RRResearch

Sorry for lack of posts...

2012-01-26T17:50:00.000-08:00

We're busy finishing the Science/arseniclife paper, and the postdoc's uptake paper, and the RA's E. coli competence paper (submitted!), and an old visitor's competence paper, and my article about teaching genetics....

Sudsy gel

2012-01-22T17:23:00.001-08:00

Why did I put SDS into the buffer of this agarose gel before I loaded it? So the DNA from the lysed cells wouldn't rise up out of the wells and spread out over the surface of the gel buffer, of course!

I'll tell you more tomorrow, if my experiment works out

The Discussion for the post-doc's DNA uptake paper

2012-01-20T19:14:00.000-08:00

The post-doc and I have been struggling, independently and together, to create a good Discussion section for his paper on the sequence specificity of DNA uptake. We have lots of things we could write about, but many of them aren't well connected to each other or to what the paper is about. But now that I've done some good work on the end of the Results, I think I've finally come up with a Discussion that might work.

The Results ends with the analysis of possible interactions between bases at different positions in the uptake signal sequence motif he derived. We motivate this analysis as a possible explanation of the discrepancy between his uptake sequence motif and the one I derived years ago for the uptake sequences in the H. influenzae genome. I'm reproducing the two motifs below and below them his figure of his interaction analysis.

He's now done an uptake experiment that validates (confirms the predictions of) the interaction analysis. It shows that having mutations at two interacting positions (positions 4 and 11, I think) does indeed reduce DNA uptake much more strongly than predicted by the effect of each mutation singly. This motivated me to clarify for myself the implications of the interaction analysis.

The diagram is at the top of this post. The center four positions of the core (left segment) are greyed out, because their effect on uptake is so strong that we can't make confident inferences about their interactions with other positions. The black brackets above each segment indicates that all of the bracketed positions participate in interactions with all the positions in the other bracketed segments, as indicated by the blue arrows. However the positions within a single bracketed segments do not interact with each other, unlike the minor covariation interactions (figure below) we found long ago between adjacent positions in the genomic USS sequences (pdf of the paper here).

Anyway, back to the Discussion...

First we can explain how the interaction analysis nicely reinforces the hypothesis that the uptake sequences in the genome are there as a direct and unselected molecular-drive consequence of the bias of the uptake machinery. This is an 'exception that proves the rule' situation, where the initial finding that the simple uptake-bias motif didn't match the genomic USS motif created doubt about the hypothesis, and the subsequent demonstration that interactions explain the discrepancy increased our confidence in it.

Then we can say that this leaves only the uptake bias itself in need of an explanation, and that we propose that it exists as part of a solution to the mechanistic problem of getting stiff, highly charged DNA molecules through the narrow secretin pore. Because cells efficiently take up closed circular DNAs we know that uptake doesn't usually initiate at a fragment end, but must initiate internally on DNA fragments (see this very old post). We hypothesize that the uptake bias favours sequences that are readily kinked, and that this kinking occurs mainly as consequence of interactions between the uptake sequence and mutually-interacting proteins of the uptake machinery (the uptake motif is itself only slightly bent, at the T-tracts). One reason to think that proteins mediate the interactions is that adjacent positions don't interact with each other.

Perhaps we can here pose a specific model of what parts of the uptake sequence interact with what parts of the machinery... This should take into account that the T-tracts interact with the core positions but not with each other.

Finally we can discuss the known or possible uptake biases of other species. First the other Pasteurellaceae, then the Neisserias, and finally bacteria where uptake bias may have been overlooked.

Growth of GFAJ-1 under phosphate limitation (correction)

2012-01-20T16:09:00.000-08:00

Erika Check Hayden's otherwise-excellent Nature News report on our work contained one error, the statement that "Redfield was unable to grow any cells without adding a small amount of phosphorus".

Here's the email I had sent her in response to an earlier query about phosphorus concentrations:

Hi Erika,

The amount of phosphate in the medium used by Wolfe-Simon et al for their published growth analysis is indeed uncertain. Their ICP-MS analysis found that most of their media preparations contained 3-4 µM phosphorus, but one batch contained <0.3 µM and a solution containing only the AML60-medium salts had 7.8 µM. Because we don't know which batch was used for the results in their Figure 1, 3-4 µM is a good estimate of the phosphorus contamination, but the actual amount could have been substantially lower or higher.

My cells did grow in medium with no added phosphorus*, to about 5 x 10^6 cells/ml. This is about 1/4 of the density reached by GFAJ-1 in Wolfe-Simon et al's '-P/+As' medium. Adding 3 µM phosphorus to my medium increased GFAJ-1 growth fourfold, to the same density as reported in Wolfe-Simon et al's experiments. Simple algebra thus suggests that my unsupplemented medium contained about 1 µM phosphorus. The correspondence of the cell densities reached in my supplemented (3 µM) and their unsupplemented medium supports the estimate of 3-4 µM contaminating phosphorus in their medium.

My cells, like theirs, were clearly phosphorus-limited, because they grew to much higher densities when additional phosphorus was provided (see my recent RRResearch post and their Fig. 1).

