If I do an experiment that uses 50 nm magnetic beads, am I using nanotechnology? If the experiment involves a new use for these beads, am I developing nanotechnology? More generally, is all molecular biology nanotechnology?
Anyway, what am I going to do with these beads? I want to find out whether the USS polarizes the direction of DNA uptake. That is, when the USS sequence interacts with the DNA uptake machinery on the cell surface, does only the DNA on one side of the canonical orientation get pulled into the cell, at least until the end has been brought in?
My latest model of the mechanism of uptake initiation predicts that at the initiation step a type 4 pseudopilus pulls in the DNA on the right side of the USS core, while the core is bound tightly to a receptor protein on the surface (this could be the secretin pore-forming protein). If the core remains tightly bound, uptake could continue only on this side until the end of the fragment is reached. This would require that the force exerted by the pseudopilus on the DNA be less that that needed to pull the USS core free from its receptor. Bringing in the end of the DNA would allow transport across the cytoplasmic membrane, which would reverse the direction of uptake. Under this version of the model, this reversal of direction and the stronger pull exerted on the DNA by the cytoplasmic membrane machinery would detach the USS core from the receptor and bring in the DNA on the left side.
This model of uptake is much more detailed than the available evidence supports. I see this as a strength, not a weakness. The model makes many very specific and testable predictions, and at this stage of investigation it's more important for a hypothesis to be testable than to be correct. Maybe that's true at any stage.
I still like the idea of using laser tweezers to investigate the polarity question, but my physicist collaborator points out that this single-molecule method may lack the resolution we need to answer the question. Tweezers are best at measuring forces, not movement of cells.
So I'm back to thinking about blocking the ends with beads. Our previous attempts used what we now realize were giant beads (streptavidin-agarose), far bigger than the cells and much too big and porous for the task. This improved approach will use beads that are only 50 nm across, which should be plenty big enough to prevent uptake as the secretin pore is thought to be only 6-7 nm across at its widest. They can be purchased with streptavidin bound to their surface, making it easy to attach them to DNA just as we did with the agarose beads.
Our original experiments labeled one end of a short USS-containing fragment with 32P and blocked the other end with a (giant) bead. We predicted that the orientation of the the USS with respect to the bead would determine whether the 32P got inside the cell, but found that orientation had little effect. I think we should repeat these experiments using the 50 nm magnetic beads, paying careful attention to the kinetics. The original experiments used 220 bp fragments; because such short fragments might behave differently than long ones, we should also repeat the analysis with fragments that are one or a few kb long (easy - we just use the whole USS plasmid rather than its 220bp insert).
Maybe we can then combine use of the beads with the tweezer analysis, examining forces rather than distance changes as originally planned. If the tweezer analysis of the force acting on the DNA shows that the inner membrane machinery exerts a stronger force than the pseudopilus, then blocking the right end of a long fragment with a bead (left end held in the laser trap) should allow weak-force uptake but prevent the transition to strong-force uptake. Flipping the orientation of the USS should. . . . . no, the transition would still be blocked, this time by the large trapped polystyrene bead at the other end. The flipped USS orientation would affect movement of the cell rather than the kind of force.
Field of Science
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Change of address1 year ago in Variety of Life
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Change of address1 year ago in Catalogue of Organisms
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Earth Day: Pogo and our responsibility1 year ago in Doc Madhattan
How bacterial conjugation works
I just had a lively discussion with the guy in the next office about whether the single-stranded DNA that enters a bacterial cell by conjugation becomes double-stranded before it recombines into the chromosome. The textbook he's using to teach his new bacterial molecular genetics course say yes, but this seems rather improbable to me.
Another faculty member found a paper from about 10 years ago that concludes that the DNA becomes double-stranded before it recombines. But I don't think this double-strandedness was necessitated by the findings of their experiments; rather it seems to be mainly driven by the assumptions of the researchers.
The paper had another surprise. It showed the DNA being transferred into the recipient cell as a loop, with the 'leading' end being held back in the 'donor' cell, and the loop being pushed into the recipient. I've only seen it represented differently, with the leading end moving into the recipient cell as if it was pulling the DNA strand along behind it. But now I think more carefully, the protein attached to the leading end isn't expected to have any power to 'pull' anything once it's in the recipient cell, so the DNA must be being transported just by the machinery in the membrane of the 'donor'.
I'm putting 'donor' in quotes, because that's how most people think of the process of conjugation. They've been taught to think of the cell that has a plasmid kindly donating a copy of it to a cell that lacks the plasmid; if the plasmid is attached to chromosomal DNA the recipient cell is blessed with a copy of part of the donor's chromosome. But plasmids are genetic parasites, and the 'donation' is really an infection. The recipient is passive, and the plasmid DNA is forcibly inserted into it by the 'donor'. To do this the plasmid uses the proteins it codes for, and proteins already produced by its cell.
Another faculty member found a paper from about 10 years ago that concludes that the DNA becomes double-stranded before it recombines. But I don't think this double-strandedness was necessitated by the findings of their experiments; rather it seems to be mainly driven by the assumptions of the researchers.
The paper had another surprise. It showed the DNA being transferred into the recipient cell as a loop, with the 'leading' end being held back in the 'donor' cell, and the loop being pushed into the recipient. I've only seen it represented differently, with the leading end moving into the recipient cell as if it was pulling the DNA strand along behind it. But now I think more carefully, the protein attached to the leading end isn't expected to have any power to 'pull' anything once it's in the recipient cell, so the DNA must be being transported just by the machinery in the membrane of the 'donor'.
I'm putting 'donor' in quotes, because that's how most people think of the process of conjugation. They've been taught to think of the cell that has a plasmid kindly donating a copy of it to a cell that lacks the plasmid; if the plasmid is attached to chromosomal DNA the recipient cell is blessed with a copy of part of the donor's chromosome. But plasmids are genetic parasites, and the 'donation' is really an infection. The recipient is passive, and the plasmid DNA is forcibly inserted into it by the 'donor'. To do this the plasmid uses the proteins it codes for, and proteins already produced by its cell.
Why is H. influenzae's CRP so feeble?
One discovery from the grad student's work on Sxy and CRP is that the H. influenzae CRP protein binds CRP sites much less strongly (with much lower affinity) than the E. coli CRP protein does. This is a bit surprising. The two proteins have quite similar sequences, and all the amino acid residues expected to directly contact the DNA are identical.
He's going to contact a lab that has done extensive structural analysis of E. coli CRP, to see how difficult it would be to see how well H. influenzae CRP will superimpose on the E. coli structure. One possibility he suggested is that the dimerization domain of H. influenzae CRP could be weak. This would cause the protein to spend less time assembled into the dimers that most readily bind DNA.
Until now we (at least I) had thought that the affinity of CRP for different genes was determined by how well the gene's CRP site matched the protein's requirements for binding. This would have been optimized for each gene by natural selection acting on mutations in its CRP site. But now I'm wondering whether natural selection has also acted differently on E. coli and H. influenzae CRP proteins to fine tune their affinity for all the sites in their respective genomes.
H. influenzae has only about 40% as many genes as E. coli, and about 40% as many CRP sites regulating them. But I can't think of any way that would favour a 100-fold difference in CRP affinity for the same CRP site, which is what the grad student has found.
The assays were done under exactly the same conditions, but this doesn't ensure that the proteins responded identically to these conditions. I wonder if the binding conditions used for these measurements (optimized for E. coli CRP) might be unsuitable for H. influenzae CRP.
He's going to contact a lab that has done extensive structural analysis of E. coli CRP, to see how difficult it would be to see how well H. influenzae CRP will superimpose on the E. coli structure. One possibility he suggested is that the dimerization domain of H. influenzae CRP could be weak. This would cause the protein to spend less time assembled into the dimers that most readily bind DNA.
Until now we (at least I) had thought that the affinity of CRP for different genes was determined by how well the gene's CRP site matched the protein's requirements for binding. This would have been optimized for each gene by natural selection acting on mutations in its CRP site. But now I'm wondering whether natural selection has also acted differently on E. coli and H. influenzae CRP proteins to fine tune their affinity for all the sites in their respective genomes.
H. influenzae has only about 40% as many genes as E. coli, and about 40% as many CRP sites regulating them. But I can't think of any way that would favour a 100-fold difference in CRP affinity for the same CRP site, which is what the grad student has found.
The assays were done under exactly the same conditions, but this doesn't ensure that the proteins responded identically to these conditions. I wonder if the binding conditions used for these measurements (optimized for E. coli CRP) might be unsuitable for H. influenzae CRP.
What part of "match" don't we understand?
The overall goal of the work the post-doc and I describe in our Defining the USS manuscript is to find out whether the sequence specificity of the H. influenzae DNA uptake machinery matches the consensus of the uptake signal sequence (USS) repeats in its genome. We've done good jobs of characterizing both the uptake specificity and the USS consensus, but now we're discovering that comparing them is trickier than we had realized.
The figure shows both results, with the genome consensus shown as a SequenceLogo at the bottom, and the effect on uptake of changing each position shown by the bar chart above. Because all the positions in the core (the AAGTGCGGT at the left) have a very strong consensus in the genome, we expected that changing any of these positions would dramatically decrease uptake. But instead we see that changes to different core positions have very different effects. Changing the first position doesn't decrease uptake at all (within our limit of detection) but changing any of the central three knocks it way down.
One confounding factor comes from how the genome consensus is described; the Y-axis of the logo is not a linear scale but a logarithmic scale reflecting 'information content' of the consensus. Another may come from how the uptake effects were measured; cells were deliberately given less DNA than needed to saturate the uptake machinery.
But the biggest complication is that, although we have a simple 'molecular drive' hypothesis describing how biased DNA uptake leads to accumulation of the preferred sequence in the genome, we are only beginning to develop ways to evaluate the different components of this model. This means that we can't predict exactly what sequences will accumulate in response to any specific uptake bias.
The post-doc describes the discrepancy between the two parts of the figure as maybe resulting from 'saturation' of the evolutionary process causing USS accumulation. If a small uptake bias acting over millions of generations is enough to drive the preferred base at a particular position to a very high frequency, then a larger bias may not make much difference to the outcome. For example, say USSs with a C or G or T at position 1 are taken up 95% as well as a USS with the consensus A at that position, but this bias provides sufficient long-term drive to cause 98% of the USS population to have As at that position. If so, increasing the bias might not increase the frequency of As by very much.
So what will we say in our manuscript? We could start by discussing the implications for the role of the USS in DNA uptake. We'll point out that uptake is very sensitive to the central positions of the core. Even if the other positions in the USS (core and flanking AT-rich segments) are all perfectly matched to the genome consensus, changing any one of these central bases drastically reduces uptake. This suggests that these positions make very important contacts with the uptake machinery. Changes at any of five other core positions reduce uptake to 20-60% of the control (perfect USS), so these also probably make important contacts. Changing the initial A, or pairs of bases in the flanking segments, reduces uptake only modestly, suggesting that the functions of these in uptake are dispensable if the rest of the USS is perfect.
We could go on to discuss the implications for accumulation of USSs in the genome, as I do above, and end by pointing out that what's needed is a better understanding of the consequences of molecular drive (and maybe put in a plug for our Perl simulation model).
The figure shows both results, with the genome consensus shown as a SequenceLogo at the bottom, and the effect on uptake of changing each position shown by the bar chart above. Because all the positions in the core (the AAGTGCGGT at the left) have a very strong consensus in the genome, we expected that changing any of these positions would dramatically decrease uptake. But instead we see that changes to different core positions have very different effects. Changing the first position doesn't decrease uptake at all (within our limit of detection) but changing any of the central three knocks it way down.One confounding factor comes from how the genome consensus is described; the Y-axis of the logo is not a linear scale but a logarithmic scale reflecting 'information content' of the consensus. Another may come from how the uptake effects were measured; cells were deliberately given less DNA than needed to saturate the uptake machinery.
But the biggest complication is that, although we have a simple 'molecular drive' hypothesis describing how biased DNA uptake leads to accumulation of the preferred sequence in the genome, we are only beginning to develop ways to evaluate the different components of this model. This means that we can't predict exactly what sequences will accumulate in response to any specific uptake bias.
The post-doc describes the discrepancy between the two parts of the figure as maybe resulting from 'saturation' of the evolutionary process causing USS accumulation. If a small uptake bias acting over millions of generations is enough to drive the preferred base at a particular position to a very high frequency, then a larger bias may not make much difference to the outcome. For example, say USSs with a C or G or T at position 1 are taken up 95% as well as a USS with the consensus A at that position, but this bias provides sufficient long-term drive to cause 98% of the USS population to have As at that position. If so, increasing the bias might not increase the frequency of As by very much.
So what will we say in our manuscript? We could start by discussing the implications for the role of the USS in DNA uptake. We'll point out that uptake is very sensitive to the central positions of the core. Even if the other positions in the USS (core and flanking AT-rich segments) are all perfectly matched to the genome consensus, changing any one of these central bases drastically reduces uptake. This suggests that these positions make very important contacts with the uptake machinery. Changes at any of five other core positions reduce uptake to 20-60% of the control (perfect USS), so these also probably make important contacts. Changing the initial A, or pairs of bases in the flanking segments, reduces uptake only modestly, suggesting that the functions of these in uptake are dispensable if the rest of the USS is perfect.
We could go on to discuss the implications for accumulation of USSs in the genome, as I do above, and end by pointing out that what's needed is a better understanding of the consequences of molecular drive (and maybe put in a plug for our Perl simulation model).
The post-doc and I got about half-way through our Defining the USS paper today. We fixed up the Methods section and almost all of the Introduction, and walked ourselves through the Results. Maybe because we hadn't looked at them for a couple of weeks, we had some new ideas.
We had been commenting on how the USSs in coding sequences show significantly different USS motifs, depending on which reading frame they're in. But now we look again, we realize that the real story isn't how they are different, but how similar they are. Despite coding for entirely different amino acids, they all have strong matches to the canonical USS core and flanking segments. The differences are so small that they might even be attributable to random effects due to small sample sizes.
This might mean that needing to code for proteins doesn't significantly constrain USS sequences. Rather, USSs accumulate only in places where the existing protein coding constraints meet their need for uptake efficiency.
We also clarified our analysis of the correlations between the bases at different positions in the USS, especially for USSs that are likely to be acting as terminators. Because terminators act by folding into hairpins, we expected to see that the parts of these USSs that come together when folded would show correlations. But they don't. Instead the strong correlations are all between bases that are adjacent in the sequence or separated by only a single base, just as they are in the non-terminator data set. We think this means that the correlations arise from the need for adjacent base pairs to physically cooperate when the USS is kinked during uptake, and that selection for terminator function is much weaker.
We had been commenting on how the USSs in coding sequences show significantly different USS motifs, depending on which reading frame they're in. But now we look again, we realize that the real story isn't how they are different, but how similar they are. Despite coding for entirely different amino acids, they all have strong matches to the canonical USS core and flanking segments. The differences are so small that they might even be attributable to random effects due to small sample sizes.
This might mean that needing to code for proteins doesn't significantly constrain USS sequences. Rather, USSs accumulate only in places where the existing protein coding constraints meet their need for uptake efficiency.
We also clarified our analysis of the correlations between the bases at different positions in the USS, especially for USSs that are likely to be acting as terminators. Because terminators act by folding into hairpins, we expected to see that the parts of these USSs that come together when folded would show correlations. But they don't. Instead the strong correlations are all between bases that are adjacent in the sequence or separated by only a single base, just as they are in the non-terminator data set. We think this means that the correlations arise from the need for adjacent base pairs to physically cooperate when the USS is kinked during uptake, and that selection for terminator function is much weaker.
Defining the uptake signal sequence
The big goal for today is to make progress on the manuscript where we pin down the specifics of the Haemophilus influenzae 'uptake signal sequence' (USS). This DNA sequence can be defined in two ways, both as the sequence motif preferentially taken up by competent H. influenzae cells, and as a sequence motif that's strikingly overabundant in the H. influenzae genome. The two are of course related; we think that the abundance on the genome is a direct consequence of preferential uptake and random (unbiased) recombination between incoming DNA and the chromosome.
The most glaring hole in the manuscript right now is the Methods section where I need to describe the bioinformatics methods I've used to analyze the USS motif in the genome. The post-doc responsible for the DNA uptake experiments has already written her part of the Methods, and today we're going to sit down together and write my part, as well as work on polishing the Results (I think this is in OK shape) and the Discussion (needs work!).
The most glaring hole in the manuscript right now is the Methods section where I need to describe the bioinformatics methods I've used to analyze the USS motif in the genome. The post-doc responsible for the DNA uptake experiments has already written her part of the Methods, and today we're going to sit down together and write my part, as well as work on polishing the Results (I think this is in OK shape) and the Discussion (needs work!).
Polishing the Sxy manuscript
We're fixing the final details on the Sxy manuscript; I'm hoping to have it submitted in the next 24 hours. Below I'll try to summarize what it says, in less technical terms than the Abstract uses.
We already know that the Sxy protein regulates expression of competence genes; here we're examining how Sxy itself is regulated. Our most powerful tools are regulatory mutations that turn Sxy on when it would otherwise be off. The paper starts by describing new mutations that, taken all together, strongly suggest that expression is controlled by changes in RNA folding. We conclude this because all of the mutations change how RNA can fold, but only one of them changes the Sxy protein sequence (and that in a trivial way).
The paper then presents data showing that the RNA folding changes don't affect how much Sxy RNA is made, but how efficiently the RNA is used to make Sxy protein. Then more data shows that normal cells translate their Sxy RNA more efficiently when they're starved, but the mutant cells translate it efficiently even when they're not starved.
The cell uses the genes/proteins that Sxy regulates to take up DNA, so these results help us understand how being starved causes cells to take up DNA. We have argued that cells take up DNA as food (not to get different versions of their genes as others have assumed), so these results strengthen the evidence that DNA uptake is an adaptive response to starvation.
We already know that the Sxy protein regulates expression of competence genes; here we're examining how Sxy itself is regulated. Our most powerful tools are regulatory mutations that turn Sxy on when it would otherwise be off. The paper starts by describing new mutations that, taken all together, strongly suggest that expression is controlled by changes in RNA folding. We conclude this because all of the mutations change how RNA can fold, but only one of them changes the Sxy protein sequence (and that in a trivial way).
The paper then presents data showing that the RNA folding changes don't affect how much Sxy RNA is made, but how efficiently the RNA is used to make Sxy protein. Then more data shows that normal cells translate their Sxy RNA more efficiently when they're starved, but the mutant cells translate it efficiently even when they're not starved.
The cell uses the genes/proteins that Sxy regulates to take up DNA, so these results help us understand how being starved causes cells to take up DNA. We have argued that cells take up DNA as food (not to get different versions of their genes as others have assumed), so these results strengthen the evidence that DNA uptake is an adaptive response to starvation.
What's UP, Sxy?
One of the grad students has been doing experiments to clarify how the transcriptional activator protein Sxy turns on genes that have its recognition sequence, the "CRP-S" site. He's given me a draft of a paper he's writing about this work.
Sxy works by interacting with another transcriptional activator protein called CRP. CRP's job is to bind to DNA at CRP sites and, by bending the DNA, help RNA polymerase start making RNA. CRP-S sites are much harder to bend than normal "CRP-N" sites, and we have been thinking that Sxy acts by helping CRP to bind (if it can't bend the DNA it lets go).
But several pieces of his data tell us that binding isn't enough. A protein from E. coli (we work mostly in another bacterium, Haemophilus influenzae) binds CRP-S sites fairly strongly in the test tube, but when it's in H. influenzae cells it can't turn the CRP-S genes on without Sxy's help. And changing the CRP-S sequence so H. influenzae's CRP can bind it without Sxy in the test tube doesn't enable CRP to turn the gene on in cells without Sxy's help.
So the grad student is suggesting that Sxy may also affect how RNA polymerase interacts with the DNA. RNA polymerase binds to DNA more effectively if its 'tail' makes contact with an AT-rich sequence called the "UP" sequence. Many genes have an UP sequence, and at normal CRP-N sites CRP is thought to help RNA polymerase's tail bind to it.
He's found that the genes with CRP-S promoters have what look like three precisely-spaced UP sequences beside their CRP-S sites, and in his draft paper he proposes that Sxy acts partly by helping RNA polymerase make contact with these sites. We could test this by changing these sequences to not fit the UP consensus, and seeing if this makes the promoter unable to be activated by Sxy.
This work doesn't address the question of whether Sxy does help CRP bend the DNA. But we won't be able to address this until we have conditions where purified CRP and Sxy both interact. Right now we have the CRP but not the Sxy - it's a very difficult protein to work with.
Sxy works by interacting with another transcriptional activator protein called CRP. CRP's job is to bind to DNA at CRP sites and, by bending the DNA, help RNA polymerase start making RNA. CRP-S sites are much harder to bend than normal "CRP-N" sites, and we have been thinking that Sxy acts by helping CRP to bind (if it can't bend the DNA it lets go).
But several pieces of his data tell us that binding isn't enough. A protein from E. coli (we work mostly in another bacterium, Haemophilus influenzae) binds CRP-S sites fairly strongly in the test tube, but when it's in H. influenzae cells it can't turn the CRP-S genes on without Sxy's help. And changing the CRP-S sequence so H. influenzae's CRP can bind it without Sxy in the test tube doesn't enable CRP to turn the gene on in cells without Sxy's help.
So the grad student is suggesting that Sxy may also affect how RNA polymerase interacts with the DNA. RNA polymerase binds to DNA more effectively if its 'tail' makes contact with an AT-rich sequence called the "UP" sequence. Many genes have an UP sequence, and at normal CRP-N sites CRP is thought to help RNA polymerase's tail bind to it.
He's found that the genes with CRP-S promoters have what look like three precisely-spaced UP sequences beside their CRP-S sites, and in his draft paper he proposes that Sxy acts partly by helping RNA polymerase make contact with these sites. We could test this by changing these sequences to not fit the UP consensus, and seeing if this makes the promoter unable to be activated by Sxy.
This work doesn't address the question of whether Sxy does help CRP bend the DNA. But we won't be able to address this until we have conditions where purified CRP and Sxy both interact. Right now we have the CRP but not the Sxy - it's a very difficult protein to work with.
Welcome, visitors from the Just Science links
Unlike most of the other science blogs you might check out, this is a real 'research blog'. In it I write about the experiments and analyses we do in my lab, and about what I think they mean. I've been doing this for about 6 months, and I find that writing it helps me think more clearly about our work.
I don't expect casual visitors (or any visitors except those from my lab) to follow the complexities of the science, but it might give them a sense of what more-or-less ordinary* scientists do and how we think.
*No, of course I don't really think I'm any more ordinary than any other scientist (we all think we're special), but I'm a Canadian so I feel obliged to pretend.
I don't expect casual visitors (or any visitors except those from my lab) to follow the complexities of the science, but it might give them a sense of what more-or-less ordinary* scientists do and how we think.
*No, of course I don't really think I'm any more ordinary than any other scientist (we all think we're special), but I'm a Canadian so I feel obliged to pretend.
DNase I: friend and foe
My posting has been a bit infrequent because I've been focused on getting drafts of our two grant proposals ready for pre-submission critiquing by the volunteer reviewers (blessed be their names and their progeny). But that's done, with a lot of help from a grad student and a postdoc. I've committed myself to the Week of Science Blogging, so I'll be posting at least daily for the next week.
We put a fair bit of effort into crafting a tightly focused "Significance" section to end each proposal (after the detailed descriptions of the experiments we propose). It's important to avoid ending with strings of platitudes, so ours is a bit on the edgy side. On the other hand we can't afford to sound too arrogant (this is a Canadian granting agency).
Some science: We had been hoping to be able to use preparations of cell membranes to study how a radioactively-labeled 'test' DNA fragment interacts with the uptake machinery. But we had overlooked the problem of all the cellular DNA that will be released when we break the cells open to get the membranes.
When the cells are first opened the uptake proteins will bind the abundant cellular DNA, and unless we get rid of the DNA they've bound, they will never bind our test DNA. In principle we could get rid of the cellular DNA by adding a lot of DNase I, an enzyme that breaks down DNA. But then we would need to get rid of ALL of the DNase I , as otherwise it will break down our test DNA!
The postdoc (who discovered the problem) is going to do some preliminary testing to see if this analysis can be salvaged, perhaps by repeatedly washing the membranes to get rid of the DNase I.
We put a fair bit of effort into crafting a tightly focused "Significance" section to end each proposal (after the detailed descriptions of the experiments we propose). It's important to avoid ending with strings of platitudes, so ours is a bit on the edgy side. On the other hand we can't afford to sound too arrogant (this is a Canadian granting agency).
Some science: We had been hoping to be able to use preparations of cell membranes to study how a radioactively-labeled 'test' DNA fragment interacts with the uptake machinery. But we had overlooked the problem of all the cellular DNA that will be released when we break the cells open to get the membranes.
When the cells are first opened the uptake proteins will bind the abundant cellular DNA, and unless we get rid of the DNA they've bound, they will never bind our test DNA. In principle we could get rid of the cellular DNA by adding a lot of DNase I, an enzyme that breaks down DNA. But then we would need to get rid of ALL of the DNase I , as otherwise it will break down our test DNA!
The postdoc (who discovered the problem) is going to do some preliminary testing to see if this analysis can be salvaged, perhaps by repeatedly washing the membranes to get rid of the DNase I.
Does local nicking ease uptake?
Posting has become less frequent because proposal writing and teaching are taking my attention away from blogging about research. But yesterday I had a nice idea of another way to probe how USS sequences facilitate uptake.
We're hypothesizing that USSs are sequences that are more easily taken up because they are easily deformed to pass through the narrow secretin pore. If so, a single-stranded nick might take the place of part of the USS, because it should be easy to bend the DNA sharply at the site of a nick.
I remembered old work showing that restriction enzymes, which normally cut both DNA strands at their recognition sites, will cut only one strand in the presence of high concentrations of the DNA-binding dye ethidium bromide. So if we replace part of a USS with a restriction site, we can test whether a nick at that site increases uptake by competent cells. This should be quite easy, as our standard USS for such experiments is cloned into the middle of the restriction sites of plasmid pGEM, giving us lots of built-in sites to work with.
We're hypothesizing that USSs are sequences that are more easily taken up because they are easily deformed to pass through the narrow secretin pore. If so, a single-stranded nick might take the place of part of the USS, because it should be easy to bend the DNA sharply at the site of a nick.
I remembered old work showing that restriction enzymes, which normally cut both DNA strands at their recognition sites, will cut only one strand in the presence of high concentrations of the DNA-binding dye ethidium bromide. So if we replace part of a USS with a restriction site, we can test whether a nick at that site increases uptake by competent cells. This should be quite easy, as our standard USS for such experiments is cloned into the middle of the restriction sites of plasmid pGEM, giving us lots of built-in sites to work with.
A snapshot of uptake specificity?
We've been using DNA uptake assays with fragments having specific differences from the USS consensus to measure how differences at each USS position affect uptake. But these are a lot of work, and I'm considering an alternative approach that would gather all the information in one big experiment.
We'd start with the plasmid that has a perfect USS insert, and subject it to high-efficiency random mutagenesis in the USS segment (say about 40-50bp). This would be done using a mutagenesis kit and a batch of degenerate oligos for this segment. Each degenerate oligo would have a small probability (say 9%) of having a 'wrong' base at each position (3% of each 'wrong' base), so that on average each oligo in the batch would have about 3-4 differences from the consensus. But the distribution would be broad, so a small fraction of the oligos would have one or no changes, and some would have 5 or more.
We'd then use competent cells to select, from the pool of mutagenized plasmid inserts, ones that can be bound or taken up (depending on whether or not we add DNase I to destroy DNA on the outside of the cells).
Then we'd use PCR to amplify the USS inserts of the taken-up sequences, and analyze their genetic diversity. I'm not up on the technology that would be most appropriate - I'll need to ask and search for genome analysis tools. In principle this could be done in two ways - we'd probably want to do both. The first way would be to use some sort of chip or array (?) to determine the proportion of each base at each position in the 40-50bp we've mutagenized. Because this wouldn't tell us anything about the correlations between differences at different positions, we'd also want to sequence some (say 1000?) of the mutagenized segments.
In principle this is just a high-tech version of analysis Sol Goodgal did about 15 years ago.
We'd start with the plasmid that has a perfect USS insert, and subject it to high-efficiency random mutagenesis in the USS segment (say about 40-50bp). This would be done using a mutagenesis kit and a batch of degenerate oligos for this segment. Each degenerate oligo would have a small probability (say 9%) of having a 'wrong' base at each position (3% of each 'wrong' base), so that on average each oligo in the batch would have about 3-4 differences from the consensus. But the distribution would be broad, so a small fraction of the oligos would have one or no changes, and some would have 5 or more.
We'd then use competent cells to select, from the pool of mutagenized plasmid inserts, ones that can be bound or taken up (depending on whether or not we add DNase I to destroy DNA on the outside of the cells).
Then we'd use PCR to amplify the USS inserts of the taken-up sequences, and analyze their genetic diversity. I'm not up on the technology that would be most appropriate - I'll need to ask and search for genome analysis tools. In principle this could be done in two ways - we'd probably want to do both. The first way would be to use some sort of chip or array (?) to determine the proportion of each base at each position in the 40-50bp we've mutagenized. Because this wouldn't tell us anything about the correlations between differences at different positions, we'd also want to sequence some (say 1000?) of the mutagenized segments.
In principle this is just a high-tech version of analysis Sol Goodgal did about 15 years ago.
Sxy and CRP
One of the grad students and I were talking this afternoon, trying to develop a model of how the regulatory proteins Sxy and CRP interact to cause transcription from promoters with CRP-S sites. We wound up with a model where Sxy does two things that increase transcription at these otherwise-unfavourable promoters.
The first thing would be that Sxy helps CRP bind to and bend DNA right at the CRP-S site. At CRP-N sites, which have an easily-bendable sequence, Sxy isn't needed for CRP binding.
The second thing would be that Sxy also interacts with another attribute of CRP-S promoters, and together with CRP helps RNA polymerase to begin transcription. This 'other attribute' is probably outside of the core CRP-S site; it could be a part of the nearby sequence that we haven't examined yet, or something about the separation of the CRP and RNA polymerase binding sites. If CRP binds to such a site without Sxy, it can't initiate transcription.
The other component of our model is that there is much more CRP than Sxy in the cell, so most of the CRP isn't associated with Sxy, but most of the Sxy is associated with CRP.
This model explains why changing the core of a CRP-S site into a CRP-N type core doesn't allow the promoter to work nearly as effectively as an authentic CRP-N site. And why doing the reverse (changing a CRP-N core into a CRP-S core) also creates a lousy promoter.
In our sketches we always had Sxy associate with CRP in solution, before either made contact with the DNA. But I'm wondering if this is necessary, or if one or the other protein might bind DNA and then its partner. Probably it is necessary. E. coli CRP alone binds very poorly to CRP-S sites in vitro; H. influenzae CRP won't bind them at all. So either Sxy must bind the DNA at CRP-S sites before the CRP gets there, or they must meet up in solution. Sxy has none of the features of typical DNA-binding proteins.
Of course, it could be that Sxy doesn't bind CRP at all, but instead binds to RNA polymerase....
The first thing would be that Sxy helps CRP bind to and bend DNA right at the CRP-S site. At CRP-N sites, which have an easily-bendable sequence, Sxy isn't needed for CRP binding.
The second thing would be that Sxy also interacts with another attribute of CRP-S promoters, and together with CRP helps RNA polymerase to begin transcription. This 'other attribute' is probably outside of the core CRP-S site; it could be a part of the nearby sequence that we haven't examined yet, or something about the separation of the CRP and RNA polymerase binding sites. If CRP binds to such a site without Sxy, it can't initiate transcription.
The other component of our model is that there is much more CRP than Sxy in the cell, so most of the CRP isn't associated with Sxy, but most of the Sxy is associated with CRP.
This model explains why changing the core of a CRP-S site into a CRP-N type core doesn't allow the promoter to work nearly as effectively as an authentic CRP-N site. And why doing the reverse (changing a CRP-N core into a CRP-S core) also creates a lousy promoter.
In our sketches we always had Sxy associate with CRP in solution, before either made contact with the DNA. But I'm wondering if this is necessary, or if one or the other protein might bind DNA and then its partner. Probably it is necessary. E. coli CRP alone binds very poorly to CRP-S sites in vitro; H. influenzae CRP won't bind them at all. So either Sxy must bind the DNA at CRP-S sites before the CRP gets there, or they must meet up in solution. Sxy has none of the features of typical DNA-binding proteins.
Of course, it could be that Sxy doesn't bind CRP at all, but instead binds to RNA polymerase....
New analysis of sxy RNase data
I just met with the grad students to discuss their re-analysis of how RNAses sensitive to base pairing cut sxy mRNA. The new analysis is much better than what we had before. Rather than subjectively scoring sites as 'strongly cut', 'weakly cut' or 'uncut', we now have quantitative measures of the intensity of cutting at each site, and of the effect of the two mutations on this cutting.
The tricky part is how we combine the RNase data with our genetic evidence and with the secondary structure predicted by Mfold. The Mfold predictions look good, and it's easy to give more credibility to a hypothesis expressed as a drawing of a structure than to one expressed only in words or numbers (our brains love pictures). But I would feel more comfortable with the predictions if we were able to use the software at a more sophisticated level, rather than just pasting in various sequences and leaving all the settings at their defaults.
The tricky part is how we combine the RNase data with our genetic evidence and with the secondary structure predicted by Mfold. The Mfold predictions look good, and it's easy to give more credibility to a hypothesis expressed as a drawing of a structure than to one expressed only in words or numbers (our brains love pictures). But I would feel more comfortable with the predictions if we were able to use the software at a more sophisticated level, rather than just pasting in various sequences and leaving all the settings at their defaults.
Hypotheses about loss of competence
The post-doc working on how and why cells lose competence has some very interesting data, but it makes us realize that we haven't yet spelled out clear hypotheses for this data to address. It's tempting to try to come up with the true explanation of why many cells/strains/species lose competence while others retain it, but that's not what we need at this stage. Instead we just need some clear overly-simplistic statements of the different possible causes and their consequences, to guide our thinking.
So here's one possibility:
Maybe cells take up DNA only because the genetic changes this sometimes causes are occasionally beneficial. (Most people think this is true, though I think the DNA=food consequence is much more important.) These benefits will be rare. So most competent cells will go for long periods taking up DNA but getting no benefit. When mutations that reduce or eliminate gene function arise in genes needed only for DNA uptake, there may be no selection against them for very long times. Depending on the particular mutations, these cells will have an advantage because they won't waste resources taking up DNA that's doing them no immediate good. So the frequency of competent cells in the population will be gradually decreasing.
(I write 'may be' because mutations that mess up one component of a complex machine may cause harm in ways that eliminating the whole machine wouldn't (like the difference between a car with no brakes and no car at all). For example, knocking out the secretin pore but keeping the rest of the DNA uptake machinery messes up the membranes of competent cells in ways that knocking out the ability to turn on competence doesn't. But lets not worry about this right now.)
But once in a while a cell that takes up DNA gets a good genetic change, one that lets it outcompete its relatives. This cell and its immediate descendants all have fully functional competence genes, so the frequency of competent cells in the population has increased.
The long-term outcome depends on how often DNA uptake produces good changes and how often deleterious mutations arise in competence genes and how harmful or beneficial these mutations are in the short term. If the good changes happen often enough, this could give populations that always contained lots of competent cells and some recently arisen non-competent ones. But if the good changes are less frequent, the cells with mutations causing loss of competence could completely take over. And once this happens there's no going back.
So here's one possibility:
Maybe cells take up DNA only because the genetic changes this sometimes causes are occasionally beneficial. (Most people think this is true, though I think the DNA=food consequence is much more important.) These benefits will be rare. So most competent cells will go for long periods taking up DNA but getting no benefit. When mutations that reduce or eliminate gene function arise in genes needed only for DNA uptake, there may be no selection against them for very long times. Depending on the particular mutations, these cells will have an advantage because they won't waste resources taking up DNA that's doing them no immediate good. So the frequency of competent cells in the population will be gradually decreasing.
(I write 'may be' because mutations that mess up one component of a complex machine may cause harm in ways that eliminating the whole machine wouldn't (like the difference between a car with no brakes and no car at all). For example, knocking out the secretin pore but keeping the rest of the DNA uptake machinery messes up the membranes of competent cells in ways that knocking out the ability to turn on competence doesn't. But lets not worry about this right now.)
But once in a while a cell that takes up DNA gets a good genetic change, one that lets it outcompete its relatives. This cell and its immediate descendants all have fully functional competence genes, so the frequency of competent cells in the population has increased.
The long-term outcome depends on how often DNA uptake produces good changes and how often deleterious mutations arise in competence genes and how harmful or beneficial these mutations are in the short term. If the good changes happen often enough, this could give populations that always contained lots of competent cells and some recently arisen non-competent ones. But if the good changes are less frequent, the cells with mutations causing loss of competence could completely take over. And once this happens there's no going back.
Mass spectrometry for the masses (=us)?
Today I consulted two very helpful experts in mass spectrometric to find out the sequences of al;l the peptides in the mix (more or less) methods. I wanted to find out if we really have a hope of identifying DNA uptake proteins by crosslinking them to DNA. It looks promising.
The plan is to incubate competent cells with DNA, and then add formaldehyde, which will create crosslinks between DNA and the proteins it's in contact with. We'll then dissolve the cells and pull out the DNA with its attached proteins. Then we'll get rid of the DNA and undo the crosslinks (by boiling the mix), leaving us with a little tube containing a mixture of DNA-contacting proteins of unknown identities. Then we'll digest the proteins with the protease trypsin, which will cut them into predictable pieces.
We'll use a combination of HPLC (high performance liquid chomatography) and mass spectrometry to find out the amino acid sequences of all the peptides in the mix. By comparing these with the known sequences of all the proteins specified by the genome, we'll know what proteins the peptides came from. By comparing results with DNAs that either do or don't have a USS (or have a variant USS) and with cells carrying different mutations, we can infer a lot about the specific interactions (I hope).
We don't need to invest in equipment for or learn how to do the HPLC and mass spec; we can pay local experts to do it for us.
One issue we'll need to grapple with is the small amounts of protein our 'fishing' technique is likely to produce. I think we can scale up, but that's always a source of problems. Another issue is stopping the cells from quickly sucking the DNA all the way inside - we may be able to control this by initially using cells with uptake mutations, or by sticking the DNA onto beads that are too big for the cells to take up.
The plan is to incubate competent cells with DNA, and then add formaldehyde, which will create crosslinks between DNA and the proteins it's in contact with. We'll then dissolve the cells and pull out the DNA with its attached proteins. Then we'll get rid of the DNA and undo the crosslinks (by boiling the mix), leaving us with a little tube containing a mixture of DNA-contacting proteins of unknown identities. Then we'll digest the proteins with the protease trypsin, which will cut them into predictable pieces.
We'll use a combination of HPLC (high performance liquid chomatography) and mass spectrometry to find out the amino acid sequences of all the peptides in the mix. By comparing these with the known sequences of all the proteins specified by the genome, we'll know what proteins the peptides came from. By comparing results with DNAs that either do or don't have a USS (or have a variant USS) and with cells carrying different mutations, we can infer a lot about the specific interactions (I hope).
We don't need to invest in equipment for or learn how to do the HPLC and mass spec; we can pay local experts to do it for us.
One issue we'll need to grapple with is the small amounts of protein our 'fishing' technique is likely to produce. I think we can scale up, but that's always a source of problems. Another issue is stopping the cells from quickly sucking the DNA all the way inside - we may be able to control this by initially using cells with uptake mutations, or by sticking the DNA onto beads that are too big for the cells to take up.
Grantspersonship
Today I spoke with a colleague with extensive experience of the granting agency I'm applying to (CIHR, the Canadian Institutes for Health Research). I was wondering whether I should split my grant proposal into two smaller and more tightly focused proposals, to be sent to two different peer-review committees.
Right now my proposal addresses two distinct aspects of H. influenzae competence. One is the regulation of competence, specifically the signals that induce the master regulator Sxy and the process by which Sxy induces the competence regulon genes. The other is the mechanism of DNA uptake, especially the role of the uptake signal sequence.
I could easily split this into two smaller proposals, each asking for less money, and each targeted to a particular peer-review committee. Because each proposal would be more focused, there would be more room in the allowed 11 pages to explain the issues specifically important for it. This would make each proposal easier for the peer-reviewers to read and understand. If only one of them got funded, we'd have plenty of money to go on with and I'd resubmit the other one for the next competition six months later.
I'm going to wait till I've gotten the opinions of a couple of other colleagues before making a final decision.
Got your pencils and rubber bands?
I've been discussing the DNA uptake issues by email with a colleague who studies type four pili. She's pushed me into checking some things I thought I knew, and into making my ideas about uptake clearer.
First the retraction issue: I had been thinking that we knew that H. influenzae and other Pasteurellacea could do 'twitching motility', which meant that they must be able to actively retract their pili. But I rechecked and found that nobody has directly shown that they move. Strains of H. influenzae that have pili do form the same 'rafts' at the edges of their colonies that cells with known twitching motility do, but nobody's watched them doing it. I don't know how difficult this would be, but in any case we can't do it until we have cells that we know are piliated.
Second, the DNA-kinking issue: I had been thinking only at an intuitive level about this (hand-waving and doodles), and her questions pushed me into coming up with a physical model.
We can reasonably the DNA and pilus with a long rubber band (= circular DNA) and a pencil (= the pilus); the relative diameters are appropriate. If a loop of DNA is going to be pulled through a pore that's only slightly wider than the pencil, then all of the loop has to become closely appressed to the pencil. For this to happen the DNA must, at some point, make a 180 degree turn in a distance no longer than the diameter of the pencil. If we were to let the DNA detach from the pencil the turn could open up to a larger diameter, but then the pencil plus DNA wouldn't fit through the pore.
The 180 degree turn is a problem because the persistence length of DNA is about 50nm (150bp), whereas the pilus diameter is only about 6nm. Nucleosomes manage to bend DNA smoothly through two full turns over distances of 165bp; this is a diameter of about 9nm (radius of curvature of 4.3nm). Perhaps not coincidentally, part of the uptake signal sequence resembles a nucleosome-positioning sequence.
So the DNA loop needs to bend on the pilus a bit more sharply than it does in a nucleosome - maybe not too big a problem. And if the DNA is lying in the positively charged grooves of the pilus, it may not stick out too far to fit through the pore (our model needs a pencil with grooves). But where the DNA makes its turn, it will have to leave its snug grooves and cross over at least one of the raised parts of the pilus, making an awkward bulge on the surface that's going to have a hard time fitting through the pore.
So uptake of a loop raises two problems. One is forcing the DNA to bend sharply, and the other is fitting the pilus back through the secretin pore once it has a loop of DNA attached to it. I don't think this is impossible, but it probably takes some specialized interactions. I think the role of the uptake signal sequence is to interact with the pilus and secretin to facilitate this.
I'm going to see if I can buy some tubing of different sizes to use for a demo of the problem.
First the retraction issue: I had been thinking that we knew that H. influenzae and other Pasteurellacea could do 'twitching motility', which meant that they must be able to actively retract their pili. But I rechecked and found that nobody has directly shown that they move. Strains of H. influenzae that have pili do form the same 'rafts' at the edges of their colonies that cells with known twitching motility do, but nobody's watched them doing it. I don't know how difficult this would be, but in any case we can't do it until we have cells that we know are piliated.
Second, the DNA-kinking issue: I had been thinking only at an intuitive level about this (hand-waving and doodles), and her questions pushed me into coming up with a physical model.
We can reasonably the DNA and pilus with a long rubber band (= circular DNA) and a pencil (= the pilus); the relative diameters are appropriate. If a loop of DNA is going to be pulled through a pore that's only slightly wider than the pencil, then all of the loop has to become closely appressed to the pencil. For this to happen the DNA must, at some point, make a 180 degree turn in a distance no longer than the diameter of the pencil. If we were to let the DNA detach from the pencil the turn could open up to a larger diameter, but then the pencil plus DNA wouldn't fit through the pore.
The 180 degree turn is a problem because the persistence length of DNA is about 50nm (150bp), whereas the pilus diameter is only about 6nm. Nucleosomes manage to bend DNA smoothly through two full turns over distances of 165bp; this is a diameter of about 9nm (radius of curvature of 4.3nm). Perhaps not coincidentally, part of the uptake signal sequence resembles a nucleosome-positioning sequence.
So the DNA loop needs to bend on the pilus a bit more sharply than it does in a nucleosome - maybe not too big a problem. And if the DNA is lying in the positively charged grooves of the pilus, it may not stick out too far to fit through the pore (our model needs a pencil with grooves). But where the DNA makes its turn, it will have to leave its snug grooves and cross over at least one of the raised parts of the pilus, making an awkward bulge on the surface that's going to have a hard time fitting through the pore.
So uptake of a loop raises two problems. One is forcing the DNA to bend sharply, and the other is fitting the pilus back through the secretin pore once it has a loop of DNA attached to it. I don't think this is impossible, but it probably takes some specialized interactions. I think the role of the uptake signal sequence is to interact with the pilus and secretin to facilitate this.
I'm going to see if I can buy some tubing of different sizes to use for a demo of the problem.
Does uptake depend on later steps?
I've been wondering how interdependent the different parts of the uptake machinery are.
We have long known that some H. influenzae mutants (knockouts of rec2 or comF) could take up DNA but not translocate it into the cytoplasm. They take up about the same amounts of DNA as do wildtype cells, but the DNA stays intact (double-stranded, not degraded). Because this DNA doesn't get cut by the nucleases we know to be active in the cytoplasm, the DNA must be accumulating in the periplasm. This means that the uptake machinery can continue to operate in the absence of translocation.
What about ComE1? This H. influenzae protein is a homolog of the well-characterized B. subtilis protein ComEA. In B. subtilis, ComEA is thought to sit on the periplasmic side of the membrane, under the thick cell wall (there is no outer membrane). The pseudopilus machinery (specified by the ComG proteins) passes DNA across the cell wall to ComEA, and ComEA passes the DNA to the membrane transport machinery (homologs of Rec2 and ComF).
ComEA is essential for DNA uptake in B. subtilis, and its homologs are essential in other competent bacteria. But the H. influenzae homolog, ComE1,is not essential - knockouts reduce transformation by only about 10-fold, and reduce uptake by only about 7-fold. Nevertheless, let's assume that ComE1 does more or less the same thing that ComEA does - accepts DNA from the pseudopilus and passes the DNA on to the machinery that moves it across the inner membrane.
Here's the question: Can the pseudopilus keep doing its job (reeling DNA in) even if ComE1 isn't passing the DNA on? Put another way, do mutations in comE1 reduce DNA uptake, or are they like mutations in rec2 and comF? I already told you the answer - a comE1 knockout reduces DNA uptake as well as transformation. We've known this for several years, but I only now put it into the context of the newer B. subtilis results.
So this means that my model of DNA uptake has to include ComE1 accepting the DNA from the pseudopilus and passing it on to Rec2 and ComF if they're available, or letting the DNA pile up in the periplasm if they're not. If ComE1 isn't there, the pseudopilus stalls. The residual transformation we see in cells lacking ComE1 probably means that about 10% of the DNA finds its way from the pseudopilus to the translocation machinery even in the absence of ComE1, and that this machinery can transport DNA that hasn't been handled by ComE1.
This interpretation makes a prediction about an experiment we've already done - that the reduced DNA uptake seen in comE1 mutants is still USS-dependent. And that's what we see. There's another prediction, that comE1 mutants should not be defective in the initiation of uptake (passing the initial loop through the pore) but only in the subsequent reeling in of the DNA by the pseudopilus. Perhaps we can test this using laser-tweezers, or by cross-linking analysis. From this perspective the comE1 mutant may be very useful in helping us dissect steps of uptake, as it would cause uptake to stall or slow.
This is all very satisfying. We've had the comE1 data for a long time, and tried to write it up, but haven't published it because it didn't seem to explain anything. I think our confusion arose from trying to distinguish between 'binding' and 'uptake' of DNA, whereas now I'm distinguishing between the 'initiation' and 'continuation' of uptake. Now we have a framework for these results, they will make a nice little paper. Unfortunately the technician and M.Sc. student who did the work are long gone so they can't help rewrite the draft, and some of their experiments should be repeated and expanded a bit. Luckily one of the present post-docs is doing similar experiments on other strains, and it would probably be quite simple for her to do the needed experiments and help finish the paper (on which she would then be first author).
We have long known that some H. influenzae mutants (knockouts of rec2 or comF) could take up DNA but not translocate it into the cytoplasm. They take up about the same amounts of DNA as do wildtype cells, but the DNA stays intact (double-stranded, not degraded). Because this DNA doesn't get cut by the nucleases we know to be active in the cytoplasm, the DNA must be accumulating in the periplasm. This means that the uptake machinery can continue to operate in the absence of translocation.
What about ComE1? This H. influenzae protein is a homolog of the well-characterized B. subtilis protein ComEA. In B. subtilis, ComEA is thought to sit on the periplasmic side of the membrane, under the thick cell wall (there is no outer membrane). The pseudopilus machinery (specified by the ComG proteins) passes DNA across the cell wall to ComEA, and ComEA passes the DNA to the membrane transport machinery (homologs of Rec2 and ComF).
ComEA is essential for DNA uptake in B. subtilis, and its homologs are essential in other competent bacteria. But the H. influenzae homolog, ComE1,is not essential - knockouts reduce transformation by only about 10-fold, and reduce uptake by only about 7-fold. Nevertheless, let's assume that ComE1 does more or less the same thing that ComEA does - accepts DNA from the pseudopilus and passes the DNA on to the machinery that moves it across the inner membrane.
Here's the question: Can the pseudopilus keep doing its job (reeling DNA in) even if ComE1 isn't passing the DNA on? Put another way, do mutations in comE1 reduce DNA uptake, or are they like mutations in rec2 and comF? I already told you the answer - a comE1 knockout reduces DNA uptake as well as transformation. We've known this for several years, but I only now put it into the context of the newer B. subtilis results.
So this means that my model of DNA uptake has to include ComE1 accepting the DNA from the pseudopilus and passing it on to Rec2 and ComF if they're available, or letting the DNA pile up in the periplasm if they're not. If ComE1 isn't there, the pseudopilus stalls. The residual transformation we see in cells lacking ComE1 probably means that about 10% of the DNA finds its way from the pseudopilus to the translocation machinery even in the absence of ComE1, and that this machinery can transport DNA that hasn't been handled by ComE1.
This interpretation makes a prediction about an experiment we've already done - that the reduced DNA uptake seen in comE1 mutants is still USS-dependent. And that's what we see. There's another prediction, that comE1 mutants should not be defective in the initiation of uptake (passing the initial loop through the pore) but only in the subsequent reeling in of the DNA by the pseudopilus. Perhaps we can test this using laser-tweezers, or by cross-linking analysis. From this perspective the comE1 mutant may be very useful in helping us dissect steps of uptake, as it would cause uptake to stall or slow.
This is all very satisfying. We've had the comE1 data for a long time, and tried to write it up, but haven't published it because it didn't seem to explain anything. I think our confusion arose from trying to distinguish between 'binding' and 'uptake' of DNA, whereas now I'm distinguishing between the 'initiation' and 'continuation' of uptake. Now we have a framework for these results, they will make a nice little paper. Unfortunately the technician and M.Sc. student who did the work are long gone so they can't help rewrite the draft, and some of their experiments should be repeated and expanded a bit. Luckily one of the present post-docs is doing similar experiments on other strains, and it would probably be quite simple for her to do the needed experiments and help finish the paper (on which she would then be first author).
Sticky tip?
I'm deep into the literature on type four pili. (They're variously abbreviated as Tfp or T4P - I'm going to go with Tfp* because a Google search for Tfp pilin finds over 10,000 hits, whereas a search for T4P pilin finds only 273.)
I was thinking that the USS-specific interaction between pilin subunits and the DNA sequence flanking the core USS would be mediated by the positively charged regions of the pilin that are exposed on the sides of the pilus, but it could instead be a region that is exposed only at the pilus tip.
When Tfp cause adhesion or twitching motility by sticking to surfaces, it's the tip of the pilus that does the sticking. Because of the way the subunits assemble, the top and part of the sides of each non-tip subunit are covered by the subunits above them, but the tops and parts of the sides of the subunits at the tip are exposed, and can interact with their environment.
If the side of the pilus was interacting with the USS outside of the pore, when the pilus retracted the pore would need to accommodate the pilus plus the two sides of the DNA loop. But the pores that have been studied aren't big enough for this; they are about 6-7nm across when open, and can fit a pilus (5-5nm), or two filaments of double-stranded DNA (2nm each), but not them all, and maybe not even the pilus plus one filament of DNA.
Thus continuing uptake may require that the pilus stay in the periplasm, and reel the DNA in through the pore (as I drew it in this post). But when uptake is being initiated the pilus must protrude through the pore to form the initial attachment to the DNA. Pulling the DNA in through the pore would be a lot easier if the DNA was attached to the pilus tip rather than to its side.
I have an initiation figure that shows the DNA binding to the side of the pilus; I'll modify it and add it to this post later.
* Except of course when corresponding with someone who prefers T4P, such as the author of a recent helpful review.
I was thinking that the USS-specific interaction between pilin subunits and the DNA sequence flanking the core USS would be mediated by the positively charged regions of the pilin that are exposed on the sides of the pilus, but it could instead be a region that is exposed only at the pilus tip.
When Tfp cause adhesion or twitching motility by sticking to surfaces, it's the tip of the pilus that does the sticking. Because of the way the subunits assemble, the top and part of the sides of each non-tip subunit are covered by the subunits above them, but the tops and parts of the sides of the subunits at the tip are exposed, and can interact with their environment.
If the side of the pilus was interacting with the USS outside of the pore, when the pilus retracted the pore would need to accommodate the pilus plus the two sides of the DNA loop. But the pores that have been studied aren't big enough for this; they are about 6-7nm across when open, and can fit a pilus (5-5nm), or two filaments of double-stranded DNA (2nm each), but not them all, and maybe not even the pilus plus one filament of DNA.
Thus continuing uptake may require that the pilus stay in the periplasm, and reel the DNA in through the pore (as I drew it in this post). But when uptake is being initiated the pilus must protrude through the pore to form the initial attachment to the DNA. Pulling the DNA in through the pore would be a lot easier if the DNA was attached to the pilus tip rather than to its side.
I have an initiation figure that shows the DNA binding to the side of the pilus; I'll modify it and add it to this post later.
* Except of course when corresponding with someone who prefers T4P, such as the author of a recent helpful review.
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