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.

Might H. influenzae competence be phase-variable?

One of the postdocs just raised an issue I've never seriously considered. Many surface structures on bacterial cells undergo what's called "phase variation". That is, a key gene controlling the structure has evolved to have a high rate of mutations that switch it from an active allele to an inactive allele, and from the inactive allele to an active one.

By "high frequency" here I mean more often than one switch per million cell divisions. Switching is thus still a very rare event, but is much higher than the background mutation rate for normal DNA sequences. Such elevated frequencies are usually caused either by short sequence repeats that cause DNA polymerase to add or miss bases in critical positions, or by specific DNA-altering enzymes that recognize the gene.

That's the proximate cause of the variation. The ultimate cause (the evolutionary cause) is thought to be natural selection created by predators or host immune systems that recognize the surface structure and attack cells expressing it. Under such pressure, a cell that has turned the structure off will have an advantage, so cells with elevated mutation rates affecting the structure are favoured. Because the structure is strongly advantageous in the absence of external attack, selection favours cells that also have a high rate of reversion mutations that switch the structure back on. Such genes are often called "contingency loci".

Competence for DNA uptake requires expressing DNA uptake proteins on the cell surface, so it's a logical target for attack by the host immune system, and thus perhaps for phase variation. But how would we detect it? In Neisseria competence is known to be phase variable, but only because it depends on the phase-variable expression of type 4 pili, a phenotype that is easily assayed in the lab. Screening H. influenzae cells for phase variation of competence is likely to be very difficult, as our only assays are uptake of radioactive DNA and transformation to antibiotic resistance.

Rather than screening for variation, a more efficient approach is to examine the H. influenzae genome for sequences that could promote such variation, and check each for its ability to affect competence. These have been thoroughly investigated by Richard Moxon and his colleagues. They found no enzymatic switches but many short sequence repeats affecting production of complex carbohydrates on the cell surface. Now we need to carefully check whether any of these could also affect genes needed for DNA uptake.

Simplifying the cyclization experiments

One of the analyses I proposed in the DNA-uptake grant proposal is designed to find out whether uptake signal sequences are unusually flexible or bent, by testing whether presence of a USS helps short DNA sequences bend around into circles whose ends can be joined by DNA ligase.

Because competition for grants is very tight this time around, we want to be prepared to submit even better proposals in September if the ones we submitted two months ago aren't successful. This means we want to have lots more preliminary data showing that the experiments we propose will actually work. So one of the post-docs has been working to get preliminary results for this circularization experiment.

The DNA fragments should be about 200bp long; shorter ones usually can't circularize at all, and longer ones have enough flexibility that the ends readily bump into each other. Even for fragments in the right size range, exact length is critical because of the 'polarity' of the ends of the double-stranded DNA. Each 'end' is really the ends of two base-paired strands, one ending in a 3'-OH and the other in a 5'-P. When the ends do bump into each other, ligase can only join them if a 3'-OH is aligned with a 5'-P (the bond must be a 3'-5 connection, not a 3'-3' or 5'-5'). Because the two strands of the DNA wind around each other every 10.4 base pairs, the length of the DNA fragments must be approximately a multiple of this length so that the ends will meet in the right alignment. The strategy is to use PCR to synthesize fragments of the right length, and she designed two sets of primers, giving fragments of 208 and 260bp (my notes say 260 but I think I have it wrong as this seems too long, unless it's the positive control).

The ends created by the PCR process are not easy to ligate, because they have inconvenient incompatible tails, so her primers include sites for digestion by restriction enzymes. Cutting both ends of the PCR product with the same restriction enzyme will generate compatible 'sticky ends' that can base pair with each other. The base pairing will hold together any ends that do bump into each other in the right orientation until ligase can seal them together permanently.

Sounds good so far. But there's one more factor. The circularization reactions must be done at low DNA concentration to decrease the frequency of ends of different molecules bumping into each other. This intermolecular reaction has 'bimolecular' kinetics, meaning that its rate depends on the DNA concentration. In contrast, circularization has 'unimolecular' kinetics, and its rate is independent of the presence of other molecules and thus of DNA concentration. Doing the reaction at low concentration is easy (need less DNA), but detecting the results of the ligation is hard, because small amounts of DNA (linear or circular) are difficult to see in the gels used to separate the different conformations that result from the ligation.

The solution is to label the DNA fragments with 32-P, making even very small quantities easy to detect. The post-doc followed a published method for labeling DNA for these experiments, which should put a 32-P at each 3' end of each fragment. The fragments are first cut with the restriction enzyme, purified using little spin-columns, and then incubated first with a phosphatase, to remove the non-radioactive P from each 3' end, and then with the 32-P nucleotide and a kinase, to put the hot phosphate on. Then the fragments are purified again.

Initially I was concerned by the low recovery; only about 10% of the input DNA was recovered after all the intervening reactions and clean-up steps. After a bit more thought I became more concerned by the labeling reactions, mostly because I've always found phosphatases to be nasty treacherous enzymes that don't know when to stop. If the phosphatase removes more than the single terminal phosphate, the fragment will not be circularizable even if the kinase then does its job correctly. Furthermore, any fragment that the kinase misses will also not be circularizable, even if the phosphatase has behaved itself. Even if one end is processed correctly, any problem with the other end will still prevent circularization. In principle these problems can be controlled for, but any experiment-to-experiment variation will invalidate the conclusions we're hoping to achieve.

Luckily, once I started worrying about these issues I realized that we can eliminate both the recovery problem and the phosphatase/kinase problems by labeling the DNA internally rather than at its ends. So the new plan is to add a labeling step after the PCR reaction. This will be essentially one extra PCR cycle, this time with one radioactive precursor nucleotide added to the mix. The resulting DNA fragments will then only need to be digested with the restriction enzyme and cleaned up once.

When the post-doc gets back from her visit home, we'll still need to solve the problem of why the gels run so oddly, but at least we'll have enough labeled DNA to do lots of tests. The gel problem may be related to the high concentration of ligase needed in these reactions. The standard ligase stock is purchased in 50% glycerol at the low concentrations needed for cloning reactions, and the circularization reactions use so much ligase that they are about 25% glycerol. I'm hoping we'll be able to buy a high-concentration ligase stock, rather than having to make our own ligase....

no transductants??

I thought I had a lot of KanR AmpR transductants from infection of the ppdD::lacZ strain (AmpR) with the P1 lysate made on the sxy::kan strain, because lots of colonies grew overnight on the Kan+Amp plates from this infection but not from mock-infected cells. But I picked 4 colonies and streaked them on various plates (Kan, Amp, Kan+Amp, Maconkey-lac) and they mostly didn't grow. Now I suspect that some ofthe plates I used may have been too old (Amp is unstable) or otherwise problematic.

My test transduction of lacY from W3110 into C600 doesn't seem to have worked either (though I did find the TTC and get cute little red colonies). And the subsequent transduction of crp::kan into the ppdD::lacZ strain doesn't seem to have worked either.

After I carefully recheck the controls I'll redo it all with fresh plates.

Doing it right (P1)

I've been futzing around with phage P1, trying to get a high-titer lysate from a single clear (P1vir) plaque, with no success. Part of the problem is the high frequency of what I guess are revertant phage, giving larger and turbid plaques, and partly it's that I've been trying to squeeze the work in between other stuff (peer-reviewing manuscripts and a proposal, preparing my freshman biology final exam, etc.).

But today's clear, and I'm ready to roll. Last night I innoculated cultures from single colonies of the E. coli wildtype strain W3110 and three mutant strains: ppdD::lacZ, sxy::kan and crp::kan. Last night I poured lawns of W3110 with appropriate dilutions of two of my not-very-satisfactory lysates. When I could the plaques this morning I'll know the titer of these lysates, and can then use one or the other to infect the four E. coli strains. I'll do these infections in broth and maybe also on plates.

Then I'll collect the lysates and titer them all overnight on W3110 lawns. I may also do a test transduction overnight, using the W3110 lysate (before I know its titer) to transduce the lacY gene into the lacY mutant strain C600. This should give me all the information I'll need for tomorrow, when I want to infect the sxy and crp knockout mutant strains with the ppdD fusion lysate, and vice versa, and select for transductants that have both the ppdD fusion and the sxy or crp knockout. Then I can test whether the high 'baseline' expression of lacZ in the ppdD::lacZ fusion is due to high baseline activity of its CRP-S promoter, as such activity should drop dramatically if sxy or crp is knocked out.

I'll need to make some minimal-lactose plates to properly score the result of my test transduction, for which I'll need to make up and sterilize stock solutions of the amino acids threonine and leucine, the vitamin thiamine, and lactose, because C600 is thi- thr- leu- as well as lacY-. (I used this strain a lot in grad school, and its genotype is burned into my brain.) And ideally I should also make up some of the TTC we used to put into minimal plates that makes the colonies turn red and easy to see. Right now I can't remember what TTC stands for, but it will come back to me, and I think we have a big bottle of the stuff somewhere.

How to compare protein sequences?

In the last post I described an analysis that depends on comparing protein sequences. There are two different ways to do the comparison; and I need to decide which is more appropriate for our analysis.

Both methods rely on first aligning the amino acids in the proteins to be compared (here we'll only be comparing two proteins at a time), and then comparing the amino acids at each aligned position. The goal of the alignment is to align amino acids that are homologous - that is, those that are similar because of descent from the same position in the ancestral sequence both proteins evolved from. (If the proteins are not themselves homologous the analysis can't be done.)

In studies of evolutionary relationships, the usual method of comparison is to simply count how many of the positions have identical amino acids, giving a "% identity" score. In studies of protein function, scores based on the functional similarity of the aligned amino acids are often used. These rely on a matrix that gives similarity scores of all pairwise combinations of amino acids. These matrix scores are themselves derived from comparisons of large numbers of aligned amino acids, giving highest scores to amino acids which most often have evolved to perform the same role in a protein. For example, valine and leucine are commonly found in homologous positions, and matrices give this pair a high score. Wikipedia gives a good explanation of the use of matrices to compare protein sequences.

I'll post later about the issues raised by our analysis problem.

Analyzing the effect of USS on the coding function of genes

While I've been doing other things a collaborator has been working hard on a comparative genomics project that will tell us how much impact uptake signal sequences (USS) have on gene function.

Reminder: USS are short sequence motifs (the longest are ~30bp) present in many copies in the genomes of naturally transformable bacteria, probably because the cells preferentially take up DNA fragments containing the motif. Most of the USS in the Haemophilus influenzae genome are in coding sequences, and we want to find out whether their presence forces genes to specify sub-optimal amino acids at positions encoded by USS.

This analysis is testing the effect of USS by comparing the amino acid sequences of proteins with and without USS. For each H. influenzae gene with one or more USSs, we first find homologous protein sequences from at least three genomes with no USS. We compare these three protein sequences with each other (that's three no-USS comparison scores), to get a measure of how strongly selection acts on the protein, especially on the segment that in H. influenzae is specified by a USS. Then we compare each of the three with the H. influenzae sequence (that's three +USS comparison scores).

Then we compare the mean no-USS score with the mean +USS score; if the scores are similar then we conclude that the USS doesn't significantly constrain the protein's function. There's a lot of random variation, so we do this for every USS-encoded gene in the the genome and then plot each pair of scores as a point on a scatter-plot. Points that fall on a diagonal line represent genes whose USSs don't constrain them, and points that fall below the line represent genes whose USSs may be causing problems.

We're not interested in specific genes, but in the general picture - we want to know whether, on average, USSs cause problems or not. A preliminary analysis done years ago suggested they don't, but the answer from this new improved analysis will be interesting in any case.

Lysates!

Well, lysates aren't really that exciting, but it's been a while since I got to do anything with phage. I used both the turbid-plaque and the clear-plaque streaks to infect two cultures, one with the ppdD::lacZ reporter fusion and one with the sxy::kan knockout. They all grew up nicely and then promptly lysed. Titering showed that the clear-plaque phage gave about 2x10^10 pfu/ml, all the small clear plaques expected of P1vir. The other phage gave a mix of clear and turbid plaques, suggesting that the colleague who made the source lysate had neglected to start from a single plaque (naming no names).

Today I'm going to make a good stock lysate, by infecting the "wild type" E. coli strain W3110. (I put wild type in quotes because the strain does not carry the lambda phage present in the original "wild" K-12 isolate. And I'm going to do the transduction that was the point of getting a P1 lysate, to create a strain carrying both the ppdD::lacZ reporter and the sxy::kan knockout). I have lysates of both strains so I'll try the transduction in both directions, each selecting for AmpR KanR (the reporter carries AmpR). And a control transduction transferring lacY from W3110 into C600.

P1 (vir?)

I have some phage P1, courtesy of a colleague. But (as usual) complications have arisen. The colleague is out of town, and his P1 collection consisted of five different lysate tubes, of unknown ages as none of the tubes had a date. So I tested them all for viable phage by streaking a drop onto a plate that had a lawn of host cells on it, and looked this morning for the plaques (spots of lysis) that phage would make.

Two of the lysates gave no plaques, so they were probably very old. Two others (#1 and #4) gave lots of conspicuous plaques, and #2 gave tiny plaques. The colleague tells me (by email) that #1 is his newest stock, but the big plaques are not what I expected.

The phage should be not normal (wildtype) P1 but a mutant called P1vir (vir = virulent). Wildtype P1 can form lysogens, becoming dormant in cells that are then immune to being killed by other P1. The large plaques I see look like plaques made by wildtype P1, because they have cloudy centers (the plaques are "turbid"), where surviving cells are growing; these are likely to be lysogens. The virulent mutant should not form lysogens, so its plaques should be clear. Our manual of bacterial genetics methods says that P1vir plaques are tiny, so I think lysate #2 is more likely to be genuine P1vir. Lysates #1 and #4 would then have been grown from P1vir that had mutated, reverting to wildtype. (Perhaps the person who made the lysate mistakenly picked a "nice big plaque" instead of the more common tiny plaques.)

Wildtype P1 is not very suitable for doing transductions because many of the survivors are likely to be lysogens, whereas we usually want to work with phage-free cells. I'll do test transductions with lysates #1 and #2; if #2 works well I won't bother doing experiments on #1 to sort out the virulence issue, but just make a good fresh lysate from a single plaque of #2. Part of this lysate will be given to the colleague who supplied the original lysates.

Before making proper lysates and testing transduction I need to grow up appropriate E. coli host strains. I did the preliminary checking on the strain carrying the ppdD::lacZ fusion. Unfortunately the computer that we keep our strain list on is in the shop, but one of the post-docs has a recent backup.

Results of beta-galactosidase assays

So I did some beta-galactosidase assays on the strain with ppdD fused to lacZ. The immediate goal was to characterize baseline transcription of the ppdD gene, with the longer goal of finding ways to turn it up by inducing expression of sxy.

The researcher who kindly sent us the strain with this fusion said that its colonies are pale blue on plates with the beta-gal indicator X-gal, but for me they were quite a strong blue. So I wasn't too surprised that my assays showed moderate beta-gal activity in all the conditions I tested. I tested cells in exponential growth and after overnight culture in LB, in LB+glucose (which should prevent production of cAMP and thus expression of the ppdD gene's CRP-S-regulated promoter, and of cells in LB+glycerol, which should allow cAMP production like plain LB but provide the same amount of extra carbon source that glucose does. I also tested cells transferred from log-phase growth in LB+glucose to minimal salts with added amino acids ("M9+caa"); a treatment that might roughly approximate the competence-inducing effect of transferring H. influenzae cells from sBHI to MIV. None of these treatments made much difference; all the samples produced between 150 and 500 units of enzyme activity per ml of cells.

So the next test is to find out whether the transcription of ppdD depends on Sxy. If it does, then this suggests that Sxy is being produced at least a bit under the standard conditions I tested. If not, the expression is genuinely baseline, and maybe I should test other genes as indicators of sxy expression.

To do this test I need to introduce our sxy knockout into the ppdD::lacZ fusion strain. So tomorrow I'll go searching for the needed P1 lysate. The colleague who has it has unfortunately gone off to Europe for a couple of months, but he tells me that one of the people in his lab can help me find it.

Preparing for the E. coli sxy induction experiments

The strain and plasmids we've been waiting for arrived on Wednesday (thank you to the Pugsley lab), so now I can get started on my attempts to induce sxy expression in E. coli. And we do already have an E. coli sxy knockout from the Japanese group, so solving the recombineering problems isn't critical (though I should get to work on that anyway).

What to do first? The initial tests will use a reporter strain carrying a fusion of the ppdD gene to lacZ. The person who sent the strain says it should be pale blue on X-gal plates, indicating weak baseline expression of lacZ. I should first characterize its lacZ expression more precisely by growing it under defined conditions and measuring the amount of beta-galactosidase (the lacZ product) with the substrate ONPG. This is a simple classic assay, done in every introductory molecular biology lab. I'll grow cells in rich medium and minimal medium, doing the assay at various cell densities.

I need to find out whether this baseline expression is independent of the transcriptional regulators Sxy and CRP/cAMP, which we know activate the ppdD promoter. This requires transferring the ppdD::lacZ fusion into strains that carry knockouts of either sxy or crp (or cya, which encodes the adenylate cyclase that makes cAMP), or transferring the sxy and crp knockouts into the ppdD::lacZ strain. We have both these knockouts, but I don't yet have the P1 lysate I'll need to do the transfers. And I need to find out the antibiotic resistances associated with each of these knockouts, so I can plan the selections. And I need to get all the strains from the grad student who's been working with them.

I do have what I need to test whether baseline expression of the ppdD::lacZ fusion is affected by cAMP levels. The grad student tells me cAMP is normally high in the rich medium LB, presumably because LB lacks glucose, causing the phosphotransferase system to activate adenylate cyclase to make lots of cAMP. If this is correct, simply adding glucose to LB should reduce cAMP levels. So should growing cells in a minimal medium with glucose as the carbon source. If the baseline lacZ expression from the ppdD promoter is due to weak activation of the promoter by CRP + Sxy, these conditions should reduce it, and adding exogenous cAMP should restore it. If it's just constitutive activity of the promoter, these conditions should have no effect.

Negative control for E. coli sxy experiments

I'm still planning the experiments to identify conditions that induce sxy expression (and possibly competence) in E. coli.

One issue I hadn't considered previously is the appropriate negative controls. If I'm using the sxy-inducible ppdD::lacZ fusion to indicate sxy expression, the best control will be a sxy-knockout mutant carrying the same lacZ fusion. We might already have such a mutant (from the wonderful Japanese group that provides clones and knockouts of E. coli genes), but if not I'll need to make one. This would be best done using the genetic technique called recombineering, which one of the post-docs has been trying to get working for us. I'll start working with her on this.

How hot can it be?

I'm going to test biotin-tagging of the ends of my big DNA fragments by doing a preliminary tagging with a radioactive (33-P) nucleotide. I need this test because I don't have specific information about the short single-stranded overhangs I expect these fragments to have. I don't know whether most fragments have overhangs, whether 3' and 5' overhangs are equally common, or how long the average overhang is.

But I realized that the unless the overhangs are very long, they will constitute only a tiny fraction of the DNA in such long molecules. So I don't expect much of my 33-P or biotin to be incorporated. I do know the specific activity of the 33-P I'll use (2500 Curie/mmol* **), and this lets me do a back-of-the-envelope calculation of how hot the DNA can get if the tagging reaction works perfectly.

[This is a long but not difficult calculation, relying on the kind of 'dimensional analysis' I learned in Grade 11 Physics class. It has Avogadro's number and Rosie's universal constant (10^18 bp/g) and arithmetic-simplifying assumptions that the average fragment is 75kb long and that a single 33-P nucleotide gets incorporated at each end. This last assumption would be right if 3' and 5' ends were equally common, blunt ends insignificant, and the average overhang about 8 bases.]

The result of this calculation: a perfect labeling reaction could incorporate enough 33-P to give only about 240 dpm per microgram of DNA. This is so low that the 33-P labeling experiment may not be worth doing at all. Using 32-P won't make a big difference, as the standard specific activity of 32-P nucleotides is 3000 Ci/mmol.

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*A Curie is 2.2 x 10^12 dpm (radioactive disintegrations per minute).

** This is close to the theoretical maximum specific activity, with a 33-P as the alpha phosphate of every nucleotide.

DNA looks ready to label

Both my gels gave OK results. But I didn't really use enough DNA in the conventional gel, and the CHEF gel ran faster (or longer) than would have been ideal. But the conclusion is that both DNA preps consist almost entirely of fragments bigger than 40kb and smaller than 150kb. This is fine for my initial experiments.

The next step is to tag the ends of the DNA molecules with biotin. At most ends the two strands are likely to be unevenly broken (i.e. either the 5' end or the 3' end overhanging), so I can use Klenow polymerase to fill in the ends that have 3' overhangs. I'll use a biotinylated deoxynucleotide in place of done of the normal nucleotides (e.g. biotin-dUTP), so those ends whose 3' overhangs include an A will have a biotin. Ends lacking an A won't be labeled, nor will blunt ends and those with 5' overhangs. But this shouldn't be a problem as the presence of fragments with one or two unlabeled ends won't interfere with attachment of the biotin fragments to the streptavidin beads. I expect 3' overhangs and 5' overhangs to be equally common, and both to bed much more common than blunt ends. H. influenzae DNA us rich in As and Ts so, if all goes well, 30-50% of the ends will be labeled.

I can't do this until we get some biotin-dUTP; I'll order it first thing Monday morning.

To check whether the labeling reaction works, I'll do a test reaction that also includes a small amount of a radioactive nucleotide; we have some in the fridge that should still be good as it is labeled with 33-P which has a 30-day half-life. I'll both check for incorporation of 33-P into the DNA directly, and run the DNA in a gel and do an autoradiogram (expose it to a radiation-detecting screen) to check that most of the label is in big fragments. This is important because, although small fragments are less conspicuous in gels (longer fragment shave more DNA in them so they fluorescence brighter). their ends label just as well as those of long fragments fragments.

On Monday I'll also order the streptavidin-coated polystyrene beads needed for the next step.

Running gels!

I've got two gels running while I type. Both will help me decide if the MAP7 DNA preps we have are suitable for my tweezers experiments. Both gels contain high and low concentrations of two different MAP7 DNA preps, along with a size standard consisting of intact phage lambda DNA (48.5kb) and a HindIII digest of the same DNA.

The first gel is a conventional agarose gel - the voltage is created by a pair of simple wire electrodes, one running across each end of the gel box. To increase the resolution (i.e. separation) of DNA fragments bigger than 15-20kb, the gel has a lower concentration of agarose than is usually used (0.6% rather then 0.8-1.0%). This makes it more fragile, so I'll need to handle it very carefully tomorrow when I'm photographing it. I'm also using a much lower voltage, which will also help spread out the big fragments and squeeze together the little ones I don't need to resolve.

The second gel is a "CHEF" pulsed field gel. This uses pulsing electric fields in different directions (120 degrees to each other) to jiggle the DNA fragments back and forth as they move through the gel. This forward-left then forward-right pushing has little effect on the separation of small fragments (less than about 20kb) but it greatly improves the separation of the big fragments. The new apparatus (belonging to the new lab next door) is much more sophisticated than the old one that's collecting dust on our top shelf, and I had to call technical support to learn how to dumb it down enough that I could control what it was doing.

In admiring this new apparatus we developed a new rule of thumb relating the cost of a piece of scientific equipment to the number of buttons it has. This one has more than 50 buttons, and cost about $40,000; our old one had about 7 buttons and cost $4500. This only applies to equipment that doesn't come with its own computer. The real-time PCR machine we share with other labs cost $70,000; it gets away with having only a single button because it's controlled by very complicated software.

Not beads-on-a-string but strings of DNA on a bead

The other project I'm taking on is the controls and preparation for our laser tweezers analysis of DNA uptake. Our previous work (really the physics grad student's work) was trying to get cells to attach to and take up a specific bead-attached DNA fragment with a single USS. If this had worked we hoped it would let us study how DNA uptake depends on the orientation of the USS. We now realize that this was too ambitious an initial goal.

Our new experimental goals are to study the forces that act on the DNA during uptake, in wildtype cells and in cells with mutations in specific uptake proteins. This doesn't require use of a defined DNA molecule with only one USS, so we can change the setup to optimize our chances of success.

We also belatedly realized that we can use Bacillus subtilis as a positive control. Laser tweezers have already been used to study forces generated during B. subtilis DNA uptake. (Because the B. subtilis uptake mechanism is quite different than that of H. influenzae this serves as proof-of-concept for our experiments but doesn't make them redundant.) B. subtilis cells are larger and more robust than H. influenzae cells, and their DNA uptake does not depend on a particular sequence. Because the conditions for B. subtilis tweezer assays have already been worked out, we will make sure we can demonstrate uptake by it before moving on to uptake by H. influenzae.

The new plan is to first make sure we have lots of DNA fragments attached to each polystyrene bead, before trying to detect uptake by either B. subtilis or H. influenzae. My first step is to get a preparation of randomly broken H. influenzae DNA that consists mainly of fragments 50-100kb long, and then to attach these fragments to the beads by sticking biotin on the ends of the DNA, and using beads pre-coated with streptavidin, which binds tightly to biotin.

I've decided on 50-100kb for several reasons. First, it's hard to work with fragments longer than this, because longer fragments break unless they're handled very gently. Second, it's easy to get fragments about this size simply by not being gentle with the DNA prep. Third, this will let me attach a LOT of DNA to each bead. Fourth, the individual fragments will be substantially longer than the cells and the beads: the beads are 1 micrometer in diameter, the cells are 1-3 micrometers long, and a 60kb DNA fragment is about 20 micrometers. But they aren't so long that they extend a long way away from the bead they're attached to.

I'll use DNA from strain MAP7, partly because we already have lots of it (made by one of the post-docs) and partly because it carries antibiotic-resistance alleles that we can use to detect whether cells have taken it up. This will be important in checking whether the beads really do have DNA attached to them.

The DNA in the prep we have may already be broken into appropriately-sized fragments. I can check for small pieces by running this DNA in a normal agarose gel; fragments bigger than about 25kb will all jam up at the top of the gel, and only smaller fragments will spread out in the gel. A better check for DNA size will be to run the DNA in a pulsed-field agarose gel, whose rapidly reversing electrical current allows even very big fragments to spread out in the gel. Luckily our neighbours in the lab have a new apparatus to run these gels. (We have a 15-year old system which I suspect no longer works.)

If this analysis says the DNA fragments are too small, I'll make a fresh prep, handling it more gently. If the analysis shows that the DNA fragments are too big, I'll just rough up the DNA prep by whirling it in the vortex mixer, or by forcing it through a narrow syringe needle.

The next step will be adding the biotin to the ends....

(Hmm, the spell-checker thinks that tweezers must be plural; it doesn't approve of "laser tweezer analysis".)

E. coli strain construction the old-fashioned way

I haven't done a P1 transduction (or any E. coli genetics) for 20 years. But I'm being sent E. coli strains carrying chromosomal lacZ fusions and will need to move these fusions into different genetic backgrounds. This is stuff I did in grad school and I'm delighted to be doing it again.

P1 is a phage (bacteria virus). Particles of phage inject their DNA into E. coli cells, where the DNA is replicated and instructs the cell to make lots of phage proteins. Then the phage proteins self-assemble into phage particles that each fill themselves with a phage DNA, and other phage-encoded proteins cause the cell to lyse (burst open), freeing the new phage to go infect more cells.

What makes P1 useful for strain construction is its tendency to make the mistake of filling itself with a fragment of E. coli DNA rather than P1 DNA. The particles that do this still go on to inject their DNA into new cells, but this DNA (of course) doesn't make new phage or burst the cell. Instead it often recombines with the chromosome of this cell. If the original cell (the 'donor') and the new cell (the 'recipient') had different versions (alleles) of a gene, the donor cell's allele will sometimes get recombined into the recipient cell. The fragment of DNA that's transferred is about 100 genes long, so even big differences (presence or absence of whole genes) can be transferred.

Say strain A has an gene that I want to put into strain B. I first replicate P1 with strain A as host. This produces about 5 ml of a phage 'lysate', usually containing about 10^10 Pi phage per ml. Most of these are normal but some have DNA from strain A. I then infect strain B with these phage, using fewer phage than I have bacteria (I think), so most bacteria get infected by only a single phage. Most of these are infected by normal phage, and go on to lyse and release new phage. To prevent these phage from killing other cell, after the first cells have had time to inject their DNA into their hosts (10 minutes?) I add some citrate, which binds up the calcium the phage need to attach to new cells. Then I put the strain B cells onto agar plates with medium that only cells with the strain A gene can grow on, and I leave them in the incubator overnight. If I haven't messed up, the next day the plates will have lots of strain B colonies with the desired gene.

Lovely pure genetics. No enzymes, no PCR, no kits, and the cells do all the work.

HOW to turn on sxy in E. coli

We think that the previous attempts at inducing E. coli's ppdD were unsuccessful because the researchers didn't know that this gene has a CRP-S promoter that requires Sxy, and didn't know what we know about regulation of sxy expression and of competence genes in H. influenzae.

We know that competence is induced by an abrupt nutritional downshift, so I'll try to induce E. coli sxy the same way. The previous researchers tested nutritionally-limiting conditions, but only in steady state (log phase) culture. So I'll grow E. coli in rich medium (LB or sBHI) and transfer it to either MIV or the simple salts used in various E. coli defined media. I'll also test the effect on ppdD of abruptly cutting off the culture's oxygen supply, and of the onset of stationary phase.

Because we have evidence that in H. influenzae sxy induction depends on depletion of nucleotide pools, I'll try various genetic tricks that may cause E. coli to suddenly run out of nucleotides for DNA or RNA synthesis. The most straightforward is culturing purine auxotrophs in rich medium, and then abruptly transferring them to a defined medium lacking purines.

cAMP induces sxy transcription in H. influenzae, but I don't think the E. coli sxy gene has any obvious CRP site. However I'll probably add cAMP to the various test media, both because we know the CRP-S promoters need it and just in case sxy transcription does too.

Turning on sxy in E. coli

(Sorry for the infrequent posts. Teaching biology to 400 first-year students, and the associated administrative crap, has been sucking all the oxygen out of my brain.)

Once classes end (only a couple more weeks...) I'm planning research in two directions. One is the preliminary work for the laser tweezers measurements of DNA uptake. I'll post about that later. The other is searching for conditions that induce expression of the E. coli sxy gene.

In Haemophilus influenzae expression of sxy induces the genes of the CRP-S operon, and these in turn enable the cell to take up DNA. E. coli has most of the same CRP-S genes, and it has a sxy homolog. The grad student (we're down to one; the other one finished) has shown that artificial expression of E. coli sxy does induce expression of its CRP-S genes, but nobody has found conditions that naturally induce these genes. Such conditions would, we expect, be conditions that induce expression of sxy. Understanding how sxy is regulated, in both E. coli and H. influenzae, is key to understanding the evolutionary function(s) of the CRP-S regulon.

So my plan is to subject E. coli to the same kinds of conditions that induce sxy in H. influenzae. To tell when sxy has been induced, I can either use a fusion of the E. coli lacZ gene to the sxy gene, or use a fusion of the lacZ gene to the ppdD gene, which is one of the CRP-S genes that we know to be strongly induced by Sxy. Then I can easily detectg expression of lacZ, either in broth by incubation with the lactose analog ONPG (turns yellow), or on plates with the lactose analog X-GAL (turns colonies blue). The grad student has emailed the French researchers who studied ppdD induction, asking if they will send us their ppdD fusion.

Long-term plans inspire short-term plans

Now that the rush of grant-proposal preparation is over it's time to consider the immediate implications for the experiments we're doing. Because the granting agency's funds are especially tight right now, our plans need to include the possibility that one or both proposals won't be funded in this round, so that we'll need to reapply in September.

(Summaries of both proposals are available on the "What we're planning" page of our lab's web pages (link in sidebar). The complete Sxy proposal is also available there, but the DNA uptake proposal is not because I left one of the figure files at home; I'll post it tomorrow.)

So this afternoon at lab meeting we discussed the proposed DNA uptake experiments, with a particular focus on identifying the aspects of those experiments that will be seen as strong preliminary work in support of a resubmitted application. This was all too easy, because we haven't yet done nearly as much preliminary work as I'd like.

Luckily the post-doc working on uptake is happy to consider new directions (we submitted her paper on Tuesday), and I'm eager to get my hands wet again after so many months of paper and proposal writing (and teaching, though that's not done for another month).

Grant proposals done!

Both my grant proposals are done. Competition is very tight this year, and I won't learn the outcomes until sometime in July.

I think the problems we propose to address are very important, the questions we pose get to the heart of the problems, and the experiments we're proposing are excellent ways to answer them. The proposals themselves are very clearly written and nicely presented; the text is well spaced and has little colour illustrations embedded in it, and is supplemented with very clear and carefully drawn illustrations. Much of the credit goes to the grad students and post-docs, whose insights and work were critical in getting the proposals done.

The proposals would have been even better if I had been able to include more preliminary data for some of the experiments, to reassure the reviewers that the planned experiments will actually work. A related problem is the need for more discussion of what problems could arise with the proposed experiments and how such problems will be dealt with.

Now we have these great research programs worked out, we need to adjust our immediate research priorities a bit. This will have the added benefit of generating some preliminary data and a better understanding of possible problems, so if the proposals don't get funded we'll be ready to submit stronger ones in September.

Nanotechnology?

If I do an experiment that uses 50 nm magnetic beads, am I using nanotechnology? If the experiment involves a new use for these beads, am I developing nanotechnology? More generally, is all molecular biology nanotechnology?

Anyway, what am I going to do with these beads? I want to find out whether the USS polarizes the direction of DNA uptake. That is, when the USS sequence interacts with the DNA uptake machinery on the cell surface, does only the DNA on one side of the canonical orientation get pulled into the cell, at least until the end has been brought in?

My latest model of the mechanism of uptake initiation predicts that at the initiation step a type 4 pseudopilus pulls in the DNA on the right side of the USS core, while the core is bound tightly to a receptor protein on the surface (this could be the secretin pore-forming protein). If the core remains tightly bound, uptake could continue only on this side until the end of the fragment is reached. This would require that the force exerted by the pseudopilus on the DNA be less that that needed to pull the USS core free from its receptor. Bringing in the end of the DNA would allow transport across the cytoplasmic membrane, which would reverse the direction of uptake. Under this version of the model, this reversal of direction and the stronger pull exerted on the DNA by the cytoplasmic membrane machinery would detach the USS core from the receptor and bring in the DNA on the left side.

This model of uptake is much more detailed than the available evidence supports. I see this as a strength, not a weakness. The model makes many very specific and testable predictions, and at this stage of investigation it's more important for a hypothesis to be testable than to be correct. Maybe that's true at any stage.

I still like the idea of using laser tweezers to investigate the polarity question, but my physicist collaborator points out that this single-molecule method may lack the resolution we need to answer the question. Tweezers are best at measuring forces, not movement of cells.

So I'm back to thinking about blocking the ends with beads. Our previous attempts used what we now realize were giant beads (streptavidin-agarose), far bigger than the cells and much too big and porous for the task. This improved approach will use beads that are only 50 nm across, which should be plenty big enough to prevent uptake as the secretin pore is thought to be only 6-7 nm across at its widest. They can be purchased with streptavidin bound to their surface, making it easy to attach them to DNA just as we did with the agarose beads.

Our original experiments labeled one end of a short USS-containing fragment with 32P and blocked the other end with a (giant) bead. We predicted that the orientation of the the USS with respect to the bead would determine whether the 32P got inside the cell, but found that orientation had little effect. I think we should repeat these experiments using the 50 nm magnetic beads, paying careful attention to the kinetics. The original experiments used 220 bp fragments; because such short fragments might behave differently than long ones, we should also repeat the analysis with fragments that are one or a few kb long (easy - we just use the whole USS plasmid rather than its 220bp insert).

Maybe we can then combine use of the beads with the tweezer analysis, examining forces rather than distance changes as originally planned. If the tweezer analysis of the force acting on the DNA shows that the inner membrane machinery exerts a stronger force than the pseudopilus, then blocking the right end of a long fragment with a bead (left end held in the laser trap) should allow weak-force uptake but prevent the transition to strong-force uptake. Flipping the orientation of the USS should. . . . . no, the transition would still be blocked, this time by the large trapped polystyrene bead at the other end. The flipped USS orientation would affect movement of the cell rather than the kind of force.

How bacterial conjugation works

I just had a lively discussion with the guy in the next office about whether the single-stranded DNA that enters a bacterial cell by conjugation becomes double-stranded before it recombines into the chromosome. The textbook he's using to teach his new bacterial molecular genetics course say yes, but this seems rather improbable to me.

Another faculty member found a paper from about 10 years ago that concludes that the DNA becomes double-stranded before it recombines. But I don't think this double-strandedness was necessitated by the findings of their experiments; rather it seems to be mainly driven by the assumptions of the researchers.

The paper had another surprise. It showed the DNA being transferred into the recipient cell as a loop, with the 'leading' end being held back in the 'donor' cell, and the loop being pushed into the recipient. I've only seen it represented differently, with the leading end moving into the recipient cell as if it was pulling the DNA strand along behind it. But now I think more carefully, the protein attached to the leading end isn't expected to have any power to 'pull' anything once it's in the recipient cell, so the DNA must be being transported just by the machinery in the membrane of the 'donor'.

I'm putting 'donor' in quotes, because that's how most people think of the process of conjugation. They've been taught to think of the cell that has a plasmid kindly donating a copy of it to a cell that lacks the plasmid; if the plasmid is attached to chromosomal DNA the recipient cell is blessed with a copy of part of the donor's chromosome. But plasmids are genetic parasites, and the 'donation' is really an infection. The recipient is passive, and the plasmid DNA is forcibly inserted into it by the 'donor'. To do this the plasmid uses the proteins it codes for, and proteins already produced by its cell.

Why is H. influenzae's CRP so feeble?

One discovery from the grad student's work on Sxy and CRP is that the H. influenzae CRP protein binds CRP sites much less strongly (with much lower affinity) than the E. coli CRP protein does. This is a bit surprising. The two proteins have quite similar sequences, and all the amino acid residues expected to directly contact the DNA are identical.

He's going to contact a lab that has done extensive structural analysis of E. coli CRP, to see how difficult it would be to see how well H. influenzae CRP will superimpose on the E. coli structure. One possibility he suggested is that the dimerization domain of H. influenzae CRP could be weak. This would cause the protein to spend less time assembled into the dimers that most readily bind DNA.

Until now we (at least I) had thought that the affinity of CRP for different genes was determined by how well the gene's CRP site matched the protein's requirements for binding. This would have been optimized for each gene by natural selection acting on mutations in its CRP site. But now I'm wondering whether natural selection has also acted differently on E. coli and H. influenzae CRP proteins to fine tune their affinity for all the sites in their respective genomes.

H. influenzae has only about 40% as many genes as E. coli, and about 40% as many CRP sites regulating them. But I can't think of any way that would favour a 100-fold difference in CRP affinity for the same CRP site, which is what the grad student has found.

The assays were done under exactly the same conditions, but this doesn't ensure that the proteins responded identically to these conditions. I wonder if the binding conditions used for these measurements (optimized for E. coli CRP) might be unsuitable for H. influenzae CRP.

What part of "match" don't we understand?

The overall goal of the work the post-doc and I describe in our Defining the USS manuscript is to find out whether the sequence specificity of the H. influenzae DNA uptake machinery matches the consensus of the uptake signal sequence (USS) repeats in its genome. We've done good jobs of characterizing both the uptake specificity and the USS consensus, but now we're discovering that comparing them is trickier than we had realized.

The figure shows both results, with the genome consensus shown as a SequenceLogo at the bottom, and the effect on uptake of changing each position shown by the bar chart above. Because all the positions in the core (the AAGTGCGGT at the left) have a very strong consensus in the genome, we expected that changing any of these positions would dramatically decrease uptake. But instead we see that changes to different core positions have very different effects. Changing the first position doesn't decrease uptake at all (within our limit of detection) but changing any of the central three knocks it way down.

One confounding factor comes from how the genome consensus is described; the Y-axis of the logo is not a linear scale but a logarithmic scale reflecting 'information content' of the consensus. Another may come from how the uptake effects were measured; cells were deliberately given less DNA than needed to saturate the uptake machinery.

But the biggest complication is that, although we have a simple 'molecular drive' hypothesis describing how biased DNA uptake leads to accumulation of the preferred sequence in the genome, we are only beginning to develop ways to evaluate the different components of this model. This means that we can't predict exactly what sequences will accumulate in response to any specific uptake bias.

The post-doc describes the discrepancy between the two parts of the figure as maybe resulting from 'saturation' of the evolutionary process causing USS accumulation. If a small uptake bias acting over millions of generations is enough to drive the preferred base at a particular position to a very high frequency, then a larger bias may not make much difference to the outcome. For example, say USSs with a C or G or T at position 1 are taken up 95% as well as a USS with the consensus A at that position, but this bias provides sufficient long-term drive to cause 98% of the USS population to have As at that position. If so, increasing the bias might not increase the frequency of As by very much.

So what will we say in our manuscript? We could start by discussing the implications for the role of the USS in DNA uptake. We'll point out that uptake is very sensitive to the central positions of the core. Even if the other positions in the USS (core and flanking AT-rich segments) are all perfectly matched to the genome consensus, changing any one of these central bases drastically reduces uptake. This suggests that these positions make very important contacts with the uptake machinery. Changes at any of five other core positions reduce uptake to 20-60% of the control (perfect USS), so these also probably make important contacts. Changing the initial A, or pairs of bases in the flanking segments, reduces uptake only modestly, suggesting that the functions of these in uptake are dispensable if the rest of the USS is perfect.

We could go on to discuss the implications for accumulation of USSs in the genome, as I do above, and end by pointing out that what's needed is a better understanding of the consequences of molecular drive (and maybe put in a plug for our Perl simulation model).

The post-doc and I got about half-way through our Defining the USS paper today. We fixed up the Methods section and almost all of the Introduction, and walked ourselves through the Results. Maybe because we hadn't looked at them for a couple of weeks, we had some new ideas.

We had been commenting on how the USSs in coding sequences show significantly different USS motifs, depending on which reading frame they're in. But now we look again, we realize that the real story isn't how they are different, but how similar they are. Despite coding for entirely different amino acids, they all have strong matches to the canonical USS core and flanking segments. The differences are so small that they might even be attributable to random effects due to small sample sizes.

This might mean that needing to code for proteins doesn't significantly constrain USS sequences. Rather, USSs accumulate only in places where the existing protein coding constraints meet their need for uptake efficiency.

We also clarified our analysis of the correlations between the bases at different positions in the USS, especially for USSs that are likely to be acting as terminators. Because terminators act by folding into hairpins, we expected to see that the parts of these USSs that come together when folded would show correlations. But they don't. Instead the strong correlations are all between bases that are adjacent in the sequence or separated by only a single base, just as they are in the non-terminator data set. We think this means that the correlations arise from the need for adjacent base pairs to physically cooperate when the USS is kinked during uptake, and that selection for terminator function is much weaker.

Defining the uptake signal sequence

The big goal for today is to make progress on the manuscript where we pin down the specifics of the Haemophilus influenzae 'uptake signal sequence' (USS). This DNA sequence can be defined in two ways, both as the sequence motif preferentially taken up by competent H. influenzae cells, and as a sequence motif that's strikingly overabundant in the H. influenzae genome. The two are of course related; we think that the abundance on the genome is a direct consequence of preferential uptake and random (unbiased) recombination between incoming DNA and the chromosome.

The most glaring hole in the manuscript right now is the Methods section where I need to describe the bioinformatics methods I've used to analyze the USS motif in the genome. The post-doc responsible for the DNA uptake experiments has already written her part of the Methods, and today we're going to sit down together and write my part, as well as work on polishing the Results (I think this is in OK shape) and the Discussion (needs work!).

Polishing the Sxy manuscript

We're fixing the final details on the Sxy manuscript; I'm hoping to have it submitted in the next 24 hours. Below I'll try to summarize what it says, in less technical terms than the Abstract uses.

We already know that the Sxy protein regulates expression of competence genes; here we're examining how Sxy itself is regulated. Our most powerful tools are regulatory mutations that turn Sxy on when it would otherwise be off. The paper starts by describing new mutations that, taken all together, strongly suggest that expression is controlled by changes in RNA folding. We conclude this because all of the mutations change how RNA can fold, but only one of them changes the Sxy protein sequence (and that in a trivial way).

The paper then presents data showing that the RNA folding changes don't affect how much Sxy RNA is made, but how efficiently the RNA is used to make Sxy protein. Then more data shows that normal cells translate their Sxy RNA more efficiently when they're starved, but the mutant cells translate it efficiently even when they're not starved.

The cell uses the genes/proteins that Sxy regulates to take up DNA, so these results help us understand how being starved causes cells to take up DNA. We have argued that cells take up DNA as food (not to get different versions of their genes as others have assumed), so these results strengthen the evidence that DNA uptake is an adaptive response to starvation.

What's UP, Sxy?

One of the grad students has been doing experiments to clarify how the transcriptional activator protein Sxy turns on genes that have its recognition sequence, the "CRP-S" site. He's given me a draft of a paper he's writing about this work.

Sxy works by interacting with another transcriptional activator protein called CRP. CRP's job is to bind to DNA at CRP sites and, by bending the DNA, help RNA polymerase start making RNA. CRP-S sites are much harder to bend than normal "CRP-N" sites, and we have been thinking that Sxy acts by helping CRP to bind (if it can't bend the DNA it lets go).

But several pieces of his data tell us that binding isn't enough. A protein from E. coli (we work mostly in another bacterium, Haemophilus influenzae) binds CRP-S sites fairly strongly in the test tube, but when it's in H. influenzae cells it can't turn the CRP-S genes on without Sxy's help. And changing the CRP-S sequence so H. influenzae's CRP can bind it without Sxy in the test tube doesn't enable CRP to turn the gene on in cells without Sxy's help.

So the grad student is suggesting that Sxy may also affect how RNA polymerase interacts with the DNA. RNA polymerase binds to DNA more effectively if its 'tail' makes contact with an AT-rich sequence called the "UP" sequence. Many genes have an UP sequence, and at normal CRP-N sites CRP is thought to help RNA polymerase's tail bind to it.

He's found that the genes with CRP-S promoters have what look like three precisely-spaced UP sequences beside their CRP-S sites, and in his draft paper he proposes that Sxy acts partly by helping RNA polymerase make contact with these sites. We could test this by changing these sequences to not fit the UP consensus, and seeing if this makes the promoter unable to be activated by Sxy.

This work doesn't address the question of whether Sxy does help CRP bend the DNA. But we won't be able to address this until we have conditions where purified CRP and Sxy both interact. Right now we have the CRP but not the Sxy - it's a very difficult protein to work with.

Welcome, visitors from the Just Science links

Unlike most of the other science blogs you might check out, this is a real 'research blog'. In it I write about the experiments and analyses we do in my lab, and about what I think they mean. I've been doing this for about 6 months, and I find that writing it helps me think more clearly about our work.

I don't expect casual visitors (or any visitors except those from my lab) to follow the complexities of the science, but it might give them a sense of what more-or-less ordinary* scientists do and how we think.

*No, of course I don't really think I'm any more ordinary than any other scientist (we all think we're special), but I'm a Canadian so I feel obliged to pretend.

DNase I: friend and foe

My posting has been a bit infrequent because I've been focused on getting drafts of our two grant proposals ready for pre-submission critiquing by the volunteer reviewers (blessed be their names and their progeny). But that's done, with a lot of help from a grad student and a postdoc. I've committed myself to the Week of Science Blogging, so I'll be posting at least daily for the next week.

We put a fair bit of effort into crafting a tightly focused "Significance" section to end each proposal (after the detailed descriptions of the experiments we propose). It's important to avoid ending with strings of platitudes, so ours is a bit on the edgy side. On the other hand we can't afford to sound too arrogant (this is a Canadian granting agency).

Some science: We had been hoping to be able to use preparations of cell membranes to study how a radioactively-labeled 'test' DNA fragment interacts with the uptake machinery. But we had overlooked the problem of all the cellular DNA that will be released when we break the cells open to get the membranes.

When the cells are first opened the uptake proteins will bind the abundant cellular DNA, and unless we get rid of the DNA they've bound, they will never bind our test DNA. In principle we could get rid of the cellular DNA by adding a lot of DNase I, an enzyme that breaks down DNA. But then we would need to get rid of ALL of the DNase I , as otherwise it will break down our test DNA!

The postdoc (who discovered the problem) is going to do some preliminary testing to see if this analysis can be salvaged, perhaps by repeatedly washing the membranes to get rid of the DNase I.