I think this is the best that can be done, since Wolfe-Simon et al. apparently did not keep good enough records to determine the actual phosphorus concentration of the medium they used for their reported experiments.

Hope this helps,

Rosie

*The initial growth problem was not due to a lack of phosphorus but to the need for an amino acid, which I solved by supplementing the medium with a small amount of glutamate.

Not me!

2012-01-17T16:25:00.000-08:00

GFAJ-1 growth curves in limiting phosphate

2012-01-17T11:43:00.000-08:00

The BioScreen is a wonderful time-saver. Over the weekend it did growth curves using media with 9 different concentrations of phosphate, each with 10 replicates, taking readings every 20 minutes for 46 hr!

This data tells me that my choice of 3 µM added phosphate was good; it gives about four times as much growth as no added phosphate, and twice as much as 1 µM, so the unsupplemented medium probably has about 1 µM contaminating phosphate.

The big surprise is that cells reach higher densities with a moderate amount of phosphate (70 µM) than they do with 250 µM or with the 1500 µM used by Wolfe-Simon et al. I don't think this has any serious implications for our analysis.

I was also surprised to see that the cultures with the higher amounts of phosphate were still growing at the end of the time course. I'm going to replicate these results with another time course, and this time I'll run it for longer (3 days? 4 days?).

The CsCl/mass spectrometry data

2012-01-16T19:53:00.000-08:00

Here's the figure the collaborating grad student sent, showing his LC-MS analysis results of two DNA samples from the first set of GFAJ-1 preparations I sent him.

Each data point is a fraction from one of the CsCl gradients he fractionated the two GFAJ-1 DNA samples on (one for the -As/-P DNA and one for the +As/-P DNA). The -P condition is actually 3 µM added phosphate - this gives growth to approximately the same density as Wolfe-Simon et al's '-P' condition.

The lines with the solid symbols show the amount of DNA in each fraction - these each show a nice DNA peak at around the 800 µl position in the gradient.

The lines with open symbols show the amount of arsenate in each of these fractions - these lines are hard to see because they're sitting right on top of the X-axis (yes, that means that the amounts of arsenate detected are ~ zero 'ion counts'). The real values aren't necessarily zero, but they're below the detection limit for this experiment.

The dashed line shows the amount of arsenate that should have been detected if 4% of the phosphate in the DNA had been replaced by arsenate, as predicted by Wolfe-Simon et al's gel analysis (data in their Table S2).

The second graph shows his standard curve for arsenate detection.

Academic publishing gets even sleazier

2012-01-15T08:26:00.000-08:00

An email from Scientific and Academic Publishing:

Dear Rosemary J. Redfield,

This is Scientific & Academic Publishing, USA. Nice to get your information from the journal PLOS Pathogens and also happy to pass on our regards to you from the editorial department of SAP.

We've finished reading the abstract of your paper Transformation of Natural Genetic Variation into Haemophilus Influenzae Genomes and will recommend it to our editors. If you are interested in our journals and want to publish it on our journals, please extend this paper and describe your latest research achievements and send it to us by our online submission system (http://www.manuscriptsystem.com). All manuscripts submitted will be considered for publication.

If this paper has been published, we also welcome you to submit other papers to us.

Welcome to visit our website at http://www.sapub.org.

In the second paragraph they seem to be first saying they'll recommend my already-published paper to their editors (for the editors to do what, read it with admiration?), and then asking me to add a bit of new material to it and submit it to them for publication. This reeks of self-plagiarization. But in the next sentence they ask for other papers instead.

Who are these guys? Their web site lists an impressive 133 journal titles. But most of the ones I clicked on are nonexistent - they have some Editorial Board members but no Editor in Chief or ISBN number, and haven't published any papers. Only one (The International Journal of Plant Research) had 'published' any papers, and these each had only Abstract and reference list- the body of the paper was apparently 'coming soon'. Perhaps this is to be expected, given that this journal too lacks an Editor in Chief. It may lack editors entirely - authors are instructed that they must format the html links for the references they cite, a function normally done by a journal's copy editors.

Their office is in California, so they're not a third-world effort. I couldn't find any information about publication charges at all, but I don't suppose they're just doing this for the glory.

Hmm, the International Journal of Genetic Engineering needs an Editor in Chief - that would look good on my CV. All I need to do is check the boxes on the handy application form they provide!

Here's the gel photo

2012-01-14T17:43:00.000-08:00

These DNAs were all stored in the fridge (4 °C) in aqueous solution (10 mM Tris 1 mM EDTA pH 8.0) for two months before this gel was run. The DNAs in the 'ss' lanes were heated to 95°C for 10 min before loading to separate the strands and reveal the effects of any single-strand breaks.

These DNAs show no sign of degradation; compare to the original photo here. In particular, the DNA fragments from cells grown with limiting phosphate and 40 mM arsenate are actually slightly longer than the fragments from cells grown with limiting phosphate and no arsenate. (I don't think this difference is significant; the important point is that the fragments aren't any shorter.)

