Do competent cells stick to an agar layer containing DNA?

Apparently not.

I made plates of agar culture medium topped with a thin layer of agar containing chromosomal DNA (1 µg/ml or 10 µg/ml).  I layered ~ 10^8 cells (log phase or competent) on top of the DNA-agar, let them sit for 10 min, and then washed the surface by flowing buffer over it (2 x 25 ml).  Then I overlaid the agar with 1 ml  of medium ± DNase I.  Finally I washed the agar surface one last time with 25 ml of buffer and plated both this final wash and the top agar itself. 

If the competent cells were able to bind to the DNA I had mixed into the top agar, I expected to see many more competent cells than log-phase cells in the top agar when DNase I wasn't used.  And when DNase I was used, I expected Wash 3 to contain most of the cells that would otherwise be with the top agar.  But neither of these effects were seen.

I don't know if the differences between the log-phase cells and the competent cells (leftmost bars vs all the other bars) are significant.  It could just be because the log cells were my first attempt.

But there are clearly no significant differences caused by treatment with DNase I, nor by use of the two different DNA concentrations.

Controls I failed to do: 

• Top agar with no DNA.  This would control for non-specific sticking.
• Transformation of the cells by the NovR allele in the DNA.  If this gives no transformants, maybe none of the DNA was exposed above the agar surface.  If it gives lots of transformants, maybe the exposed DNA was all released from the agar into the medium.

Other changes I could try:  There's no reason to put the DNA-agar on top of normal sBHI agar, since I'm not going to try to grow the cells.  Instead I can just make thin layers of agarose with DNA, varying both the agarose concentration and the DNA concentration.  I could try higher DNA concentrations, up to 100 µg/ml.  For ease of washing I could make these on glass slides rather than in Petri dishes.  (Nope - just tried that and the agar just slid off the slide when I washed water over it.)

Tangled up in goo?

I just reread an opinion piece from Richard Moxon's group, considering whether H. influenzae produces anything that should really be called a biofilm.  They agree that H. influenzae cells will grow on surfaces, but they don't think there's any real evidence that these films are the result of a developmental program that evolved because of the benefits of biofilm formation.

Although 'biofilm' can simply mean a layer of bacterial cells and biological macromolecules, associated with a solid surface or an interface, most microbiologists assume that  biofilms arise by regulated developmental programs.  Said another way, most microbiologists think that bacteria grow in biofilms because natural selection has favoured genes that cause them to do so.  They think that bacteria respond to certain stresses that arise in biofilms, or simply to the presence of a surface, by turning on sets of genes that optimize their ability to initiate a biofilm and the physical properties of the biofilm that forms.

It's not that researchers have rigorously evaluated and discarded the alternative explanation - that biofilms form just because bacteria have adhesive organelles and macromolecules are often sticky and fibrous.  But they, and grant agencies and journal editors, find the adaptive perspective more interesting, so that's what gets done and published.  The Moxon paper is complaining that the term biofilm, with its aura of scientific coolness, is being applied to H. influenzae structures that fit only my simple non-adaptive definition.

Thus at least part of the reason why my simple DNA-on-glass-tubes experiment didn't work is probably that the H. influenzae cells were never going to form a biofilm at all.

After we discussed this at lab meeting, I'm going to try something different.  I'll make a fake biofilm by putting a layer of top agar that contains DNA on top of normal agar plates, and then testing whether cells stick to the surface.  I'll also return to an experiment I did by accident when I was a post-doc, which showed that multiple cells can bind to single DNA fragments in solution.

Nothing to see here folks

My repeat biofilm experiment was seriously compromised by a tube-label failure - the identifying numbers I'd written on the culture tubes mostly washed off when I washed the crystal violet stain from the tubes. (Somehow my new red sharpie's writing didn't stick to the surface of the new glass tubes, but instead flaked off like a whiteboard marker.) 

Luckily I'd also labeled each tube to indicate the treatment, and these marks (other sharpies) didn't wash off.  The blue bars are the amount of crystal violet that stained tubes that had been pretreated with DNA.  The green bars are for tubes pretreated with just the buffer, and the pink bars are for tubes pretreated with buffer and then given 1 µg DNA/ml in solution in the cell culture.

If we set aside the murR749 hypercompetent mutant, I can make a few (weak) generalizations.  First, my DNA pretreatment didn't increase the amount of staining, and thus didn't increase the numbers of cells adhering to the sides of the tubes.  Second, adding dissolved DNA to the cultures may have slightly increased the amount of staining.  Third, the use of the sxy hypercompetent mutant or either non-competent mutant (sxy- or pilA-) didn't affect the amount of staining.

The murE749 mutant had higher staining than the other strains, especially if I ruthlessly assign the high 'unknown' to it.  This strain's staining was also higher in the first version of this experiment.

Is any of this worth following up on?  I don't think I'll bother to repeat this experiment using a more reliable Sharpie.  Should I put more effort into getting DNA attached to a surface?  I think first I should finish my post about what biofilms are...

No DNA-enhanced biofilms

Yesterday I tested whether pre-coating glass tubes with DNA helps competent cells form biofilms.  The answer is clearly No, but the results are nevertheless interesting, in a "That's peculiar!" way.

I did pretty much what I had planned (see previous post):  I added 2 ml of a DNA solution to new glass culture tubes (in high-salt, low-pH), left it for an hour at room temperature, removed the solution and let the tubes dry for several hours at 37 °C.  I then rinsed these tubes and untreated tubes with the high-salt low-pH buffer.  I then added 3 ml of either high-density (~2x10^8 cfu/ml) or low density (?~10^7 cfu/ml?) cultures of the five strains I wanted to test, and left them for either 4 hr (high density cultures) or 18 hr (low density cultures) at 37 °C, just sitting in a rack.  When the time was up I dumped out the cells, rinsed the tubes once with phosphate-buffered saline (no vortexing), and added 2 ml of a 0.1% solution of crystal violet in water.  (I tried putting the crystal violet in my high-salt low-pH buffer but it wouldn't dissolve).  I let the tubes sit for 10 min, dumped out the crystal violet, rinsed them with water, and let them dry.  Then I resuspended the dye in 1 ml of 95% ethanol and measured its absorbance.  The amount of dye that stuck to the tubes should measure the amount of cells that were stuck to the tubes.

I was hoping that the DNA treatemnt would increase the number of cells sticking to the sides of the tubes, especially for the two hypercompetent mutants (sxy-hyp and murE-hyp), but not for the non-competent mutants (sxy- and pilA-).

The graph shows that the DNA treated tubes (light blue and light green) had LESS, not more, stain.

This could have just meant than my pre-treatment didn't work - that the DNA had washed off the tubes before I added the cells.  But that wouldn't explain why the untreated tubes had more cells that didn't wash off, especially after the long incubation.  Instead, the DNA treatment appears to have done something that actively decreased cell adhesion.

My first look at the 18 hr cultures had me expecting a very different result.  In all of the cultures in the no-DNA tubes, most of the cells had sunk to the bottom of the tube, but most of the +DNA tubes were cloudy all the way up.  This led me to think that they had thick biofilms, but now I think that the cells were just more evenly suspended in the medium.  Unfortunately I didn't measure the cell densities or anything else, or do replicates, or even carefully check which of the +DNA tubes looked less cloudy than the others (I think it was the pilA mutant).

So I guess I should repeat the whole thing.  But first I should put some thought into possible explanations...  ...OK, I have no idea.  There shouldn't have been much DNA in solution in the +DNA tubes, given that I did rinse them before adding the cells.  But I could test this by having tubes where I just added 1 µg of DNA in solution to the culture.  Might the soaking in DNA have modified the physical properties of the glass surface in some way?  This time I should treat the -DNA control tubes exactly the same way I treat the +DNA tubes.  I guess I also need to do replicates this time.  I won't bother with the high-density 4-hr incubations, I'll just do overnight ones that start at low cell density.

I'll also count each culture's cfu/ml at the start of the incubations, and check the ODs at the end (with and without mixing up any settled cells.  And I'll be more meticulous in my washing steps.

Better go pour a few plates and start preparing the tubes...

DNA and biofilm formation, planning

I want to test whether their ability to bind DNA helps competent cells form or join biofilms. 

The standard way to test for biofilm formation is to incubate cells in a polystyrene microtiter-well plate, remove the unbound cells, and stain the remaining bacteria with crystal violet.  But I don't know anything about whether DNA sticks to polystyrene (I can only find bapers about chemical methods of attachment, so I don't think it spontaneously sticks), and I think it's better to do the test under conditions where biofilms don't form easily.

As a preliminary test, I first want to coat the inside of glass culture tubes with a film of DNA.  I'll then add dilute H. influenzae cells, either wildtype or constitutively competent, and incubate overnight (with or without constant mixing? probably I'll try both).  The on-line sources say that DNA binds 'avidly' to glass under conditions of high salt and low pH (below 7.5).

So I'll prepare a high-salt low-pH stock of H. influenzae chromosomal DNA (100 µg/ml DNA, 1 M NaCl, 50 mM Tris pH 7.0).  I have an old high-concentration DNA stock somewhere.  I'll add 2 ml of this to new (sterile)glass culture tubes and let them sit for 1 hr at 37 °C.  Then I'll pour out the DNA (saving it for next time), and leave the tubes to dry  at 37 °C overnight.  In the morning I'll fill the tubes with 1M NaCl 50 mM Tris pH 7.0, leave them for 30 min, vortex them and discard the wash solution.  Then I'll add 5 ml of cell culture in sBHI, at a density of ~ 10^7 cfu/ml, and incubate the tubes at 37 °C for 5 hr or overnight.  Then I'll remove the cells, and add crystal violet (0.05%) to stain the biofilm.  After 10 min I'll wash the tubes with water twice, and then dissolve the remaining crystal violet in 95% ethanol and measure its absorbance at 570 nm.

What cells will I use?  Wildtype cells, hypercompetent mutants (sxy-1, murE749), noncompetent mutants (sxy-, cya-, pilA-).  I'll have control tubes with no DNA, and tubes with added DNase I.

Submitted!

Back to the bench...

Well, not quite yet, but very soon.  The CIHR proposal is in excellent shape - we're doing a final read through and then final revisions (just polishing) on Monday, and then I'll click 'Submit' two days ahead of the deadline (a new record).

It's in such good shape that I'm already preparing to get back in the lab.  The first step is updating the Table of Contents of my lab notebook.  This is just a Word file that lists each experiment's number and date with a few words or sentences describing it.  Having it is very useful as I can search for any word to find the relevant experiments (which I might otherwise have forgotten about).  The process of updating it is also very useful, as I have to read through my notebook and summarize each experiment.  That's what I've been doing, for the work I've done since January.

This readthrough showed that I have two projects ongoing.  One is the optical tweezers work.  Now that the system is working reasonably well, it's time to make some well-characterized DNA-coated beads (measuring the amount of DNA on each bead) and start trying to get cells to grab onto the DNA.  It would be good to demonstrate this at the bench as well as trying to see it with the tweezers apparatus (abbreviated 'OT' by its creator).

The other project is the purine work on the regulation of competence.  We now have the mutants we need, properly verified by PCR, so it's time to do some more rigorous phenotyping.

Bead and cells

A figure for our CIHR grant proposal.

Phenotypes of retraction and anti-backsliding mutants

One component of the CIHR proposal we're working on is identifying mutants that are unable to retract their pseudopili ('retraction mutants') or to preventing DNA from sliding back out after it has been partly pulled in ('anti-backsliding mutants').  These mutants are expected to have similar but not identical phenotypes.  The retraction mutant should be able to bind DNA at the cell surface but not take up any of it into the periplasm, so when cells are incubated with 32P-labeled DNA, all of the 'cell-associated' DNA should be removed by DNase I.  The anti-backsliding mutant should also bind DNA, but it would be able to bring some DNA into the periplasm.  This uptake should be inefficient, especially if the next step (translocation into the cytoplasm) is blocked by another mutation or by using circular DNA.  So uptake should be reduced but probably not eliminated.

But the screens for these mutants aren't very good, so I need to think more about this.

Consider the retraction mutants first:  These mutants should be able to bind DNA, but that doesn't mean they will have the same level of cell-associated radioactivity as wildtype cells.  That depends on whether the uptake machinery is reused to take up more than one DNA fragment.  To explain this better, here's an extreme example:  Imagine that wildtype cells have only one DNA-binding structure on their surface.  After DNA binds to this structure, the DNA is passed to an uptake machine (again only one per cell) which pulls it across the outer membrane into the periplasm.  In species A, the structure is now free to bind another DNA fragment and pass it to the uptake machinery, and then another, and then another.  In species B, each structure (or each uptake machine) can be used only once.  When we compare the amounts of 32P-DNA associated with cells of species A and species B, species A will have four times as much.  Now consider a mutant of each species whose uptake machine is unable to pull DNA in at all.  These cells can still bind DNA but they can't take it up.  The species A mutant will only have 25% as much cell-associated DNA of wildtype species A cells, but the species B mutant will have 100%.

Is H. influenzae's DNA uptake like species A or species B?  We have suspected it's more like species B (i.e. uptake machinery isn't reusable), but the evidence for this is mainly just that the number of transformants doesn't keep increasing with increasing time or increasing DNA concentration, as if the uptake machinery had been used up.  But the same effect may not be seen with DNA uptake experiments (I have one example where it isn't), which would mean that the bottleneck is at a later step (DNA translocation or recombination).  I think we need to carefully investigate this first, so we'll know what phenotype to expect of our postulated retraction mutants.  The post-doc has some newer uptake data that may address this.

The other issue is how we measure cell-associated DNA.  The standard procedure is to incubate competent cells with a 'saturating' amount of DNA labeled with 32P (or 33P) and wash the cells by centrifuging them, resuspending them in fresh medium (with vigorous vortexing), centrifuging them again, resuspending them again, and probably centrifuging and resuspending them one more time.  The goal is to remove all the DNA that's in the medium but leave all the DNA that's stuck to or inside the cells.  This procedure should be fine for DNA that's inside the cells, but we don't really know how well this works for DNA that's just bound to the outside of the cells.  Does the vigorous vortexing pull loosely bound DNA off the cells?  The answer probably has something to do with Reynolds numbers, but that's beyond my expertise.  In Neisseria, does it break off the pili that the DNA may be bound to?  The RA probably can answer this, not because she knows about Reynolds numbers but because she's worked with Neisseria.

So I'm considering a different way of washing the cells.  We routinely use filtration to collect and wash cells when we're transferring them to competence medium, so why not also use it to collect and wash cells in DNA-binding assays?  We could first dilute the cells+DNA into a large volume of medium (100-1000-fold dilution, and then collect the cells by filtration (perhaps using only gentle suction to minimize shearing forces at the filter).  We can then easily wash the filter with lots more medium to make sure all the unbound DNA is removed.  Then we can just pop the filter into a scintillation vial for counting.  To detect only DNA that has been taken up, we can add DNase I to the cells, dilution mix or wash.   The dilution and washing medium should be cold or at room temperature to stop uptake of DNA that's already bound.  Clogging of the filter isn't a problem because we'd be using only ~1 ml of cells, and the medium is cheap so the dilution and washing volumes are limited only by the capacity of the collection flask under the filter.  The filters cost a couple of dollars each, but I think this would be well worth the cost.

We also can't be sure that the mutant's defect will be due to defective retraction of the postulated pseudopilus, but that should be a separate post.

What's known about T4P and DNA uptake in Neisseria

I've been rereading an important 2002 paper by Finn Erik Aas et al. (Mike Toomey's group; Molecular Microbiology 46:749-760).  They dissect the roles of the Neisseria type IV pilus components PilE (the major pilin), ComP (a minor pilin important for DNA uptake), and PilT (the retraction ATPase).  The results are important and need to be considered in the CIHR grant proposal we're rewriting. 

I was about to try to summarize them here, when I wondered whether I might have already posted anything about this paper.  A quick search for 'Aas' found a whole post from last summer, which I had completely forgotten about!  So here I'm instead going to edit/annotate/rewrite that post.

An important difference between this paper and previous papers is their careful attempts to separate of DNA binding from DNA uptake.  DNA binding is usually measured indirectly  by giving cells radioactively labeled DNA and comparing cell-associated cpm with and without pretreatment with DNase I, which removes DNA that has bound to cells but not been taken inside them.  And that's how it's done in this paper - the 'binding' values are how much of the total cpm is removed by DNase I treatment (i.e. they're determined by subtraction).

The Aas et al. paper  showed that, when 5x10^8 wildtype Neisseria cells are incubated for 30 min with 500 ng of 32P-labeled plasmid DNA containing the Neisseria uptake sequence (the DUS), only about 1-2% of the DNA sticks to the cells (i.e. is still there after 10 min on ice and three washes with cold medium).  They found that more than half of this DNA is taken up during the 30 min incubation, because it wasn't removed when the cells are incubated with DNase for 10 min at room temperature before being washed instead of being stored on ice.  If the DNA doesn't contain a DUS the cells still bind a lot of it (25-100% of the +DUS binding amount), but almost none of the bound DNA is taken up (> 0.01% of the 500 ng).  This says that cells will bind any DNA but can only take it up if it has a DUS.

The minor pilin-like protein encoded by comP is normally expressed at a very low level (so low that it's barely detectable by protein assays), but the level is high enough to show that the ComP protein is incorporated into pili along with the major pilus protein PilE.  Cells lacking PilE don't have visible pili and they didn't bind DNA or take up DNA, but cells lacking ComP have normal-looking visible pili and they bound just about as much DNA as wildtype cells.  But they hardly took up any of this DNA even if it had a DUS (they interacted with DUS+ DNA the way wildtype cells interact with DUS- DNA).

This strongly suggests that the ComP protein recognizes the DUS at the cell surface, but that ComP is not involved in the non-specific DNA binding step.   Consistent with this, overexpressing the normally-scarce ComP protein increased by 20-fold the amount of DNA bound and taken up, and proportionately increased the transformation frequency. This increased uptake was specific for DNA containing a DUS, although a modest increase was also seen for DNA that lacked DUS.  Overexpression also greatly increased the amount of ComP protein in the pili.  I think this result says that ComP can cause DUS-specific DNA binding, on top of the normal non-specific binding.  ComP, when assembled into pili, would be able to bind DUS-containing DNA but not other DNA.

However, the authors found that, when they purified ComP, it did not bind specifically to any DNA.  They tested both 'recombinant' ComP (+ His-tagged) and 'overexpressed' ComP, but they they don't show this data and they don't say how the binding assays were done.  Does this result mean that ComP really doesn't interact with the DUS?  Maybe.  The authors suggest that it acts by modifying some other protein so that it binds the DUS, but another explanation might be that ComP loses its DNA-recognizing function when it's purified away from the T4P complex.

My tentative model:  ComP is processed by the  prepilin peptidase as is the main pilin (PilE), and is assembled along with PilE into the pilus filament, where it forms a very minor component.  Once DNA binds non-specifically to the PilE part of the pilus, it can interact with ComP and, if the DNA has a DUS, be taken up when the pilus is retracted.  If the amount of ComP in the pilus exceeds the non-specific binding capacity of the PilE, DNA will bind directly to ComP (even if it can't be taken up, see below).

What about PilT?  In most bacteria with T4P, PilT provides the power to retract type 4 pili filaments, by disassembling the subunits from the pilus base.  (The exceptions are H. influenzae and its relatives in the family Pasteurellaceae, who don't have a pilT homolog.)  As pili are thought to bind DNA at the cell surface, PilT would then be responsible for pulling the pilus and its DNA into the periplasm.  PilT mutants had already been shown to have the expected phenotype: abnormally large amounts of pili, and unable to take up DNA or be transformed.  We might then expect that having more pili would make pilT mutants bind more DNA than wildtype cells, but Aas et al. found that they bound <1% of the wildtype amount.  They took up about 20% of what they bound, but I think this may be at the detection threshold. 

Overexpressing ComP in a pilT knockout increased the DNA binding (but not uptake) by about 5-fold.  This added binding was DUS-dependent.  I could add this to my tentative model:  The nonspecific binding by PilE requires PilT, but we have no idea why (I'll call this the magic PilT effect).  Specific binding by ComP does not.

They then made transcriptional fusions of pilE and pilT to the E. coli lac promoter (in different strains) so they could keep the genes turned off or turn them on by adding IPTG, as desired, and measured transformation.  When each gene was on throughout the transformation experiment, transformation was close to normal, as expected.  When it was off, there was no transformation, again as expected.  They then tried turning the genes on halfway through the transformation experiment, to find out whether PilT was only needed to pull the pili in

When pilE was off during the DNA incubation step and then turned on after the cells had been washed free of unbound DNA, transformation was down >500-fold.  This is consistent with both nonspecific and specific binding happening only in assembled pili - it's no use making the pili after the DNA has all been washed away.  When pilT was off during the DNA incubation step and then turned on after the cells had been washed free of unbound DNA, transformation was down  >32-fold; this says that some DNA can bind to cells in the absence of PilT, and then be later taken up once PilT becomes available.

Does this mean that the binding that happens before turn-on of PilT is specific for the DUS?

The authors then did the same experiment in cells overexpressing ComP.  When there are no pili (pilE off) my model predicts that no DNA will bind even though there's tons of ComP, so turning pilE back on after the cells are washed shouldn't give any transformants.  In fact it gives some, but not a lot.  When there are pili but no PilT, my model predicts that having tons of ComP will increase DNA binding, and that this will increase transformation frequencies once PilT becomes available.  My model predicts that this binding will be DUS-dependent, but that isn't shown.

Summary of the PilT bit:  Having pilT off while DNA is binding to the pili reduces transformation frequencies 32-fold when most binding is initially nonspecific, because most of this binding is caused by the magic PilT effect, but only 7-fold when more of the binding is by ComP to the DUS, because the magic PilT effect doesn't act on ComP binding to the DUS. 

But uptake requires pilus retraction.  So maybe I should conclude that, in wildtype cells, most DNA initially binds non-specifically to the many pili that have no ComP but have been magicked by PilT.  DNA bound to these pili is not released to the solution, and these pili are not retracted.  But if a DNA fragment has a DUS, the DUS can bind to one of the fewer pili (or pseudopili?) that have ComP (binding to directly to ComP or to another protein).  Once a ComP-containing pilus has bound DNA it can be retracted, pulling the DNA in and initiating uptake.  This pilus might then elongate and bind another DUS, or other ComP-containing pili might be continually assembled and accept DNAs from the PilE pili. 

I'm going to stop here, because the more I read the confuseder I get.  I'll talk this all over with the RA tomorrow morning and see if she can sort me out.

Role of competence in biofilm attachment (part 2)

Yesterday I talked to my colleague down the hall about a possible joint project.  She reminded me that one of her grad students has done quite a bit of work on DNA in Campylobacter jejuni biofilms; the student has a mutant that makes faster/thicker biofilms, and whose biofilms have much more DNA than is usual.  The biofilms also fall apart in the presence of DNase I.  She gave me some papers to read, one by the student and two others about DNA in Pseudomonas biofilms.

Despite quite a few publications about DNA in biofilms, none consider the role of competence.  So I still like the idea of testing whether biofilm formation by C. jejuni and by H. influenzae is enhanced by the ability to bind (and take up?) DNA.

I also did some reading about Campylobacter competence.  There aren't many papers, but there's some nice work characterizing genes needed for DNA uptake, and characterizing how competence varies under different culture conditions.  We'd need to send for a suitable mutant, one that was unable to take up DNA but had no growth defect.  Ideally we'd want to know that the cells couldn't bind DNA, or couldn't turn on the competence genes, but I'll have to do more reading to see if such a mutant is known.

All the H. influenzae biofilm work appears to have been done with 'nontypable' strains (clinical strains lacking a capsule, which are important causes of ear infections), rather than with the Rd strain we normally use.  I was hoping that these experiments might have used Rd as a control, but apparently not.  I'd much prefer to do these experiments with Rd, but we'll have to check whether it readily forms biofilms in culture.  If not, we might be able to use one of the nontypable strains that we've found to be readily transformable, but we'd need to introduce a hypercompetence mutation into it.

The minimal experiment would use glass culture tubes, some of which had been precoated with DNA.  Some DNA sticks to plain glass, although I can't find out how much.  A lot more sticks in the presence of saturating sodium iodide but that's not an option for living cells.  So we could try just filling tubes with moderately concentrated solutions of H. influenzae or C. jejuni DNA, maybe 10 µg/ml, leaving them for a few hours, and then letting the DNA dry on overnight.  (There's evidence that C. jejuni, like H. influenzae, prefers to take up its own DNA, although there's no obvious uptake sequence in its genome.)

We'd probably be wise to rinse the tubes with culture medium in the morning, to remove DNA that wasn't stuck to the glass surface.  Then we'd add dilute cultures of hypercompetent (sxy*) or sxy- H. influenzae, or wildtype or noncompetent C. jejuni, and let the cultures incubate overnight at 37°C.  We would also include cultures with added DNase I as another negative control.  After overnight culture (or shorter), we'd wash out the non-attached cells, stain the biofilm with crystal violet, and measure the staining.

Another experiment would measure whether being competent helps cells attach to an existing biofilm, presumably by attaching to the DNA.  For this experiment we'd grow biofilms in the ordinary way, using whatever strain makes good biofilms.  Then we'd add antibiotic resistant H. influenzae or C. jejuni cells (competent and non-competent), and give them time to attach.  Then we'd wash away all the unattached cells, break up the biofilm by adding DNase (or ???) and plate on antibiotic plates to count the newly-attached cells.  To distinguish binding to DNA from binding to other biofilm components, we could swamp the culture with free DNA - that should abolish DNA-specific attachment.  (This control could also be done for the minimal experiment.)

DNA mediated attachment in biofilms

One questioner after my CIfAR talk asked whether competence genes might have some other function for the cell.  When I raised this at lab meeting yesterday, the post-doc pointed out that competent cells are able to attach to and pull on the DNA that's a major component of biofilms, and we considered whether this might help cells to establish biofilms, to move within biofilms, or to avoid being displaced from biofilms.

We then started considering experiments we might do.  My idea was to precoat glass surfaces with DNA, incubate them with cultures of cells that can form biofilms, and then measure whether the DNA increased the initial steps of biofilm formation.  The lab down the hall studies, among other things, biofilm formation by Campylobacter, which is naturally competent, so we might work with them to test this in both H. influenzae and Campylobacter. 

What controls would we want?  We'd certainly need to test cells that can and cannot express their competence genes.  Competent cells express type IV pili, and these might allow attachment and pulling. independent of their involvement in DNA uptake, but I think doing the experiment with and without DNaseI would control for DNA-independent effects of pili.

The basic assay for biofilm formation is very simple, and the Research Associate's extensive experience turns out to include doing lots of these.  I think it's time to talk to my colleague down the hall, to see if she's interested.

Variation in core genes

I ended my talk at the CIfAR Integrated Microbial Diversity meeting by asking how much variation we should expect in 'core' bacterial genes within a single species, and I considered this more at lab meeting yesterday.  We see a lot of variation in competence phenotypes, due, we assume, to variation in the corresponding genes.  But we don't know if this variation is telling us something particular about how selection acts on DNA uptake and transformation, or is just 'normal'.

The variation could accumulate because loss of competence is sometimes a good thing, at least in the short term, but it could also be accumulate just because mildly deleterious changes are only slowly eliminated by selection.  This is especially likely to be true for genes that are only expressed occasionally.

So the variation in competence might mean that cells only need DNA uptake occasionally.  It would really help if we know more about when cells develop competence int heir natural environment...

Planning my CIfAR talk

OK, the CIHR proposal is presentable and has been sent to the internal reviewers, so now it's time to start thinking seriously about the talk I'll give at the CIfAR Integrated Microbial Diversity meeting on Sunday morning.  The talk's title is "Sporadic Sex? Scary Food?  The biodiversity of bacterial competence", but I'm going to also talk more generally about the genetic diversity of bacteria, especially the diversity within a single species.

I've started making an outline:

1. Genetic diversity in eukaryote species:  Not a lot.  The members of a species typically have very similar chromosomes, with the same genes arranged in the same order.  Different versions of the genes differ by no more than a few %, and there are very few differences in gene content or arrangement.
2. We take this for granted, but it's probably one of the many consequences of sexual reproduction.  Individuals with chromosomes whose genes have been rearranged (by, for example, an inversion) are at a big disadvantage in sexually reproducing species, because such chromosomes don't pair properly with normal chromosomes at meiosis and the resulting gametes often have two versions of some genes and none of others.
3. What about in bacterial species?  I should probably start by considering what we want 'species' to mean in Bacteria.
4. For most eukaryotes, the 'biological species concept' is the most useful way to group individuals into species.  In its simplest form, it says that all individuals that can reproduce sexually with each other should be considered members of the same species.
5. But bacteria don't have eukaryote-type (meiotic) sexual reproduction.  They don't have any process that regularly combines the sets of genes from two individuals and then shuffles them into two 'recombined' sets.  Maybe I'll talk here about what bacteria do have that puts genes into new combinations, or maybe I'll leave this till later.
6. But we can define bacterial species as groups of individuals for which at least 50% (75%?  90%?) of their genes are recently descended from a common ancestor (recent enough that sequence divergence is no more than a few %), and for which most of these genes are syntenic (arranged in the same order).  This definition works for most of the groupings that microbiologists consider for other reasons to deserve being called species.
7. For this 50% or 75% or 90% of the genome, the members of a bacterial species are a lot like a eukaryotic species.
8. But these species are far more genetically diverse than are eukaryote species, because of the other 10% or 25% or 50%.  Many of these sequences are stupendously diverse...

That's as far as I've gotten.  I mustn't forget that I only have 25 minutes to talk, plus 5 for questions...

Improving the CIHR proposal's focus

I've been thinking more about one reviewer's concern that our CIHR proposal was 'unfocused'.  Although part of the problem was that we were proposing to do too much, I think the underlying problem was indeed a genuine lack of focus.

We identified three major gaps in the understanding of DNA uptake by gram negative bacteria, and proposed a series of experiments that would help fill in these gaps.  BUT, we didn't tie these experiments together very well, and we weren't proposing to completely fill any of these gaps.  They were all parts of a big problem, but they weren't the whole solution (that will likely take many years of work by more than one lab).

Ideally, our proposal should identify one well defined problem that our set of experiments will address, and then we should show how all of our experiments will come together to solve it.  I think the way to do this is to focus on first determining whether uptake consists of distinct steps, and then on characterizing the steps as much as possible.  So we'll propose a distinction between the initiation of DNA uptake, an event that happens at one place on the DNA fragment (we think at the USS), and the subsequent uptake, an ongoing process that gradually pulls the entire fragment into the periplasm.

We can further ask whether there is an initial binding step that's distinguishable from the initiation of active uptake.  That is, does the uptake machinery at the cell surface first bind to DNA, either at the USS or first elsewhere and then at the USS, and then kink the DNA and pull the initiating loop across the outer membrane?  The alternative is that binding is not an independent step, but that DNA is kinked and pulled immediately on contacting the uptake machinery.

All of our experiments contribute to clarifying these distinctions, in one way or another.  So we just need to situate them in this context, and emphasize how they reinforce each other.  The second goal, of characterizing the steps, is open-ended, but I don't think this creates a 'focus' problem.

The distinction between binding and initiation has implications for the evolution of uptake specificity too.  We've previously considered two alternative explanations for uptake specificity.  Either the protein responsible for uptake specificity is an add-on, evolved to screen DNA for relatedness before allowing it to be taken up, or the uptake specificity is intrinsic to the initiation process, occurring because specific sequences are easier to process.  Identifying a separate binding step would favour the former.

Identifying the retraction protein

I've added a new (but obvious in retrospect) question to our CIHR proposal:  Which protein creates the retraction force that pulls the DNA across the outer membrane?  

The PilT protein that does this in other bacteria, but H. influenzae and the other Pasteurellaceae have no pilT gene.  We expect that the same retraction force is responsible for both initiation and continuation of uptake, and we can't begin to investigate its function until we know its identity (duh!).  We have a full set of nonpolar knockout mutants, covering all the genes that are implicated in competence by either direct mutations or by their membership in the CRP-S regulon, so we will screen these for the expected phenotype: no transformation, no DNase I-resistant DNA uptake, but binding of DNA to competence-induced cells (DNase I-sensitive radioactivity bound to cells.  

Testing for transformation and for DNA uptake are routine in our lab, but binding will be a bit trickier because we don't know what to expect.  Tests for DNA binding are hard to interpret in normal competent cells, because DNA is taken up soon after being bound.  So 'binding' is measured as the relative difference in cell-associated radioactivity ± treatment with DNase I ((noDNaseCPM - DNaseCPM)/noDNaseCPM).  This isn't very sensitive, because noDNaseCPM isn't much larger than DNaseCPM.  We don't know how much DNA should bind to the surface of cells that can't pull it in, nor how strong this binding should  be.  But DNaseCPM should be negligible (background in these experiments is usually <1% of DNaseCPM), so even if binding is only 10% of uptake  it should be readily detected.

Should this binding be USS-specific?  I think yes, but we won't throw out any uptake-minus mutants that bind DNA non-specifically. 

This screen will be the first set of experiments in the proposal.  The second set of experiments will be the DNA-protein crosslinking, and we will test any candidate retraction proteins for crosslinking to DNA, and test the corresponding knockout mutants for effects on the binding of other proteins.  A genuine retraction knockout should increase the crosslinking of the binding/initiation proteins that act upstream of  (before) it.

What if this screen for retraction mutants doesn't find anything?   We won't conclude that no retraction protein exists, but rather that there is no real binding step distinct from the initial initiation that pulls the first DNA into the periplasm.

Quick, change focus!

(I wrote this two days ago, but forgot to click 'Publish post'.)

The optical tweezers apparatus is now working well, so I'm off across town tomorrow to give it a try, with the expert help of my biophysicist collaborator.  But can I remember what I had learned so far, and what the status of my DNA-on-beads preps is?  No.  So I'm glad I wrote some blog posts and kept a tolerably good notebook.

I can attach biotinylated DNA to streptavidin-coated polystyrene beads.  The problem that the DNA caused the beads to clump together was resolved by agitating the beads better while they were incubating with the DNA (by putting the tubes on the roller in an orientation where they would be turned end-over-end).  However this resulted in beads that were difficult to pellet for washing, a problem that hasn't been solved but can be circumented by washing the beads by filtration and then concentrating them with a disposable protein-concentrator (Amicon).

• I need to find the bead-DNA preps I made and take them home with me tonight.

I can get cells to adhere nicely to coverslips if I rub the coverslips with ~10 µl of poly-L-lysine solution until dry.  It's easy to then assemble the treated coverslips into 'chambers' for the tweezers work.  I mark an 'X' near the center of the coverslip (on the coated side) with a fine-point sharpie; this makes it easy to be sure I'm focusing in the right place with the tweezers apparatus.  To prepare chambers with cells in them, I add the cell suspension to the chamber, incubate it coverslip-side down for 1-2 minutes, and then rinse thoroughly with 5-10 volumes of competence medium (rapping several times to dislodge weakly bound cells) before sealing with candlewax.  The attached cells are still alive and able to grow and divide if given culture medium.

• I will prepare some more coverslips and chambers before I go home tonight, and take them with me.

I have frozen competent cells (H. influenzae and B. subtilis) in the -80 °C freezer at the biophysics lab.

• But I suspect there may be contamination in the fridge stock of competence medium I use to wash the chambers and to wash and resuspend the cells in after thawing (to remove the glycerol antifreeze), so I should take some fresh stock with me and leave it in the -20 °C freezer rather than in the fridge.

Tomorrow the first objectives will be to make sure I can trap cells with the laser, and I can see the cells on the coverslip.  If that works, we can try bringing a bead to a cell and see if it attaches!

Tweezers progress (I've advanced to a new problem!)

The optical tweezers apparatus is finally functioning, so I spent yesterday trying it out.  I can't yet do what I want to do, but enough of the steps are working that I've been able to get to a new problem.

The various optical components have been adjusted, so now I could (sometimes) trap beads at the laser focus.  This is quite fussy; most of the time the beads are pulled into the focus point (the trap) and then spit right out again.  My colleague thinks this is probably because of a bit of astigmatism in the (cheap) laser, and because she's now using an oil-immersion lens rather than a water immersion lens.  I don't understand why the oil immersion lens would be worse, as the oil is specially designed to have the same refractive index as the glass on either side of it (the objective lens and the cover slip) so oil causes less refraction than water.  But anyway, the trap works best when it is within 20 microns of the coverslip/oil/lens.  Unfortunately it's tricky to get this distance because the micrometer used to move the chamber forwards and back has quite a bit of wobble ('hysteresis').

I could also focus on the cells I had attached to the coverslip, and the images looked much clearer than they had before the optics were adjusted.  Again though, the wobble in the micrometer made this somewhat imprecise.  And once I'd trapped a bead I could use the micrometer to move the chamber away from the objective, thus bringing the trap position to the surface of the coverslip where the cells were. 

At this point, if the beads had DNA on them, the cells might attach to the DNA.  But the beads I had brought were too small.  I had made them in a bit of a rush the night before, and although I was quite sure I'd taken the beads from the little bottle labeled "2.1 micron" I must have somehow used the one labeled "1.2 micron".  So I was using 2 micron beads without DNA, taken from an old tube that was in the drawer of the tweezers lab.

But this is where the new problem became evident.  When I brought the beads to the coverslip where the cells were, the beads quickly became stuck to the coverslip.  This wasn't exactly a surprise.  I knew this was likely to be a problem, and had been trying out ways to prevent it a couple of months ago.  But I hadn't found any that worked.  So I need to go back and try some more, both treating the coverslip after the cells have attached, and pre-treating the beads after the DNA has attached.

I may also be able to solve this problem by not petting the beads touch the cover slip in the first place.  In principle I should be able to bring the bead to a position that's still a few microns above the coverslip.  Depending on how far the DNA extends from the bead surface, the cell will be able to contact the DNA but the bead will not contact the coverslip.  The problem here is the poor control of the chamber position (the wobbly micrometer).  My colleague has what appears to be a better micrometer (bought on eBay!), but she's not sure it is compatible with the present setup.

I'm going to make one more try at this before submitting the CIHR grant (due in a month).  This time I'll take more time to carefully prepare my 2.1 micron beads with attached DNA.  The problem of beads sticking to the sides of the tube when being washed hasn't been solved but it can be minimized.  I'll add biotin to the first wash to block the streptavidin, and finally resuspend them in TE with added BSA to reduce nonspecific binding. 

A plan to identify competence proteins that interact with DNA

We're revising the first Specific Aim of our CIHR proposal because one of the reviewers (correctly) thought that it was too unfocused.

Originally we had:

Aim I. The functions of DNA uptake proteins

Q. 1. Which genes are needed for DNA uptake?

Q. 2. Which proteins contact DNA during uptake?

Under Q. 1 we proposed to create nonpolar knockout mutants of every competence gene (all 25 CRP-S genes and 2 other genes implicated in DNA uptake), and characterize their phenotypes.  Under Q. 2 we proposed to systematically test whether they bound DNA, using His-tagging and crosslinking.

The RA has almost completed making the knockouts.  Those of them with cotranscribed downstream genes (i.e. likely to have polar effects) are not nonpolar yet, because the selection for excision of the selective cassette turns out not to work in our lab strain (it's the standard lab strain Rd).  But we expect she'll be able to manually screen for excision (or we can hire an undergrad to do it).  We expect that this will be completed before the proposed start date of the grant. We now describe this in the Preliminary Results section of the proposal, and we've eliminated what was Q. 1.

We now also have a prioritized list of candidate DNA-contacting proteins, based on several attributes.  One is the presence of protein-export signals and other clues in the sequence annotation.  Another is what is known about the functions of homologs.  Another is what is known from the available mutants; mutants are available for only some proteins, and for some of these the mutant is expected to be polar on another gene that could account for its phenotype.

Aim I. Identifying DNA uptake proteins

Q. 1. Which proteins contact DNA during uptake?

Rationale:  Our in vivo crosslinking strategy can succeed where previous in vitro approaches have failed.  We will begin these studies with the two top candidates on our list, secretin and pilin.  In parallel we will use two simple steps to identify additional strong candidate proteins.

Methods:  1. Basic crosslinking assay:

• First put a His-tag on the gene in the chromosome.  Test whether cells with this tag arre still able to take up DNA.  But a test for DNA crosslinking might be worth doing even if the tag does interfere with function, if the protein is still assembled into a pore (secretin) or pilus (pilin).  So we should also test retrieval of the tagged secretin or pilin after formaldehyde crosslinking,with and without reversing the crosslinks before running the SDS-PAGE gel.  This will tell us whether the protein is assembled into its normal complex.
• Incubate the mutant cells with 32P-labelled DNA (probably the 222-bp USS-C fragment).  The standard way to form DNA-protein crosslinks is by adding formaldehyde, but this has the BIG disadvantage of also forming protein-protein crosslinks.  We hope to instead purchase photoaffinity nucleotides and incorporate one or more of these into the DNA we give to the cells.  This will allow us to specifically induce DNA-protein crosslinks (no protein-protein crosslinks) by irradiating the cells with UV (do we need a UV laser?  I think a colleague has one.)  After a very short time (1 minute?) UV to form crosslinks.
• Wash the cells to remove the external DNA, then lyse them and load them on a Ni-NTA resin column to bind the His-tagged protein.  Wash the column to remove everything else.  Elute the protein and check whether (i) the expected protein has eluted and (ii) any radioactivity has eluted.  If the protein is there but the radioactivity isn't, there was no crosslinking between this protein and the DNA.  If radioactivity elutes with the protein, investigate further.
• This experiment needs several controls.  The most important is probably a positive control for crosslinking of the DNA with a known protein.  How about SSB - it binds single-stranded DNA in the cytoplasm?  or DprA - it binds incoming DNA and protects it from nucleases?  A good negative control will be cells that aren't UV'd, as will DNA without the photoreactive nucleotide, and cells without the protein tag.

2.  Identifying other strong candidates (2 methods):

1. We will do simple transformation and DNA uptake assays on all the non-polar mutants.  These are already standard in our lab, and can be done in one term by an undergraduate or M.Sc. student.  Only proteins that are needed for normal DNA uptake will be retained as candidates.
2. We will use formaldehyde crosslinking , followed by gel electrophoresis and HPLC-mass spectromtry to identify proteins that are crosslinked (directly or indirectly) to DNA.  Any proteins not crosslinked will not be strong candidates.  The DNA will be tagged with biotin so that it and all crosslinked proteins can be recovered by attachment to streptavidin-coated magnetic beads (Dynabeads), and after recovery the crosslinks will be reversed so the proteins can be identified.  This is not a very specific test for uptake proteins, as it will also give proteins that bind incoming DNA in the cytoplasm, but missing proteins can be safely excluded.

3.  Characterizing the proteins that crosslink specifically to DNA:

Here;'s where the planning peters out a bit...

What do we do with these proteins once we've found them?  We test them for in vitro DNA binding, using bandshift and Southwestern assays. 

What if secretin and pilin don't crosslink to DNA???? Does this mean that they don't contact DNA at all?  This would be quite surprising.

Bacterial pseudogenes and within-species diversity

Last night Jon Eisen posted about a new paper by Chih-Horng Kuo and Howard Ochman, about the evolutionary fates of bacterial pseudogenes (PLoS Genetics: The Extinction Dynamics of Bacterial Pseudogenes).  I don't (yet) understand their conclusion very clearly, but it ties in well to the issues around the diversity of bacterial competence that I need to sort out for my CIfAR talk next week.

What do we know about within-species genetic diversity in bacteria?  The big issue is core genome and accessory genome.

In most (all?) species, different strains have a core set of genes in common; usually these make up about 80% of each strain's gene set (typical range ~70%-90%).  These core genes are usually syntenic.  They are very similar across the different strains, usually no more than a few percent different in DNA sequence, and almost identical in protein sequence, consistent with recent descent from a common ancestor.  These shared-by-descent genes are what justifies grouping the strains as representatives of a single 'species'.

The rest of each genome gene set comprises genes that are absent from some or most other strains.  It's not just that the alleles of these genes are very divergent, but that the genes have different ancestries.  Many of these accessory genes are in large blocks ('islands') with evidence of a mechanism by which they have been transferred from another distantly related species (e.g. phage, integron or transposon sequences, flanking tRNA genes).  This within-species genetic diversity is not seen in typical eukaryote genomes, perhaps because of the homogenizing effect of meiotic sexual reproduction.

Also unlike most eukaryote genomes, bacterial genomes usually contain only a small amount of non-gene sequences, usually about 10% of the genome.  This is almost entirely intergenic; introns are very rare and usually contain other genes (excisionases and mobilization genes).

What about pseudogenes?  Pseudogenes are DNA sequences that are closely related to functional genes but have mutations that destroy the function.  They are usually identified by comparison with the functional sequence in a close relative ('allele' if in the same species, 'homlog' if in another species).  Although function could be destroyed by mutations that change one or more critical amino acids, these can't be recognized without biochemical characterization of the gene product, and in practice pseudogenes are identified by the presence of a stop codon or indel that would prevent translation into a full-length protein.

Bacteria do have pseudogenes; in 2005 Lerat and Ochman examined 11 genomes from 4 genera and found that1%-8% of the open reading frames were pseudogenes.  Most pseudogenes were unique, defective in one genome and apparently functional in the genomes of close relatives, but some pseudogenes were shared between several Staphylococcus pyogenes strains and between two Vibrio vulnificus strains, and two were shared between the closely related V. vulnificus and V. parahaemolyticus.  Because shared pseudogenes were uncommon the authors concluded that old pseudogenes are rare.

The new paper examines the evolutionary histories of pseudogenes in five strains of Salmonella.  The strains all did have pseudogenes, from 0.3% to 3.7% of their functional genes.  All but 32 of the 378 pseudogenes identified had only a single defect, suggesting that they had arisen recently.  Consistent with recent origin, very few pseudogenes were shared (maybe 3?).  Most were created by small deletions or by point mutations that created stop codons.  The authors don't explicitly consider the core gene/accessory gene distinction, but because the pseudogenes were identified by alignment of not just the gene itself but of the genes flanking it on both sides, I think these are pseudogenes of the core gene set common to all strains, not of accessory genes present in only one or two strains.  (I just emailed the authors to check this.)  Many of the genes have no assigned or suggested function.

Kuo and Ochman ask why there are not more old pseudogenes.  But first I want to consider the basics -  what we might expect to happen after the first mutation happens.  If the functional gene makes an important contribution to fitness, we expect cells with the mutation to die or be quickly outcompeted by other cells, so the mutation will be gone from the population.  These pseudogenes are so short-lived that they are unlikely to be present in sequenced genomes.  If the functional gene makes little or no contribution to fitness in the present environment, the mutant cells may persist and even found a lineage (or, more likely, still go extinct).  The pseudogenes that are detected in sequenced genomes must be of this type.  Because the pseudogene's sequences are no longer under selection for the coding function, additional mutations that change its sequence may be selectively neutral, or they may be beneficial if they eliminate a harmful effect of the pseudogene.  What could such harmful fitness effects be?  The non-functional gene could produce a toxic product, being translated into a defective protein that interfered with the regulation or function of other proteins.  It might be transcribed but not translated, using up transcriptional resources.  Even if it is never transcribed, the cells still has to replicate and maintain this DNA, and it's often thought that bacterial cells have compact genomes because selection favours deletions of nonfunctional DNA that reduce this burden.

Kuo and Ochman conclude that(from the Abstract)

We found that after their initial formation, the youngest pseudogenes in Salmonella genomes have a very high likelihood of being removed by deletional processes and are eliminated too rapidly to be governed by a strictly neutral model of stochastic loss. Those few highly degraded pseudogenes that have persisted in Salmonella genomes correspond to genes with low expression levels and low connectivity in gene networks, such that their inactivation and any initial deleterious effects associated with their inactivation are buffered.

There are two points here, one I agree with and one I don't.  I agree that most pseudogenes are of recent origin, and their results do suggest that genes that are highly expressed and/or well connected are less likely to persist once they become pseudogenes.  The Discussion emphasizes the toxic-product hypothesis, which makes sense.

But I don't agree that deletion must be the reason we see few old pseudogenes in genome sequences.  It's true that deleting a pseudogene will eliminate both any toxic-protein cost and the cost of maintaining the unneeded DNA.  But it doesn't eliminate the cost of the original mutation that created the pseudogene.  Unless we have independent evidence that the DNA of pseudogenes is removed from genomes by deletion, we should probably suspect that instead cells carrying pseudogenes are removed from populations by selection.

Bottom line:

Is the DNA of new pseudogenes quickly lost from genomes by deletion, creating strains that are more fit than those with the pseudogene (but probably not more fit than the ancestor with the functional gene)?  This predicts that sequenced genomes should contain many sites where 'core' genes have been deleted.

Alternatively, are cells containing new pseudogenes quickly lost from populations because the cells compete poorly with cells that retain the functional gene?  This predicts that sequenced genomes will typically all contain the same core genes.

The figure below shows what we might expect to see when comparing 5 closely related genomes under each hypothesis  The orange parts of each bar represent genes that are intact in most genomes but are a pseudogene in one genome.