Why are we still using an archaic procedure to make H. influenzae competent?

Last week I did an experiment in parallel with one of the post-docs. Her preparations of competent H. influenzae had not been very competent at all, and we wanted to check if there was a problem with her procedure. We both produced cells that were reasonably (not excellently) competent, so we concluded that her past problems were not due to faults in her procedure.

In recent posts I've been describing my preliminary steps to finding a way to induce E. coli's 'cryptic' competence genes, and this and the above have started me thinking about the procedure we use for H. influenzae. It's a simple sprocedure: cells growing in a rich medium called sBHI are abruptly transferred to a starvation medium called M-IV. After about 90-100 minutes in this medium the cells are as competent as they are going to get. I deally this means that all ofthe cells are competent, each ready to take up a few hundred kb of chromosomal DNA.

This procedure was developed about 45 years ago, by more-or-less systematic fiddling with growth conditions. It hasn't been altered, simply because nobody has had a strong reason to bother trying. But now we know much more about what is going on when competence genes are induced, and we should be able to improve on this procedure. Improvements would certainly be useful, especially for the post-doc's ongoing survey of competence in 'wild' strains of H. influenzae. Being able to improve the induction procedure is also a good test of whether we really do understand how the genes are regulated.

What should we try? Adding cAMP should take the place of carbohydrate starvation, activating CRP to make its contribution to expression of CRP-S promoters. If we're right that nucleotide starvation is needed to induce sxy translation, we would like to try simpler ways to mimic this; I wonder if there's a specific inhibitor pf purine (or pyrimidine) synthesis? But this won't be much use if the cells are still in rich medium, as they are getting their nucleotide precursors from the medium not from de novo synthesis. Maybe we could simulate nucleotide starvation another way (slowing polymerase with an antibiotic resistance mutation? as we suggested in one of our grant proposals???).

Done and to do

Yesterday I did a big time-course experiment with the ppdA::lacZ fusion strain, to get a more detailed picture of the results in my last post. The information is now in my notebook: 50+ samples, each with sampling time and OD600 (cell density at that time) and volume assayed and assay time and OD420 (ONPG hydrolyzed by beta-galactosidase), but I haven't yet entered it into Excel and analyzed it.

The next important experiment is to demonstrate that the changes in expression are dependent on both CRP and Sxy. I will do this by doing simplified time-course analyses using cells carrying the ppdA fusion plasmid and a knockout of either crp or sxy. I grew up both these strains yesterday and hope to have time to test them this afternoon. If the changes are indeed due to increased activity of the ppdA CRP-S promoter, they shouldn't happen in the knockout backgrounds. I won't do this experiment until after I have analyzed yesterday's data, so I'll know which parts of the time course are most informative.

I didn't have time yesterday to repeat the transfer-to-starvation-medium analysis; I'll wait till I have the knockout results before doing this. That way I'll be sure that the expression changesI'm seeing are indeed due to the cause I want to investagate (changes in activity of CRP-S promoters).

Biocurious about tweezers

In between time courses I've been reading a M.Sc. thesis by the student who was developing the laser-tweezers analysis of DNA uptake. (You may know him as PhilipJ of Biocurious.)

I'm not an examiner of this thesis; I'm reading it to learn more about how laser-tweezers work. It's very well written, and pitched at exactly the right level for a neophyte like me. Any day now I should decide I've taken the time courses far enough for now, and get back to my attempt to follow in Philip's footsteps.

Time course of ppdA expression

On Saturday I did a time course examining whether the expression of the ppdA::lacZ fusion (and I hope thus of the sxy gene) changed as growth conditions changed while the cells were growing in rich medium (LB) and after transfer to a starvation medium.

The green line shows how the cells grew. First the density fell sharply because I diluted the cells 1/250 in fresh LB (+Amp to maintain the plasmid carrying the fusion). After a brief period of slow growth while the cells adjusted their metabolism to the improved conditions ("lag" phase), the cells grew exponentially, doubling about every 30 minutes ("log" phase). After about 220 minutes growth slowed down as the medium became depleted of nutrients (still growing so not yet in "stationary" phase).

The blue line in the second graph shows the expression of the lacZ fusion in LB. The first sample was from the dense culture (it had been left on my bench overnight). Once those cells were diluted into fresh LB the amount of beta-gal activity slowly fell, presumably because expression of the lacZ fusion decreased. Part of the reason the decrease is slow is that the cells still contain beta-galactosidase that will either be degraded or diluted out by cell growth. The amount of beta-gal activity remained low while the cells grew, until the cell density got quite high (OD600 about 1.0), when the activity began to increase. I know that the low activity reflects lacZ expression by cells in exponential growth, and not residual enzyme from earlier induction, because I diluted part of the log-phase culture 100-fold into fresh LB+Amp, and found the same activity after these cells had spent an additional 2 and 3 hours in exponential growth (blue circles).

But once the cells got so dense that growth slowed, the beta-gal activity increased, and by 300 minutes had become at least as high as that of the overnight cells from my bench. This is a weaker version of the gene induction we see as Haemophilus influenzae approaches stationary phase.

In H. influenzae, rapid transfer to a medium lacking most nutrients causes strong induction of CRP-S genes, including the homolog of ppdA. To mimic this I transfered the log-phase cells to a medium consisting of minimal salts (M9) plus the same small amount of amino acids we use for H. influenzae. The two red bars show that this transfer caused a more dramatic increase in beta-gal activity.

So, these are weak but nice results. They suggest that CRP-S genes in E. coli are regulated by similar factors to those regulating CRP-S genes in H. influenzae.

[Just to check, I also induced lacZ expression in wildtype (lac+) cells by adding IPTG. Even though these cells have only one copy of the lacZ gene (in the chromosome, rather than one on each of many copies of a plasmid), after 60 minutes they expressed 7 times as much beta-gal activity as the starved ppdA cells did after 120 minutes. This is slightly less than the ppdA cells expressed after Sxy was overexpressed from the pASKA plasmid, and suggests that the CRP-S promoter is being only partially induced by my starvation conditions.]

What next? I need to repeat this time course, starting with a proper overnight culture and taking more time points, especially after log phase. I need to get a longer log-phase series too. And a time course in the starvation medium.

Which E. coli is best?

I haven't yet tested whether different growth/non-growth conditions alter the expression of the ppdA fusion (though I did get the sxy manuscript resubmitted today), but a paper I came across reminded me of another issue I need to consider.

The paper is an opinion piece titled "Laboratory strains of Escherichia coli: model citizens or deceitful delinquents growing old disgracefully?"; it just came out in the journal Molecular Microbiology (Mol. Micb. 64:881-885). The authors argue that the standard K-12-derived strains of E. coli that microbiologists and molecular biologists typically use are not at all representative of the strains out in the natural environment. For one thing, lab strains have lost about 20% of their genomes. These are (by definition) non-essential genes, but their loss no doubt affects cellular metabolism, so that the metabolic interactions we see in K-12 strains may be quite different than those in natural strains. Furthermore, the culture conditions we use (rich broth, lots of oxygen, no competitors) are unlikely to ever occur in nature. Their long maintenance under these and other unnatural conditions means that the lab strains will have evolved by accumulating cryptic mutations that are beneficial under lab culture conditions but that may have very different and perhaps harmful effects in the natural environment.

I already knew this. It has important implications for my search for conditions that induce expression of competence genes in E. coli. Right now I'm working with standard lab strains, but it's all too possible that one of their ancestors lost the ability to express the genes encoding homologs of H. influenzae's competence genes, or to assemble the proteins into functional DNA uptake machinery.

So I should perform my tests on a less lab-adapted, more 'natural' strain as well as on the K-12 strain the ppdA::lacZ fusion is in. But this raises two problems. First, which strain should I use? I do have a fairly-ancestral K-12 strain, but really I should use a 'wild' strain recently isolated from the environment. The NCBI Microbial Genomes page lists 8 completely sequenced E. coli genomes; their sizes range from 4.6 million bp (K-12) to 5.6 million bp (O157:H7). And the various wild strains have genomes that are quite different from each other - does this mean I would need to test many strains before giving up? I'm also not meticulous enough to be trusted with a strain that's seriously pathogenic to humans, so the two O157:H7 strains are out, as is the uropathogenic strain*.

Second, I'll need to transfer the necessary genes into this strain; first a lacZ mutation making it Lac-, then the ppdA::lacZ fusion so I can assay induction by Sxy. This might mean that I need to get my P1 transductions working after all (I had been thinking I could let them slide now that I've found that the ppdA::lacZ fusion can be used as an indicator of CRP-S induction by Sxy). I don't need P1 to move in the ppdA fusion because it's on a plasmid, but transduction would certainly be the easiest way to move a lac- mutation in. If I'm lucky, maybe some of the wild strains are naturally Lac-. (Probably not; most screens for wild E. coli start by treating anything that's Lac- as not E. coli.)

* NCBI also lists 6 sequenced 'Shigella' genomes - we now know that the bacterial strains assigned to the genus Shigella are really variants of E. coli. But I have absolutely no intention of working with these very pathogenic bacteria.

Manuscripts progress

Last week I finally submitted our manuscript about how CRP acts at CRP-S sites in H. influenzae and E. coli. The most interesting result in it was obtained by the gsnpiw just before he left for Belize. He showed that CRP-S sites contain regulatory sequences just upstream of the FCRP-S site that are needed for transcriptional activation; these sequences aren't typically present in CRP-N promoters.
Still to resubmit is our manuscript about Sxy. It too has been enhanced by a new result from the gsnpiw. The experiments are described here). He only had time to do it once before he left, so we're not treating them as part of the paper's Results section, but the results are sufficiently good to be briefly described in the Discussion section. They show directly that mutations in sxy do alter the ability of the sxy mRNA to be translated. Two of the reviewers had suggested we do a different experiment ("toeprinting") which would have tested whether the mutant mRNAs differ in their ability to serve as templates for a polymerase. In my cover letter to the editor of our manuscript I'll explain why we think our new experiments are more informative than toeprinting would have been. I'll also explain that they're not ready to be part of the Results, and that, if the editor thinks that the experiments need to be completed and included in the Results, we'd like a two-month extension of our revisions deadline so they can be completed after the gsnpiw returns from Belize.

I spent much of yesterday making changes to the manuscript and figures and writing responses to the many points raised by the reviewers. A downside of getting four thorough reviews of a manuscript is the very large number of issues they raised. I'm not complaining, as almost all of of these issues lead to improvements; either the reviewer is right, and we make the suggested change, or the reviewer misunderstood what we meant, and we clarify our writing to prevent the misunderstanding.

Only a few points remain to be dealt with. We used lacZ fusions to examine the effect of sxy secondary structure, and one reviewer wants more background information about the behaviour of the reference fusion. This data is in the PhD thesis of a former grad student (an author on the paper), so we may be able to simply refer to it there rather than adding the data to the manuscript's Results.

Time course and IPTG dependence

I've finished the two experiments I described in the last post: (1) a time course to find out how long it takes for induction of sxy by IPTG to give production of beta-galactosidase by the ppdA::lacZ gene fusion, and (2) a 'dose-response curve' to see how much IPTG is needed to induce the fusion. And I'm posting the resulting graphs, along with a sketch of the E. coli cells I'm using for this experiment.

The sketch shows that the cells contain two plasmids. The one on the left is pASKAsxy; it carries the E. coli sxy gene (red) under the control of the Plac promoter (blue arrow). This promoter is normally OFF in these cells so the sxy gene is not expressed, but the promoter can be activated by adding a lactose-analog called IPTG. The plasmid on the right carries the lacZ gene, which codes for the enzyme beta-galactosidase. Here the lacZ gene has been placed under the control of the ppdA gene's promoter, so when ppdA would be expressed, the cells make beta-galactosidase. Beta-galactosidase normally digests lactose, but I can easily detect it because it also digests another lactose analog called ONPG, producing a bright yellow chemical (ONP).

So I give these cells IPTG, wait a while, give them ONPG, and measure how much yellow colour they have made. That's what I did yesterday. (Measuring the yellow colour was complicated by a problem with our spectrophotometer - it kept forgetting what the standard brightness was - but I managed to keep it working well enough to get my data.)

Here's the time course. I set up three cultures. One got no IPTG, one got the standard amount of IPTG (1.0 mM) and one got tenfold less (0.1 mM). It's easy to see that tenfold less IPTG gave just as strong a response as the standard amount, that neither dose produced a significant increase in beta-gal until at least 40 minutes had elapsed, and that there was a bit more beta-gal at 90 minutes than at 120 minutes.

So I have the most basic information I need. When I go on to test induction of the ppdA promoter by treating cells in ways that I think might induce expression of the normal chromosomal sxy gene, I now know that I should allow at least an hour to see the effect of my treatment.

There are no error bars because I only tested each combination of dose and time once. I'll need to repeat the experiment with more replicates to get publishable results. I'll probably also want to use more time points in the interval between 40 minutes and 90 minutes, and to try a denser culture as well as the quite dilute one I used this time.

It is worth repeating this experiment and getting some more details, because I find this long delay quite surprising. One control I didn't do is to examine IPTG induction of the normal lac operon (the lac promoter driving expression of the lacZ, lacY and lacA genes), but this is a very standard experiment done in most introductory biochemistry lab courses, and as I recall the ONPG starts to accumulate within a few minutes. A Google search just turned up a paper about mutations that affect transcription, which used IPTG induction of lacZ as an assay; their control showed that beta-gal began to accumulate after 2.3 minutes. If we assume that induction of the pASKAsxy promoter by IPTG starts producing Sxy by 2.3 minutes, why does it take so long for Sxy to induce the ppdA promoter and cause beta-gal to start accumulating? Is the sxy mRNA translated very inefficiently, so accumulation of enough Sxy protein takes a long time? Is the ppdA promoter activated only once a lot of Sxy has accumulated? The results may have implications for competence induction in H. influenzae too. We've found that cells need about 45 minutes to become competent, but I have been assuming (perhaps wrongly) that this time is mainly needed for assembly of the uptake machinery. We have some fusions of lacZ to H. influenzae competence gene promoters; I (or someone) should test the kinetics of induction of these.

Here are the results from the dose-response analysis. In this graph the X-axis is IPTG concentration, not time (and note the log scale on both axes). IPTG was added at time 0. We see that any IPTG concentration above 10 micromolar gives full induction (about 40-fold) after 120 minutes, and that even the highest concentration doesn't give any induction at 25 minutes. This may not be very useful information; because I don't have an easy to measure Sxy production directly, it may just tell me about the sensitivity of the lac repressor to IPTG.

So what next? I'm going to grow the cells containing the ppdA::lacZ plasmid (no pASKAsxy) and try the same changes to culture conditions I previously tried with the ppdD::lacZ fusion (which I now know couldn't have worked because it isn't inducible by Sxy).

What next (turning on Sxy in E. coli)?

OK, the fusion of the ppdA promoter to lacZ produces 100-fold more beta-gal when Sxy is overexpressed. What are the important things to do with it?

1. Find out how sensitive the ppdA promoter is to Sxy: The pASKAsxy plasmid produces a great deal of Sxy when its promoter is induced with IPTG, but we know that most of this Sxy aggregates into insoluble and probably nonfunctional 'inclusion bodies'. So I could try inducing with decreasing concentrations of IPTG, and measure the effect on beta-gal production. It may be that only very slight induction of Sxy will give a big induction of the ppdA promoter. Ideally the amount of Sxy would be measured too, but we don't (yet) have the antibody to E. coli Sxy that would let us easily make these measurements. The beta-gal vs IPTG assay will be easy. If I find that ppdA induction needs only a tiny bit of IPTG, I'll have more confidence that natural induction will need only modest induction of Sxy expression from its normal promoter.

2. I should also do a time course of induction, so I know how quickly the steps happen (IPTG causes sxy transcription, Sxy protein is made, Sxy causes transcription of ppdA::lacZ, beta-gal is made). I can do this with the standard (1 mM) IPTG concentration I used yesterday. Yesterday I waited 2 hours after adding IPTG before assaying beta-gal; now I'll try 1 min, 2 min, 5 min, 10 min, 30 min, 60 min, 90 min, 120 min. The results will let me optimize the assay conditions I'll use when looking for culture conditions that induce Sxy expression from its normal promoter.

What culture conditions should I test first? My previous finding that the baseline expression of this fusion is independent of Sxy and CRP means that I shouldn't bother looking for conditions that reduce expression, so I won't try adding glucose to the medium to turn off CRP. I will try adding cAMP to the medium to make sure CRP is fully active. I will do a time course, following growth of the culture in LB, looking especially at expression as the cells' growth slows at high cell density. I will transfer the cells from LB to minimal salts (with a small amount of amino acids) to mimic the inducing effect of transferring H. influenzae from rich medium to MIV. Maybe I'll try growing the E. coli in BHI before such a transfer, as BHI is a much richer medium than LB. Perhaps I should get an E. coli mutant that can't synthesize purines (or pyrimidines), to see if transferring it to minimal gives induction.

Other things I might do:

Should I try to get a strain carrying this fusion in the chromosome rather than on a plasmid? This would eliminate concern about possible variations in the number of copies of the plasmid, due to differences in culture conditions. And if the chromosomal insertion was stable I could eliminate the selective antibiotic from the test cultures. This sounds like a job for recombineering - I wonder if we can still get this working.

Should I sequence the ppdD::lacZ fusion and the hofM::lacZ fusion to see if they have mutations that would explain their failure to be induced by Sxy?

Results of the obvious next step

As I posted a few days ago, I planned to introduce a sxy-expression plasmid into my cells with CRP-S promoter fusions to lacZ, to see if the CRP-S promoters are indeed induced by Sxy. I did that yesterday, and I just finished calculating the amounts of beta-galactosidase activity these cells produced with and without induction of sxy expression.

Results: Of the three fusions I tested (all that I have), one produced almost 100-fold more beta-galactosidase when Sxy was present, one produced only slightly more, and one produced even less! The exclamation mark is because I expected them all to respond similarly.

The fusion that was induced by Sxy is to the ppdA promoter (really the promoter of a 4-gene operon). The H. influenzae homologs (not pilB but comNOPQ) are in a similar operon, which is very strongly induced in competent cells and strongly dependent on CRP and Sxy. The ComN and ComO proteins are known to be required for competence. This fusion is carried on a plasmid. The 100-fold induction when Sxy is overexpressed is what I was hoping to see. This means that the low expression when Sxy is not induced is indeed baseline, so I'm comfortable with this baseline expression not being dependent on Sxy or CRP.

The results with the other fusions are surprising. The one that produced only slightly more is a plasmid-borne fusion of lacZ to the hofM promoter (really the promoter of the hofMNOPQ operon). This is homologous to H. influenzae's comABCDE operon, and like comNOPQ it is required for competence and needs CRP and Sxy for induction. Its CRP-S promoter element doesn't look significantly worse than that of ppdA, but it was induced only three-fold by Sxy overexpression.

The fusion that produced even less beta-galactosidase when Sxy was induced is to the ppdD promoter; it's integrated into the chromosome rather than being carried on a plasmid. This is the most surprising result, as the grad-student-now-postdoc-in-waiting ('gsnpiw') used quantitative PCR to show that ppdD mRNA is induced about 100-fold by overexpression of Sxy from the same plasmid I just used. Induction of Sxy in the fusion-carrying cells not only failed to induce the ppdD::lacZ fusion, it actually made these cells quite sick; the cells with IPTG grew to less than half the density of the uninduced cells, and produced only about half as much beta-galactosidase per cell. Perhaps the chromosomal insertion in these cells has somehow made them sensitive to transcriptional activation of the ppdA gene.

Anyway, the good news is that I can use the ppdA fusion as an indicator of Sxy expression, in my search for conditions that induce Sxy and thus induce expression of CRP-S genes. I'll set aside (for now) the questions about why the other fusions don't behave similarly.

(Sorry I've been too lazy to include figures in my recent blog posts; I promise to improve.)

Cryptic effects of antibiotics and resistance alleles

This morning we had a meeting with members of another lab to discuss progress on a shared project. There isn't as much progress as I had hoped, but at least we now know what still needs to be done and who will do it.

The goal is to understand how gene expression changes when cells are exposed to antibiotics at concentrations so low they don't even slow growth of the cells, much less kill them ("sub-inhibitory concentrations"; abbreviated sub-MIC).

The first part of the project was to use microarray analysis to compare the amount of mRNA produced by each gene, when cells were grown with and without sub-MIC of the antibiotics rifampicin and erythromycin. Rifampicin inhibits production of mRNA by RNA polymerase, and erythromycin inhibits production of protein by ribosomes. This work was begun by a previous technician in our lab, and completed by an undergraduate working in the other lab. (The undergraduate just learned that she's been accepted into medical school here - Congratulations, Wendy!) I have a draft version of a conference poster with some results of this analysis, but we'll need to reanalyze the data in preparation for writing the paper we plan.

The complementary part of the project turns out to be still quite a long ways from completion. The plan is to compare the gene expression by normal (antibiotic-sensitive) cells growing without antibiotic to the expression by cells that carry a mutation making them resistant to the antibiotic (i.e. of a RifR strain and of an EryR strain). This will let us compare the set of gene-expression changes caused by resistance mutations to those caused by the antibiotic. We expect these two sets of changes to be quite different, because each represents a complex outcome of different adaptive and accidental responses to a different change.

Part of our lab's contribution was to isolate the necessary resistance mutations; that's done and we have sequenced the altered DNA so we know exactly what the changes are. The Rifampicin resistance mutation is in the rpoB gene; it creates a S->P amino acid substitution at position 509. An identical mutation is known to cause Rif resistance in Staphylococcus aureus. Isolating the erythromycin resistant strain was a lot more trouble, but it's sequenced too. It's in the L22 protein (part of the 'large' subunit of the ribosome; I forgot to note down the exact position).

Our collaborators have analyzed mRNA from the EryR strain, but unfortunately the cells were being grown in the presence of erythromycin rather than in antibiotic-free medium, so we can't use this analysis as we planned. I offered to make more mRNA, this time from cells grown without antibiotic, so the microarrays can be repeated.

Several other problems were discovered.

First, the array slides used for this analysis were quite old and had been stored in air at room temperature, which causes degradation of the DNA fragments spotted on them. When repeating the analysis we will need to use new arrays, and these must be ordered from the research group in London that makes them.

Second, the very expensive license (>$4000 per year) for the GeneSpring software used to analyze microarray results has expired. It belonged to another lab that had kindly let our collaborators use it. Luckily another lab in our group of labs is thought to have just purchased a new license, so we're going to approach the head of that lab to ask if we can use their software (on their computer, as it can't be copied to other computers) in exchange for a small financial consideration.

Alternatives to GeneSpring exist, and I think some are open access, but I suspect they require quite a bit more sophistication to use well. A Google search for "alternatives to GeneSpring" led me to BRB Array Tools, which is free from NIH and runs as an Excel add-in. It was developed by 'professional statisticians' (why do I not find this reassuring?). One strength of GeneSpring is its ability to integrate the array information with the genome sequence and metabolic pathways of the organism - this is especially valuable for simple compact genomes such as H. influenzae's.

A third problem is that neither research group (ours or our collaborators) has anyone with much experience with GeneSpring (much less any equivalent free software). I've used it (a few years ago), but I've forgotten most of what I learned. Luckily we don't need to use its more sophisticated abilities, just the basic analyses, but even so it's going to take a major investment of time to analyze the data once we get it all.

The obvious next step

I did a sloppy lab meeting presentation this afternoon on my inconclusive results so far with the lacZ fusion strains. We discussed whether I could just use the moderate constitutive expression from the CRP-S promoters as the baseline, and look for conditions that increase it (and test that the induction does depend on Sxy and CRP).

One of the post docs reminded me that we have a plasmid carrying an inducible version of the sxy gene, and made the excellent suggestion that I should introduce this plasmid into these strains and see if overexpressing sxy causes a dramatic increase in beta-galactosidase activity. If not, then the system is unlikely to be much use. But if it does (and it should) then I'll know the range of signal I can expect, and can get to work testing possibly-inducing conditions.

She already has a stock of the plasmid, so I just need to make the recipient cells competent and transform this plasmid in. I'll test both the strain with the chromosomal fusion and the two strains carrying the plasmid-borne fusions. She says that the origin of replication of the sxy-expression plasmid is compatible with (i.e. different from) the origin of the fusion plasmids.

Where'd the regulation go?

I wanted to find out why the plasmids with fusions of CRP-S promoters to lacZ (hofM::lacZ and ppdA::lacZ) give such high lacZ expression. Because CRP-S promoters require both CRP and Sxy for high activity, I transformed the plasmids into strains with crp or sxy knocked out. But I also needed crp+ sxy+ cells of the same genetic background (frozen as RR1321; I forget its original number).

So yesterday I made cells of this strain 'chemically competent' to take up plasmids by incubating them in a cold solution containing rubidium chloride. (The cells didn't become 'naturally competent' - that's the long-term goal of these experiments.) I scaled down the cumbersome procedure described in the methods manual. This made it much faster because the cells could be collected by filtration and further concentrated by microcentrifugation, rather than using a big slow centrifuge. I wasn't sure this would work well but it did - this morning I found about about 100,000 transformants on my ampicillin plates).

So now I had all the strains I needed to test whether expression of the fusion depends on CRP and Sxy. Here's the beta-galactosidase 'activity' produced by each strain (in 'Miller units'):

Parent with no plasmid: 3 units
Parent plus either plasmid: 194 and 230 units
crp knockout plus plasmid: 301 and 406 units
sxy knockout plus plasmids: 361 and 463 units.

The differences between the various plasmid-containing cultures are probably not significant, as in this quick experiment I didn't take pains to make sure the cultures were all at exactly the same growth stage. This is unlikely to be important because my previous analyses found no little dependence on growth stage.

The conclusion? Expression of these fusions is independent of both CRP and Sxy. This is a troublesome result. The most likely explanation is that the inserts that carry the CRP-S promoters also contain quite a bit of sequence upstream of the promoter, and these sequences may be responsible for the high basal expression.

I will first repeat the experiment, this time having all the cultures at the same growth stage. Then I'll email the researchers who kindly gave us these plasmids to ask if they found high basal expression too. And I'll start looking the sequences of the promoter-containing inserts, in preparation for engineering them to remove extraneous sequences.

Transformations (good) and transductions (bad)

My transformations worked. The crp knockout cells transformed with the ppd::lacZ or hofM::lacZ fusion plasmids gave me about 200 AmpR KanR colonies each, which should all contain the appropriate plasmid because the no-DNA control gave no colonies. The sxy knockout cells must have not been very competent, as they gave only 1 and 2 transformants with the two plasmids, but the no-DNA control gave none so these too are probably right.

I've streaked all the sxy knockout transformants and 3 of each of the crp knockout transformants. Before I go home (if the new colonies are big enough to see), I'll inoculate them into LB+Amp+Kan, so that tomorrow I can (1) do plasmid minipreps to check that they have the right plasmids, and (2) do beta-galactosidase assays to measure promoter activity in the mutant backgrounds.

I realized this morning that the proper comparison for these strains should not be with the plasmid-carrying strains I already tested but with plasmid-carrying strains that have the same genetic background as the knockouts. We do have the parental strain of the knockouts, so I've streaked it out and tomorrow will make it competent and transform it with the plasmids.

I was growing cells and pouring plates and making P1 lysates today, in preparation for various carefully done transductions, but discovered that the LB broth I was using for today's cultures is contaminated. So I'll need to start this over with fresh medium - probably not for a few days, as two manuscripts are urgently in need of attention. Both need to be completed within the week, as the senior author is about to leave for two months of ecological R&R in Belize. One is the sxy manuscript - data is still being generated and we have four reviewers' comments to address. The other is the manuscript describing what Sxy does at CRP-S promoters. Again data is still being generated, but it's otherwise almost ready to submit.

Final revisions of the sxy manuscript

The sxy manuscript we submitted in late February came back a month or so later with four (!) thorough and quite favourable reviews. (The exclamation point is because most papers get two reviews.) We're now doing the revisions, and hope to resubmit later this week.

It's taken us two months to get to this point because two of the reviewers asked that we improve the evidence for the role of secondary structure as a regulator of sxy mRNA translation by doing an analysis called "toeprinting". The name is by analogy with "footprinting", where a DNA-binding (or RNA-binding) protein protects a specific segment of DNA (or RNA) from cleavage by a nuclease. In toeprinting, an RNA-binding protein or other obstacle is detected as a position where a polymerase stalls while copying the mRNA.

The PhD student working on sxy regulation decided that a different technique would be more appropriate, so he's directly measured the translatability of wildtype and mutant sxy mRNAs in an in vitro system. The experiments were delayed while he finished and very capably defended his thesis, so now he's finishing the experiments as his first postdoctoral work. He's also revising the figures as requested by various of the reviewers, and has already done some of the rewriting as part of his thesis. So tomorrow we'll sit down and see how much we can finalize.

P1 lysates bad but alternatives good

Most of the old P1 lysates I had left on my bench were dead, though one was OK and the ones in the fridge were still pretty good. So I made fresh lysates yesterday on the wildtype strain W3110, but the titers are lower than I expected, and maybe not even high enough to give acceptable transduction.

So I'm going to do one more try. I found what looks to be an excellent protocol on the Open Wetware pages. I'll follow their instructions exactly (no more "I'm so smart I can safely cut corners"). I'll also try the lysate I generated yesterday, and make fresh lactose plates for the selection. Two days ago I streaked the donor and recipient strains on lactose plates to check how the donor and recipient grow. The donor grows well, giving small colonies after 24 hrs and big ones after 48 hrs. The recipient does grow a bit on lactose, giving tiny colonies after 48 hours, presumably because it lacks the lactose uptake protein but can grow very slowly on lactose that leaks in to the cell. These tiny colonies are what I saw in my last transduction.

Remember why I'm doing these transductions? I want to combine the ppdD::lacZ fusion (indicator of CRP-S promoter activity and thus of Sxy and CRP activity) with a sxy knockout and with a crp knockout to find out whether the high constitutive expression of the fusion depends on its CRP-S promoter. And I'm using the ppdD::lacZ fusion because I want to find out how to induce Sxy activity and CRP-S promoter activity in E. coli. If I find that the constitutive activity is dependent on Sxy and CRP, I will suspect that such promoters are normally moderately active. If the activity is not Sxy and CRP dependent, I will conclude that the fusion is not a good indicator and not work with it any more.

In Tuesday's post I described checking some plasmids, to see that they had the expected inserts. We asked for these plasmids (gift from other researchers) because they too contain fusions of CRP-S promoters to lacZ, and can be used in the same way as the ppdD chromosomal fusion. The two promoters are from the ppdA gene and the hofM gene, which are both homologs of CRP-S genes that H. influenzae requires for DNA uptake.

So yesterday I compared the amount of beta-galactosidase (product of lacZ) produced by cells carrying these plasmids to that produced by the ppdD fusion. Even though cells have many more copies of the plasmids than of a chromosomal fusion, they produced quite a bit less beta-galactosidase. I don't need to use P1 transduction to check whether this expression is dependent on Sxy and CRP. Because I already have preps of the plasmids, and I have strains carrying the sxy and crp knockouts, I can just transform the plasmids into the knockout strains, selecting for the ampicillin resistance of the plasmids and the kanamycin resistance of the knockouts. Even better, one of the post-docs has offered me some already-competent cells of the crp knockout.

P1

Yesterday I finally sat down at the bench and wrote up the results of my first round of P1 transductions. Three transductions, three failures.

Two of the transductions were to construct double-mutant E. coli strains carrying both the ppdD::lacZ fusion (reporter for activity of the CRP-S promoters, and thus of Sxy) and either a sxy::kan knockout or a crp::kan knockout. The goal is to test whether the high baseline expression of the reporter is due to Sxy-dependent CRP-dependent activity of the CRP-S promoter, or to baseline promoter-independent expression (e.g. from other sequences in the fusion).

Both of these transductions produced lots of colonies on the Amp+Kan selective plates. But one of the negative controls (same cells, but no P1) produced just as many, suggesting that something other than transduction was responsible for the colonies. As controls for the selection I had streaked the reporter and knockout cells onto the same plates, and onto Amp or Kan plates. These told me that the Amp+Kan plates were faulty, perhaps because the Amp was old.

I would have concluded that I should just do it again, with fresh plates, but my other transduction was a positive control for transduction (one that I knew should work), and it didn't work. This experiment tried to transduce the lacY gene from the wildtype strain W3110 into the lacY mutant C600, selecting for Lac+ by plating on minimal salts with lactose as the only sugar. The negative control was C600 with no P1. The positive control of plating the cells on minimal salts plus glucose worked well - everyone grew fine. I can't remember whether I also plated the donor cells on minimal lactose - this would have been a good control. Colonies on the lactose plates were initially very tiny, and the negative control (no phage) cells gave just as many as did the cells with phage (in fact more, probably because the phage killed some cells). So there's no evidence that this positive-control transduction worked either.

What to do next? Check growth of W3110 on the lactose plates? Repeat the lacY transduction? First I must recheck the titers of my supposedly-transducing lysates (plated last night). And go back over my notes, looking for any corners I might have cut. This is just another example of the truth of my favourite saying:

"Most scientists spend most of their time trying to figure out why their experiments won't work."

Plasmid checking

Today I made it to the bench, to do plasmid preps to check that some gift plasmids had the expected structure. I needed to check this before freezing stocks of them in our lab strain collection. The plasmids contain fusions of E. coli CRP-S promoters to the E. coli lacZYA genes. I'll be using them as reporters to test my attempts to induce expression of the E. coli sxy gene. The genes are hofM (the E. coli homolog of H. influenzae's comA) and ppdA (the E. coli homolog of H. influenzae's pilB)

So I did minipreps using our nice Sigma kit, and digested the plasmids with a pair of enzymes that would give one of two patterns. I didn't know which of three restriction sites the inserts were in, so I could only predict two possible patterns from my digests (either one 11kb fragment, one 1.2kb fragment and tiny fragments of either ~300 or ~600 (hofM or ppdA); or one 11kb fragment and fragments of ~1.5kb and 1.8kb (hofM or ppdA).

But for the first time in my research career, I absentmindedly plugged the gel box electrodes in backwards - black cable into red connection and red into black. When I went back to check 25 minutes later, the tracking dye was running backwards out the top end of the gel. Rather than starting over, I just reversed the electrodes and let it run back the way it was supposed to go for a couple of hours, hoping that some information would be usable. Much to my surprise, the gel turned out great (the migration process must be perfectly reversible even though the DNA travelled twice through the wells). If the inserts had given the two tiny fragments they would have been lost during the backwards excursion, but luckily the cloning had generated the 1.5 and 1.8kb fragments instead.