Because these large fragments typically migrate at the resolving limit of the gel, all I can say with confidence is that the fragments in all four preps are all significantly larger than 30 kb. This is the size range we expect for chromosomal DNA in a normal DNA prep. I don't have size standards for single-stranded DNA (I should have heated the lambda fragments but forgot to) so all I can say about the length distribution of single strands is that the four preps are all very similar.

This result tells is that DNAs from arsenate-grown cells are not undergoing degradation in storage due to slow hydrolysis of arsenate diester bonds in the DNA backbone, as suggested by an earlier anonymous commenter.

Generating final data for the #arseniclife paper

2012-01-14T05:11:00.000-08:00

1. Cells for new DNA preps: For the replicate DNA preps (for the replicate LC-MS analysis), yesterday I inoculated GFAJ-1 cells into two 50 ml cultures in AML60 medium with 1500 µM PO4, with and without 40 mM AsO4, and into two 500 ml cultures on AML60 medium with 3 µM PO4, with and without 40 mM AsO4. Most of these cultures are growing nicely, so tomorrow I think I'll have enough cells for the DNA preps. Well, the 1500 µMp 40 mM As culture isn't growing at all, but I don't think we need to replicate this one anyway. I need to get at least 50 µg of DNA from each prep, to give the grad student enough for his CsCl gradients. Last time one of the cultures (3 µM PO4, no AsO4) wasn't dense enough to give me the DNA I needed, but so far it looks as dense (or not-dense) as the parallel culture with AsO4. I'll prep the DNA today and if I don't have enough I'll just set up more cultures. I'd be able to prep the DNA from them on Sunday, so still would have the DNAs ready to send on Monday.

2. Troll-suggested control: I've run the gel of the two-month-old DNAs from cells growth with and without arsenic, both native and denatured, and there's no difference in fragment length, with all double-stranded fragments being at least 30 kb in length. So there's no evidence of arsenic-bond strand breakage during long-term storage at 4 °C. I'll post a gel photo later (the image I saved isn't right).

3. Presentable growth curves: A lab in our research cluster has a BioScreen incubator/plate reader I can use to automate my growth curves. But the test cultures I set up in an ordinary microtiter plate aren't growing consistently, so I'll have to mess around a bit before I can do the growth curves.

Writing the #arseniclife paper

2012-01-11T14:13:00.000-08:00

The grad student working on the mass-spectrometry analysis of GFAJ-1 DNA is still making sure his results meet his high standards, but as soon as they are ready he'll send them to me and I'll post them here. In the meantime, since he and his supervisors have concluded that the DNA contains no arsenic, we've started writing our paper. We're going to submit it to Science as a Brevia. These are very short peer-reviewed articles (one page, one figure), which we think suits this work very well.

But first we need to replicate our results. My plan is to generate some detailed growth curves for cultures with various levels of phosphate, with and without 40 mM arsenate. For this I'll use a BioScreen machine that belongs to a neighbouring lab. This machine automates collection of optical density data from cultures growing in wells of 100-well plates. I'll also grow big batches of cells for new DNA preps, using the same media and culture conditions as before.

This should only take a few days, and I hope to have the DNAs ready to send to my collaborators on Monday.

Two steps forward, one step back (the postdoc's uptake bias paper)

2012-01-11T08:54:00.000-08:00

The postdoc's manuscript on uptake bias is inching towards completion. He's added most of the references and updated the figures, and we've only discovered one new analysis that needed to be done. But including this analysis at the right place in the Results makes writing the rest of the Results a lot more straightforward, so we're ahead of the game.

What is this analysis? Removing, from our dataset of 10^7 sequence reads of DNA fragments that the competent cells took up, some sequences that may have been interpreted incorrectly. The incorrect interpretation happens because the sequence responsible for their uptake isn't correctly aligned in our analysis. Here's a figure explaining the problem:

The top sequence is the consensus of the fragment we used. The lower-case bases at each end were not degenerate and function as controls. The first step in the analysis was to align each sequence read to this consensus at its left end, and below the consensus we see three correctly aligned reads, with their core uptake sequence indicated by the yellow arrows.

Below these are two reads that were misaligned because they contained either an insertion or a deletion of a single base. We think these insertions and deletions arose during synthesis of the pool of degenerate fragments. Although these fragments still contain good uptake sequences (red arrows), the incorrect alignment doesn't recognize this. Instead, the fragments appear to have been taken up despite having very poor agreement with the consensus.

Below these misaligned reads is a sequence that is correctly aligned but that contains a second match to the core consensus, indicated by the green arrow. This second match was created by several changes downstream of the consensus uptake sequences, but it isn't recognized by the analysis because it is out of alignment and, in this case, in the other orientation. The presence of two uptake sequences means that we can't attribute their uptake to the one sequence that's correctly aligned.

Sequences with these artefacts couldn't be removed from the dataset before the original analysis, because they couldn't be identified until we were able to score each fragment for matches to the 'uptake motif' that the initial analysis produced. