how the mutation can have a big effect on competence, but leaves us instead wondering why it doesn't have a big effect on cell growth.

I need to do several things:
  1. Update my reading to find out what's been learned about MurE function since we published our paper way back in 2000.
  2. Dig into the new RNA-seq data to see what it tells us about RNA changes in the murE749 mutant.  This will require finally learning some R and/or getting help from other lab members.
  3. Isolate new murE mutations that also cause hypercompetence.
Lots of fun!

Yes, I'm still here

The last-chance-for-everyone CIHR grant proposal deadline is Friday at 8:30 am!  After that's in, I promise to get back to the bench and back to proper research blogging.

Is there DNA in oreos?

I have to weigh in on this.

I spend a lot of time discussing the idea that bacteria can use DNA as a source of nutrients, and audiences are always surprised when I show this graphic and point out that DNA is ubiquitous in our foods.  And these are scientifically sophisticated molecular biologists and microbiologists.

So it's not at all surprising to me that 80% of the general public would check 'Yes' when asked, in a set of survey questions about food labelling and regulation, whether foods containing DNA should be labelled as such.  Instead of laughing at their ignorance we should think about how much expert knowledge is needed to evaluate this issue.

Many people, if guided by a series of prompting questions, could figure out that there's probably some DNA in at least some natural foods. But would you expect someone who hadn't taken high-school biology, or took it a long time ago, to know the answers to any of these questions?
  • Is there DNA in meat?
  • Is there DNA in leaves?
  • Is there DNA in potatoes?  In rice?  In noodles?
  • Is there DNA in fruit?  What if you don't eat the seeds?  In fruit juice?
  • Is there DNA in beer?  In wine?  In scotch? 
  • Is there DNA in flour?  In butter?  In olive oil?  In oreos?
  • Is DNA destroyed by being cooked?
  • Does DNA break down (like some vitamins) when food is stored?
  • Does DNA dissolve in water?