Tweezers-prep progress

1.  I tested cell adhesion to the coverslips the RA had cleaned for me - it wasn't dramatically better or more consistent than either uncleaned or acid-washed coverslips (after all had been coated with poly-L-lysine using his spreading method.  Overall plenty of cells are adhering, given that I'll only need a small number of cells adhered for each chamber.  I think part of the trick was inverting the slides for 10 minutes before washing away the nonadherent cells, so the cells settle on the coverslip.  I'm also giving the slides a few brisk taps during the washing, so that loose cells detach and wash away.

2.  Incubating the chambers with concentrated BSA protein didn't significantly reduce the number of H. influenzae or B. subtilis cells that stick to the poly-L-lysine coated coverslips.  Nor did it prevent the polystyrene beads from sticking to the coverslips.  A Google search for treatments that might block/inhibit the poly-L-lysine surface revealed only that people are coupling poly-L-lysine and polyethylene glycol or other materials into 'block' copolymers, and testing the ability of poly-L-lysine to 'inhibit' cell growth and various enzymatic reactions.  I didn't test BHI directly, but cells growing in BHI did adhere to the coverslips so I don't think it's much of an inhibitor either.  I don't know how big a problem the unwanted sticking is going to be, but I'd welcome suggestions for other possible blocking agents to test.  Just for fun I'm going to test milk.  Not reconstituted powdered skim milk, but whatever's in the food fridge (2% milk, I think).

3.  I made my big batch of competent cells and froze 34 tubes (0.5 ml), as well as 14 tubes of log-phase cells.  But I haven't yet tested how competent they are.

4.  Several tests showed that the H. influenzae cells that are stuck to the coverslips are otherwise healthy.  In the first test I filled the chambers with sBHI plus low-melt agarose and incubated them overnight.  I didn't seal the ends of the chambers but put the slides in a humidified box, but the chambers dried out completely overnight.  But I could see that there had been lots of cell doublings before the medium dried up.  So next I tested sealing the chamber ends with nail polish.  The tweezers lab people use wax from a candle, because they worry that the nail polish might be toxic, but I didn't have a candle.  The nail polish is indeed toxic (probably because acetone is quite water-soluble); the cells close to the ends of the chamber didn't divide at all.  But the cells in the middle part of the chamber grew well, producing nice tight microcolonies by the end of the day.  I brought in a candle, and today I'll see if I can improve my skill at applying melted wax to chamber ends.

5.  I think today I'll also test whether the cells on the coverslips can be transformed, by washing in MAP7 DNA, and then DNaseI and then sBHI agarose with added novobiocin.  Provided I have thousands of cells stuck to the coverslip, I should be able to find rare transformant colonies.  I'll try this with B. subtilis too, selecting for Trp+.

Uptake sequence manuscript done, back to the tweezers experiments

Yesterday I finished revising all the components of our manuscript on uptake sequence evolution, and sent them to the co-author who's handling the resubmission.  it was already a good manuscript, but now it's very very good, so if the editor and reviewers don't like it we'll just sent it somewhere else.

I've been working on the components of the optical tweezers experiments, with modest progress.  The tweezers apparatus still had problems when I was there the other day (it wasn't aligned properly?), but I have lots of other parts to work on.  Here's a list:

• The RA of the tweezers lab showed me a way to pre-clean coverslips, and I need to test the ones he prepared to see if they give more consistent results than the ones I've been using. (Coat his and mine with poly-L-lysine, make into chambers, test binding of competent cells.)
• I need to test whether washing treated coverslips with culture medium (BHI) or protein (BSA) blocks cell and bead binding, because once I have cells bound to the surface I don't want beads to stick to it.
• I need to make a big batch of competent cells and freeze them in 0.5 ml aliquots, because the cells I made before were contaminated.  (The contaminants were flagellated cells that otherwise looked just like H. influenzae - they were the spinning wiggling cells I described in my last post.)
• I need to test whether cells that have stuck to a coverslip are still able to grow.  To do this, my plan is to replace the medium in the chamber with medium containing low-melt agarose, and once this has set incubate the chamber at 37 °C for a few hours and then examine the cells under the microscope, looking for clusters of cells (microcolonies).  As a control I'll first try this with cell-free chambers and agarose medium that already contains growing cells.
• For today I'm just going to work with H. influenzae, but I should also test these factors with B. subtilis cells.

Haemophilus doesn't have flagellae!

I'm still testing treatments to get cells to stick to coverslips (with more success):

I used more poly-L-lysine this time, but the important thing I realized was that I need to tap the slides briskly to loosen the cells that are just sitting on the surface but not attached,a nd to rinse a lot of medium through the chamber.  This let me see that there were usually substantially more B. subtilis cells attached to the coverslip than to the glass slide.

So then I tested whether my competent H. influenzae cells would also bind to the coverslips.  They did, better than the B. subtilis cells.  I could see that many of the cells were only attached at one end, with the other end moving in the medium.  But what was surprising was how vigorously the free parts of the cells were moving around.  Not just bouncing back and forth (can Brownian motion be that vigorous?), but sometimes spinning around almost like cells tethered by their flagella (see this movie, which I think is Salmonella but might be E. coli).

But H. influenzae definitely doesn't have flagella.  So I'm plating the cells I have onto LB and BHI plates with and without hemin and NAD, to check that some flagellated imposter hasn't snuck in to my culture.  (It would have to make colonies that look just like H. influenzae colonies, and be sensitive to novobiocin...)

Sticking cells to coverslips (one step forward, two steps back?)

I coated some coverslips with poly-L-lysine, using the rub-until-dry method I was shown.  I then assembled the coverslips into 'chambers' like those described here, and tested whether competent B. subtilis cells would stick to them.

I wanted to use chambers for these tests because that's what I'll be using with the optical tweezers.  Another benefit turned out to be that I could compare how well cells stuck to the coated coverslip surfaces to how well they stuck to the untreated surfaces of the glass slides that form the bottoms of the chambers.

I let the cells sit in the chamber for 5-10 minutes at room temperature, and then washed them out by flowing about 10 volumes of medium through the chamber, introducing it in tiny drops from a pipette tip at one end and absorbing the flow-through with a square of blotting paper at the other end.  Then I looked at the chamber under the microscope, comparing the cells on its upper and lower surfaces.

In most tests I saw little or no difference between the treated and untreated surfaces; quite a few cells were stuck on both.  The largest volume of poly-L-lysine solution I used did give some patches where many cell stuck, but these were at the edges of the coverslip, where the rub-until-dry treatment hadn't reached well.  Cell density (on both surfaces) was also generally higher at the edges of the chambers - I think this may just v=be because the washing is less effective at the edges.

I also tested my DNA-coated beads.  They didn't stick any better than the cells did, which is good.  But there were lots of clumps of beads - perhaps I didn't vortex them well.

Changes/improvements:

In a previous experiment I had better results with coverslips that had been presoaked in acid alcohol, so I'll try this again.

I'll try higher concentrations of poly-L-lysine.

I'll try using more dilute cells.

I'll try H. influenzae cells as well as B. subtilis cells, because their surfaces have very different chemistries.

I'll try incubating the chambers+cells upside down before washing the cells out, so the cells will settle on the coverslip rather than the slide.  Though, for the tweezers experiments, it doesn't matter which surface the cells are stuck on.

I'll try marking a reference spot on the coverslip, so I can track whether cells present after the first wash are removed by the second one.

I'll reread the papers that did tweezers studies of competent cells, and email their authors for advice.

Transformation by DNA on beads

Yesterday I incubated some of my competent H. influenzae cells with DNA carrying a novobiocin-resistance gene.  One tube got the normal prep of chromosomal NovR DNA.  Two others got DNA that had been cut with one of two restriction enzymes (either EcoRI, average fragment size 6 kb or XhoI, average fragment size 12 kb) and then had biotin incorporated at the ends of the fragments, and three others got streptavidin-coated polystyrene beads with the biotinylated DNA bound to them.

I had calculated that the first batch of DNA+beads had about 250 ng of DNA per ml, so I used the same volumes of beads for the others, and diluted the non-bead DNAs to 250 ng/ml before using the same volumes.  I incubated the DNA plus cells for 15 minutes and then plated the cells on Nov agar, and (diluted) on plain agar.  Today I counted the colonies and calculated the transformation frequencies.

Point 1.  Cutting the DNA with XhoI reduces its transforming ability by 4-fold, but cutting with EcoRI reduces it by about 100-fold.  So I checked where these two enzymes cut relative to the NovR (gyrB) gene.  EcoRI cuts inside the gene, but XhoI only cuts at sites 1 kb and 3 kb on either side of it. 

Point 2.  DNA attached to beads transforms!  This wasn't a sure thing, for lots of reasons.


Point 3.  DNA attached to beads transforms 10-30-fold worse than the same amount of free DNA.  This is not surprising because most of the DNA on the beads will be inaccessible because it's tangled up with or behind other DNA fragments.

Now I'll celebrate by spreading some poly-L-lysine on some cover slips, using the new method I was taught yesterday, so tomorrow I can test whether cells bind to these and whether they stay alive after binding.

Tweezers progress

Yesterday I took my new batches of frozen competent cells (B. subtilis and H. influenzae) across town to the biophysics lab, so I'll have cells there to test without needing to make fresh ones.  While I was there I learned a better way to coat cover slips with poly-L-lysine, which I hope will give more reproducible attachment of competent cells.  And I spent more time making microscope-slide chambers and learning about the tweezers apparatus (mostly standing by while the expert grad student made adjustments to the optics and electronics).

I also attended a seminar about DNA bending.  Short fragments of double-stranded DNA (~100 bp) have been reported to circularize much more efficiently than predicted by their expected persistence length (paper by Cloutier and Widom), and one proposed explanation is the formation of tiny 'bubbles' in the DNA structure – places where several base pairs have separated although the DNA backbones remain intact.  Even though such short single-stranded regions are expected to be very transient they can have a big impact on the probability that the ends of the DNA will meet, allowing DNA ligase to join them and circularize the DNA.

Blocking with BSA makes a big difference

The streptavidin-coated magnetic beads that hadn't been blocked with BSA aggregated into big clumps when mixed with cells, but the treated beads remained separate.  The clumping was independent of whether the beads had been incubated with biotinylated DNA, so I think the clumping was because most cells stuck to more than one bead and most beads stuck to more than one cell. 

So then I checked the streptavidin-coated polystyrene beads I'll use for the tweezers experiments.  Luckily adding cells didn't make them clump.

Now I'm making fresh batches of competent H. influenzae and B. subtilis to freeze, so I'll have consistent cells to work with.

Do cells bind to DNA stuck on beads?

I need to plan the experiment(s) where I test whether competent cells bind to DNA on magnetic beads.

I have 1 ml each of two types of 1µ and 2.8µ magnetic 'dynabeads'.  The beads have been coated with streptavidin, and one batch of each size has then been blocked with BSA, which apparently reduces the surface charge and makes them better for binding protein but not so good for nucleic acids and I think more prone to clumping in high-salt buffers.  I don't know how the different surfaces will affect non-specific cell binding, something I want to avoid (Invitrogen recommends  including 0.01-0.1% Tween 20 to reduce non-specific binding, where that won't interfere with the assay).  And I don't know how the surface properties of these  beads compare to the properties of the polystyrene beads I'll be using for the tweezers experiments.  (I need to keep reminding myself that I'm only using the magnetic beads to check whether cells will bind to DNA on beads, so I shouldn't waste a lot of time optimizing the assay.)

I can separate the beads from 150 µl of liquid by simply drawing the mixture up in a pipette tip, holding the tip in the nanobead magnet rack for 10 seconds, and slowly expelling the liquid; almost all the beads remain behind on the side of the tip.  I can then resuspend the beads by drawing clean buffer up into the tip, away from the magnet.

So:  First mix one aliquot of beads with DNA (Invitrogen says to do this in TE + 1 M NaCl).  To start I'll just use one size of beads, arbitrarily the 1 µ ones.  I know that the stocks contain 10 mg beads per ml, but I don't know how many beads this is.  I was going to find that out, by diluting some beads and looking at them under the microscope (we have a hemocytometer).  But here's a rough calculation:  If the density of the beads is a bit higher than that of water, then a single bead 1 µ in diameter will have the same mass as a 1 µ cube of water, which is 10^-9 mg.  So 10 mg of beads/ml is ~10^10 beads/ml.  As I did with the polystyrene beads, I'll wash the beads several times to remove unbound DNA.  I don't think I need to check that DNA has bound by using the sensitive fluorescence assay - I'll leave that to do if I don't see a difference in cells associated with beads with and without DNA.

Then I'll thaw some frozen competent H. influenzae cells, wash away the glycerol and resuspend them in BHI.  Then I'll  mix them with DNA-treated beads and control (no DNA) beads, at a concentration of, say,  ~ 5x10^8 cells and beads per ml.  I'll incubate cells plus beads briefly (1 min at 37°C?) and then wash the beads three times, saving the eluate/supernatant/whatever it should be called.  Then I'll plate the beads (several dilutions) and the eluates.  I'll also look at the beads under the microscope (maybe do that first).

1. Make washing buffer, wash beads, resuspend in BHI, in MIV and in PBS.  
2. Check under microscope to see if they're clumping.
3. Incubate beads with DNA, wash well.Thaw and wash cells.  Resuspend in BHI or MIV.
4. Mix cells with beads, incubate briefly and wash.
5. Assess binding by plating and/or microscopy.

Revisions almost done

Today I finally finished what I very much hope is the last major revision of the uptake-sequence variation manuscript.  I've rewritten half of the Introduction and all of the Discussion, using my new non-adversarial, everyone-wins framework.  I've pulled together all the data for the supplementary table that serves only to show how quantitative our results are.  I've redrawn one figure with new data (same results as the old data).  I've gone back through the Responses to Reviewers, changing the responses to match what we've now improved.  I've started to draft a cover letter to the Editor.

Then I emailed everything to both my coauthors and to the three people in the lab (postdoc, RA and visiting researcher), asking for only essential changes and polishing of the writing.  With luck it will be resubmitted sometime next week, and with even more luck the critical reviewer will find our changes acceptable.

Now I've run out of excuses for not cleaning up my office.

Framing the uptake sequence problem (Intro and Discussion)

I think the last post may have been a bit incoherent, but it led me to a new perspective on the problems posed by uptake sequences, one that I think gives a much better frame for the manuscript.

INTRODUCTION (new frame):

(After introducing uptake sequences and uptake biases...)

Why bacteria take up DNA is controversial, and presence in two bacterial groups of DNA uptake sequences and their associated uptake biases pose problems for both major hypotheses.

It's generally assumed that bacteria take up DNA to get benefits from homologous genetic recombination, and that uptake sequences plus biases are a mate-choice adaptation to maximize these benefits by excluding DNAs that are not from close relatives.  Although this is intuitively appealing, it is evolutionarily problematic, both because it requires simultaneous evolution of bias in the uptake machinery and genomic sequences matching this bias, and because the genomic sequences can only be 'selected' after the cell carrying them is dead.  (There's also the bigger problem that the presumed benefits of recombination are expected to be, on average, very small or nonexistent.)

The alternative hypothesis is that bacteria take up DNA as a source of nutrients (initially nucleotides but also carbon, nitrogen and phosphate), for which the very existence of uptake sequences plus bias is counterintuitive.  If DNA in the environment is valued only as nucleotides on a string, all DNAs should be equally useful.  Although the sequence bias might play a mechanistic role in DNA uptake (such biases are typical of proteins that bind DNA, even ones whose functions are sequence independent), the high density of the preferred sequences in the genome is perplexing.

The phenomenon of molecular drive may resolve the worst of these problems for both hypotheses, by providing a hypothesis-neutral explanation for uptake sequence abundance.

(Explain molecular drive here.)

If molecular drive is indeed an inevitable consequence of biased DNA uptake and homologous recombination, its action may remove the biggest obstacles for both hypotheses.  Below we use a computer simulation of genome evolution to test its requirements and consequences.

METHODS

RESULTS

DISCUSSION (new frame):


Summarize the findings.  They are robust.

What's been gained:  Proponents of the mate-choice hypothesis now need only explain how natural selection for the benefits of recombination would favour uptake specificity in the genes encoding the uptake machinery - the corresponding uptake sequences will inevitably accumulate in the genome as the specificity strengthens.  Proponents of the DNA=food hypothesis need only explain how sequence bias would evolve for mechanistic benefits in DNA uptake; uptake sequences in the genome can be ignored.

The above paragraph is really too adversarial a perspective.  It's now much clearer what information is needed to explain uptake specificity.  First we need a much more detailed characterization of the real uptake biases (Neisserial and Pasteurellacean).  Second we need to know what role uptake bias plays in the process of uptake in each organism.  Does a dedicated cell-surface protein pre-screen DNA fragments for uptake sequences before uptake is initiated?  Do uptake sequences provide structural flexibility for DNA bending or kinking during initiation of uptake?  Do uptake sequences play any role after initiation?  Do they affect DNA synapsis or other stages of recombination?

We also need new explicit models of the evolutionary forces that would act on uptake genes and preferred sequences.  Can selection for genetic benefits of recombination be strong enough to cause evolution of uptake bias?  Or, vice versa, can exclusion of unrelated DNAs reduce the costs of DNA uptake?  Analysis of protein sequences in genomes with and without uptake sequences suggests that their evolutionary costs are small, but a theoretical framework for this is lacking.  Because the model presented in this paper tracks only a single focal genome, it is not suitable for investigating effects on organismal fitness (whether due to the costs of uptake sequences or to the genetic benefits of recombination).

Discussion draft

(Here I'm just trying to lay out the philosophical framework of the Discussion.)

Uptake sequences might seem to be a minor trivial problem (only two relatively unimportant bacterial groups), but they  have important implications for the evolution of sex.  The key question is, are uptake sequences (= sequence-biased DNA uptake plus abundant preferred sequences in the genome) evidence of selection for benefits of homologous recombination?

Start with recap of what needs to be explained.  It's difficult to disentangle the hypotheses:  Are bias and accumulation the two interdependent components of an adaptation to promote uptake of fragments that can contribute to beneficial recombination, as often assumed?  The big problems with this are the weakness of the hypothesized benefits of recombination in current models of the evolution of sex, and the difficulty of selecting for sequences that only act after the owner is dead.  Alternatively, each component (uptake bias and uptake sequences in the genome) could have a separate explanation; for example, uptake sequences might have a cellular function, and sequence bias might play a mechanistic role in uptake.

But an acceptable model for bacterial uptake sequences must explain the very strong correspondence between the sequences that are preferred by the uptake machinery and those that are overrepresented in the genome.  Given an abundant sequence with a cellular function, might bias favouring it evolve because of genetic benefits of homologous recombination (mate choice), or because of mechanistic benefits of evolving high-affinity DNA-binding proteins specializing in a commonly available sequence?  Conversely, given an intrinsically biased uptake machinery due to mechanistic constraints or the need for high-affinity DNA binding, might the preferred sequences accumulate in the genome regardless of recombination benefits?

This last is the simplest hypothesis to test.  Our results show accumulation of uptake sequences like those in real genomes, provided only that DNA uptake is strongly biased and homologous recombination sometimes occurs.  The results don't require that the homologous recombination provide any genetic benefits.  However, the effects we have found will act regardless of whether the benefit to the cell comes from the DNA's nucleotides or its genetic information.  They also don't need selection for homologous recombination, as that occurs anyway in repair-capable cells

So now we have shown that, provided DNA is taken up and bias exists, uptake sequences will accumulate.  If DNA uptake is selected because it provides food, then uptake sequences have not evolved as markers of sequence homology, and we expect to find that uptake is biased for non-genetic reasons.  If DNA is instead taken up for its genetic information, we need to study how the accumulation of uptake sequences affects the genetic benefits, and how the benefits can affect the evolution of the gene causing the bias..

Our finding that uptake sequences accumulate without selection removes one of the problems with the uptake-bias for sex model (that of needing to simultaneously select for bias and uptake sequences.  Instead now just need to test whether uptake bias will evolve (gradually) because of benefits of homologous recombination, with uptake sequence accumulation following passively along.  Our model of uptake sequence evolution is not designed to evaluate this because it is not population-based (it tracks only a single focal genome).  (Can I mention here my new modeling of the effect of selection at one position?)

An alternative approach is to investigate the role bias plays in the uptake mechanism.  Finding that bias is created by a mechanism-independent protein that pre-screens sequences for uptake sequences would support the hypothesis that bias exists to promote homologous recombination.  Conversely, finding that bias makes a mechanistic contribution to the process of uptake would be consistent with non-genetic functions of DNA uptake.  Our present focus is on properly characterizing the true uptake bias of the H. influenzae uptake system,  identifying the gene or genes responsible for the bias, and finding out the role of sequence biases in the mechanism of uptake

Still (STILL!) working on the uptake sequence variation manuscript

(If the damned thing takes much longer we're going to have to ask the Editor for an extension!)

It's my fault - I'm stalled at fixing up the Discussion.

The problematic reviewer felt that the model didn't make testable predictions, so I wanted to include a brief discussion of how it could be used to evaluate more complex hypotheses about uptake sequence evolution.  Unfortunately, a proper test that includes selection will require the model to follow a population of cells or genomes, rather than a single focal genome.  Such a model would necessarily be  more complicated than ours, and if it was set up like ours the run times might be prohibitively long.  (Of course a clever programmer might find ways to streamline it without losing scientific relevance.)

But I thought of a simple test of whether uptake sequences accumulate near to positions that are under fitness selection in the diverging sibs of the focal genome.  Modifying the program to do this took only about 12 lines of code, but getting this code to work properly took me most of a day (spent chasing curly brackets and finding out the correct way to use the Perl 'substr' function).

This new version of the program includes selection at position 5000 of the evolving genome.  As before, each DNA fragment in the environment is first scored for sequences matching the uptake motif, and the resulting score determines its probability of recombining with the focal genome.  But now, if the fragment overlaps position 5000, it is also checked for its base at that position.  Fragments with an A at position 5000 keep their original uptake-sequence score for the recombination step, but fragments with the other bases have their scores reduced by a factor of 0.7, 0.4 or 0.1 (for G, C and T respectively).  This is intended to simulate poor survival of cells with these bases.  If the focal genome has an A it will have a near-normal recombination around position 5000 (except for the 1/100 fragments that carry mutations to less favoured bases), but if it has one of the other bases it will have reduced recombination except for the higher recombination of fragments carrying mutations there).  Recurrent mutation at position 5000 (in the focal genome and in the divergent fragments) may create a recurrent benefit of recombination, and if uptake sequences promote beneficial recombination, might select for uptake sequences close to position 5000.  On the other hand, if recombination more often brings in harmful mutations, uptake sequences close to the selected position might be selected against.

So I examined the final locations of uptake sequences in genomes from a bunch of runs that started either with 10 kb random-sequence genomes or with 20 kb genomes pre-seeded with uptake sequences (one very close to position 5000).  In the random-sequence genomes there were just as many uptake sequences around position 5000 as anywhere else, and in the pre-seeded genomes the uptake sequence at position 4982 was no more and no less stable than any other uptake sequence.

This isn't a very good test, in lots of ways (in fact it's quite awful), but I think it will show the Editor that  the model is indeed testable, and that we have made a reasonable effort to satisfy the reviewer.  In the manuscript's Discussion I'll describe it in less detail than I have above, and I won't present any data ("Redfield, unpublished").  And I'll explain that a proper test that incorporates selection for beneficial alleles will require a population-based version of the model.

[I also still have to assemble some new data into a replacement for one of the figures, and to go back over the latest changes one more time before sending them to my coauthors on last time.]

 Some comments about my Evo-WIBO talk plan from a reader:

...what I'm really curious about is the sense I get that you feel a phenotype must be some sort of evolutionary goal (i.e., why would we have an a priori expectation that enzymes would evolve to accomplish homologous recombination?) Gender doesn't seem to pop onto the Natural History landscape full blown and ready to be appreciated. So why should HR? I really like the notion that HR might proceed from a DNA replication and repair background. 

I didn't mean to imply that phenotype is a goal, neither generally or with respect to homologous recombination (HR).  But most other microbiologists and molecular biologists have been assuming but not rigorously evaluating) that HR exists because of selection for its sometimes-beneficial consequences. 

And is it not possible that natural competence is currently an orphan process that exists for food uptake but was once a piece of a primordial sex process that developed further in other lineages but was cast aside in bacteria? (photosynthesis may have been cast aside in oomycetes in favor of parasitism).

That seems backwards to me, because selection for the food benefit is so straightforward and selection for sex so problematic.

To me, HR has to be more beneficial than horizontal gene transfer for a lineage to find it worth the trouble. When organisms are extremely simple the selective disadvantage of maintaining DNA that isn't carrying its weight should lead to its elimination. The notion of an allele implies the existence of a gene - but a gene not in the sense of a capable ORF but in the sense of two or more ORFs in a population that perform the same function in manner that the environment will influence and that selection can act on. If said variant ORFs come to be in the same cell, then HR can go to work on them.

Homologous recombination isn't really any 'trouble', to the extent that it happens as an accidental (i.e. unselected) consequence of enzymes selected for their effects on DNA replication and repair and of accidental transfer of DNA fragments by genetic parasites or of DNA uptake for food.  And in bacteria there's very little evidence that it ever occurs any other way.

The value of taking a different tack on a problem is prescient. And physics offers a host of tools and a philosophical background that could really help. To me the challenge of the 15 minute presentation is to illustrate how having data that describe the physical process of DNA uptake should allow mathematical model development for the process which then allows development of testable hypotheses. There are "big organism" examples of this approach bearing fruit.

I don't think that the phenomenon of natural competence needs mathematical models at all (nor do any other of the phenomena that sometimes lead to to recombination in bacteria). My point for the talk is that many hypotheses can be directly evaluated by more thorough investigation of the phenomena in question.

Defending 'functional design' analysis at Evo-WIBO

In a couple of weeks I'll be giving a short talk at the regional Evo-WIBO meeting.  My title is What's an evolutionary biologist doing in a physics lab?.  I think I'm going to combine a description of my specific scientific question (the physical properties of DNA uptake by Haemophilus influenzae) with a rehash of the defense of 'functional design' that I made in a post last month.  So I might subtitle the talk A defense of functional design analysis.

I'll only have 15 minutes including question time, so I'll need to keep it simple.

• The simple answer is, I'm measuring the physical properties of DNA uptake by the bacterium Haemophilus influenzae.  I'll show you how this is done at the end of my talk, with a nice explanatory animation.
  
• Why is this of evolutionary interest? Because it's one of the final pieces of the Do bacteria have sex? puzzle.
• Why aren't I using more evolution-style approaches, behaving like a proper evolutionary biologist?  How will knowing physical forces answer evolutionary questions?  Shouldn't I be using the comparative method?  Since these are bacteria, why aren't I doing Rich Lenski-style lab evolution experiments?
• A defense of 'functional design' analysis:
• Understanding 'natural history' (the stamp collecting side of biology?) is fundamental to investigating evolutionary forces.  Before we try to explain how natural selection has acted on any phenotype or behaviour, we first need a solid understanding of what the phenotype or behaviour is.
• First a big-organism example: The head-nodding lizards.  We can use the usual methods of natural history.  What does it do, when does it do it, what are the typical outcomes?
• Next, a bacterial example: For bacteria, we need to use the methods of molecular biology.  Consider RecBCD (3 proteins that work together).  How was it discovered, what was its function initially thought to be?  What was later learned about the phenomenon (not by evolutionary biologists).  Molecular biologists often treat both 'functions' as equivalently important.  How should evolutionary biologists think about it (consider relative strengths of selective forces).
• Similar history of thinking about nearly all the genes that contribute to homologous recombination in bacteria.  The molecular biology isn't my work, but I spell out the implications for evolutionary biologists.
• Main conclusion:  Many (and perhaps all) bacteria don't have 'sex'; that is, they don't have any genes that evolved to promote homologous recombination with alleles from other cells of the same or closely related species.  Two of the three processes that move DNA from one cell to another are caused by genetic parasites, and the genes responsible for the physical recombination all have important functions in DNA replication and repair.  True of E. coli.
• I say 'perhaps all' because the function of one of the three processes that move DNA is still controversial.  That's natural competence

How best to test binding of competent cells to DNA on beads?

Now I have lots of biotinylated DNA, and a well-tested procedure for binding DNA fragments to streptavidin-coated styrene beads, I'm ready to test whether competent bacterial cells (B. subtilis or H. influenzae) will bind to the DNA on the beads.

How to do this isn't straightforward.  One problem is that the beads are about the same size and density as the cells (B. subtilis cells a bit bigger, H. influenzae cells a bit smaller), so once mixed they can't be easily separated.  That means I have no way to wash unbound beads away from cells, or unbound cells away from beads.  Another problem is that B. subtilis cells are known to cut DNA fragments as part of the uptake process, and in principle this might terminate uptake.  Though maybe not, as the cutting is part of the process that initiates uptake across the inner membrane.  H. influenzae cells don't cut DNA.

I could just mix competent cells and DNA-coated beads, both at low densities, on a microscope slide and watch for them sticking to each other.  Alternatively, we have some streptavidin-coated paramagnetic beads I could use - this would allow me to pull out the beads and see if cells had stuck to them.  But these 50 nm (super-tiny) beads, too small to see individually, so I'd have to plate them to see if there were cells there.  We might also have some micron-sized ones; I'll look around. 

OK, I found our 'starter kit' of 1 and 2 micron paramagnetic beads.  The only problem is, we were too cheap to pay the $150 for the starter version of the magic magnetic rack that holds microfuge tubes against magnets so the beads stick to the side and the liquid can be removed. So I tested various magnets from around the lab, and all of them pulled the rusty-brown beads to the side of the tube in a couple of minutes.  Doing this in a way that holds the tube steady so I can remove the liquid...not yet.

I'm away for a few days, but when I get back I may send an email out asking if anyone in the building has a Dynabeads rack I could borrow for a little while.

How much DNA is on the beads?

The NanoDrop tech support person said that the styrene beads wouldn't hurt the NanoDrop spec, and agreed that light scattering might be a problem.  It was, and that combined with the detection threshold of the Nanodrop meant that my measurements didn't give any evidence of DNA on my beads.  So today I used the PicoGreen assay to look for DNA on the beads.  It's much more sensitive, and not bothered much by light scattering due to the beads.

But first I should describe what my samples were and how I made them.  I incubated some 1.26 µ streptavidin-coated beads with a diluted solution of my biotin-labeled DNA, diluted because I didn't want different beads binding to the two ends of a fragment, and I didn't want steric interference by the DNA on the beads.  I incubated the beads with the DNA for 30 minutes, gently mixing at 37C on our roller wheel.  Then I pelleted the beads, and washed them twice with 1.0 ml of TE, each time rolling the beads plus TE for 10 minutes, and resuspended the washed beads in 100 µl TE (call these Beads1).  I also added another aliquot of beads to the DNA solution I'd already used with the first beads, and put these beads through the same incubation, washing and resuspension steps (call these Beads2).

Beads1 and Beads2 had very similar DNA concentrations, about 250 ng/ml.  This isn't very much DNA (but see below), but because they're the same I know that the low binding isn't because my biotin-labeling failed.  If the labeling had been the problem, then Beads1 would have had little DNA because they had bound up all the labeled DNA in the tube, and Beads2 would have had much less DNA. (I could check this by incubating more beads with the same DNA sample.)  Instead the low labeling may be because of the amount of strepavidin on the beads, or its reduced accessibility once bound DNA fragments are getting in the way of other DNA fragments.

So how much DNA is this per bead?  Here's a very back-of-the-envelope calculation:  The bead concentration in the resuspended Beads1 and Beads2 preps is about 0.1%, assuming that no beads were lost in the washing steps.  Let's consider 1 ml of Beads1 (or Beads2), just because it makes the arithmetic clearer.  With 0.1% beads, 1 ml of Beads1 solution is about 1 µl of packed beads (and yes, that's about how big the pellets appeared).  The beads are about 1.25 µ in diameter, and 1 µl is a cube that's 1000 µ on each side, so 1 µl of packed beads is a cube with about 800 beads per side, or about 5x10^8 beads.  At 250 ng/ml, the same ml of Beads1 contains about 250x10^9 kb of DNA (using Rosie's universal constant of 10^18 kb/gram of DNA).  The average fragment size of EcoRI-cut H. influenzae DNA is about 6 kb, so this is about 42x10^9 fragments.  I conclude that the average bead has about 85 DNA fragments bound to it.  That's pretty reasonable for my experiments, so I can go ahead and use these beads and this DNA to test cells binding to DNA on beads.

I also measured the DNA concentrations in the two washes from each aliquot of beads.  The first washes had about 20 ng/ml DNA, and the second washes had fluorescences not significantly higher than background, so I know that the signals from Beads1 and Beads2 were due to bound DNA.

One control I didn't do was to make a standard curve using known amounts of DNA mixed with 0.1% beads.  I should try this tomorrow.  I've also saved the samples I measured, and I'll also try reading them again tomorrow using the high-sensitivity setting of the plate scanner.  (Later - I was wrong; there is no high-sensitivity setting.) That's if I can figure out how to do this; the scanner software is very non-intuitive, and so far I've spent most of my time trying to find files I thought I'd saved.

At the bench today

Today I did two reactions that labeled the ends of digested chromosomal DNA with biotin (one of EcoRI-digested DNA and one of XhoI-digested DNA).  The next step is to clean up the DNA, to get rid of the Klenow polymerase and the EcoRI/XhoI and the unincorporated nucleotides.  it's especially important to get rid of ALL of the unincorporated biotin-dUTP, because this will otherwise bind to the streptavidin-coated beads and prevent the biotinylated DNA from binding to them.

Because I want to wash the DNA well to get rid of the biotin-dUTP, a column cleanup is best.  We have new cheap cleanup columns from a company called Epoch, to replace the relatively expensive Sigma genelute columns we've been using.  (These in turn replaced very expensive columns from Qiagen.)  What makes the Epoch columns such good value is that instead of charging for little bottles of salty water like the other companies (their 'secret sauce' buffers), Epoch just provides the recipes so users can make their own buffers.

The RA had already tested the new columns with a PCR cleanup, but I needed to test them with large fragments of chromosomal DNA, because DNA fragments bigger than 10-20 kb tend to stick to this kind of column.  Bottom line: both columns release almost all the DNA fragments smaller than 20 kb. 

I also wanted to compare the overall recovery of DNA from the columns, especially if they were heavily loaded with a lot of DNA (their stated binding capacities were either 10 µg or 20 µg, depending on which document I read).  But my DNA wasn't as concentrated as I thought, so the most DNA I put on a column was probably about 14 µg.  Recoveries were good, >80% even with more than 10 µg on the column.

So now I have about 35 µg of biotin-tagged EcoRI-cut chromosomal DNA, and about 25 µg of biotin-tagged XhoI-cut chromosomal DNA.  The next step is to measure binding of this DNA to the streptavidin-coated beads.  I can do this accurately now that the RA has shown me how to use Picogreen to measure very low DNA concentrations, and I've checked that beads don't interfere with these measurements.  I wasn't sure if I could put samples containing beads onto the NanoDrop spec, but I just read their explanation of how it works and I don't see any problem.  Maybe I'll email their Customer Service people just to be sure, as the NanoDrop we use belongs to the lab next door.

What's up with the manuscript about uptake sequence variation?

We're revising it, though not drastically.  One of the reviewers didn't have many concerns, but the other was full of philosophical objections, which we're meeting with calm reason and more analysis.

One bit of data we'll now include is the density of uptake sequences in the equilibrium genomes we discuss.  But when I went back to extract this data from the appropriate runs, I found that one run didn't have the data because it hadn't terminated when it was supposed to; there was a typo in the specified termination cycle (2000o0 rather than 200000), so it would have kept running forever if I hadn't stopped it.

And when I went to redo that run without the typo, I discovered that the set of 12 runs it belonged to had all had another error; instead of recombining 1000 fragments each cycle they had only recombined 100.  Fixing this won't change the conclusions at all; the runs will just all converge on a modestly higher score.   So I requeue'd all 12 runs, and then requeue'd them all again to terminate after 50,000 cycles rather than 200,000, because with ten times more recombination per cycle they may not need nearly as many cycles.  I was thinking that having more recombination would let them run faster, but I forgot that, with more recombination, each cycle will take longer.  Hmm, maybe I should even set them for only 20,000 cycles.   I'll see how far they've gotten tomorrow morning.

Are the purR knockout mutants not really purR knockout mutants?

The meticulous RA thought it would be wise to use PCR to check the genotypes of the purR::kan knockout mutants I used for my time course last weekend.  (I had already checked that they were both resistant to kanamycin.)  So she designed and ordered some primers that would flank the insertion that was described in the notebook of the grad student who originally made the mutant, and did colony PCR on all four of the strains I had used.

Much to my surprise, all four strains produced bands of the size expected for purR+ cells (about 1.0 kb), and none of them produced bands of the size expected for the purR knockout (about 2.2 kb).  Either there's something wrong with the PCR analysis (and she's very meticulous so I doubt that), or the strains aren't what we've been thinking they are.

I had made these strains by transforming cells with DNA I had isolated from cells grown from the old frozen stock of purR cells made by the grad student (at least, that's what I thought I was doing), and selecting for kanamycin-resistant transformants.  Could I have used the wrong DNA?  Or grown up the wrong cells from the freezer?

We know that the original cells made by the grad student had the correct mutation, both because he had carefully checked them out and because a technician had later thawed a vial and done a microarray analysis of RNA.  This showed that the mutant dramatically overexpressed all the genes that were predicted to be repressed by PurR in wildtype cells.

So tonight I've streaked out more cells from the last freezer vial of the original purR knockout, and on Monday the RA will test them by PCR.  I also located the DNA I had used for that transformation, so she can test that by PCR too.  If these cells give the expected 2.2 kb band, we'll assume something went wrong with my transformation.  If they give the 1.0 kb band, we'll carefully check out the new PCR primers and probably run a quantitative PCR of a PurR-repressed gene on RNA from the original mutant and from one of the new mutants (with wildtype cells as control).  Or, because the RA has recombineering working well now, she might just remake the purR mutant with her new primers.

If the mutants I used for my time course turn out to not be purR-, I think we'd still be really interested to find out where their kanR cassette is, because we don't have any other mutants with this interesting phenotype.  That can be done by cloning out the kanR cassette and flanking sequences (the old-fashioned way or by inverse PCR) and then sequencing the DNA on one or both sides of the cassette.

What on earth is 'constructive neutral evolution'?

Ford Doolittle gave a talk here today in the Biodiversity seminar series, which is attended by all the evolutionary biologists.  It was titled 'Irremediable Complexity', and was promoting a concept originally published by Arlin Stoltzfus under the title 'On the possibility of constructive neutral evolution' (here, but probably behind a paywall).  I haven't read it but it's been more influential than Ford said, cited 103 times.

Arlin's title is not at all self-explanatory; here's what I now think the words are intended to mean:  'Evolution' means 'a change over time in how a function is accomplished'.  'Constructive' means 'the change is that the function is accomplished in a more complex way'.  And as a result of some helpful questions at the end, I now think that 'neutral' means 'the function itself is under stabilizing selection but not under adaptive/directional selection, and how it is accomplished (the change in complexity) is not under selection at all'.  Ford didn't define 'complexity' until the question period; he then suggested that one measure of a function's complexity might be the number of components required for it.

Here's the executive summary:

Once organisms have evolved to have many components, some components will inevitably interact with others in 'accidental' ways that have, at least initially, not been shaped by selection.  Once these accidental interactions exist, they will modify how selection acts on mutations that affect the function, sometimes making things worse but sometimes mitigating the effects of what would otherwise be deleterious mutations eliminated by selection*.  These mitigating effects will weaken stabilizing selection on the function, sometimes allowing the mutations to be preserved (especially if populations are small).  Preservation of the mutation effectively creates selection for maintenance of the formerly-unselected interaction.  The function has become more 'complex (by Ford's definition), but there hasn't been any selection for the complexity.  If mutations with these kinds of effects recur repeatedly, the function will become increasingly complex without having been in any way improved.

As evidence that this type of complexity-building is common and important, Ford cited several molecular examples where a process has become ridiculously ('stupidly') complex but doesn't work any better that simpler versions.  The RNA editing of trypanosomes is not well known but is a compelling example.  So are introns and the spliceosomal machinery that lets eukaryotes cope with them.  Simpler examples are the 'maturation proteins' that assist type I and II self-splicing introns.  The ribosome itself may be a (not very stupid) example, where proteins have gradually taken over activities originally handled by the catalytic RNAs.

The issue didn't seem very important to the evolutionary biologists in the audience, I think because they don't constantly deal with the just-so-story functions that molecular biologists typically ascribe to any complicating feature of a process.  To many molecular biologists, every base pair in the genome, every intermolecular interaction, and every small RNA in the cell is the product of adaptive selection.  There are no accidental interactions.  Shit never just happens.

*On the other hand (not considered by Ford at all), mutations whose effects are made worse by the accidental interaction will be more efficiently eliminated by the stabilizing selection on the function.  I don't think this can be said to reduce the complexity of the function, because the interaction was accidental and thus not included in the complexity count.  I don't know if it would it create selection against the interaction.

**Psci Wavefunction has blogged about this concept in some detail, here and here.  I confess that I haven't read these very long posts through, but perhaps now I will.  (She asked an excellent question after the talk.)

*** Somewhere in his talk Ford was describing clade selection; using the example of how a propensity to speciate can cause a lineage to have many more species than other lineages.  He said that more species means more individuals, but that's certainly not true.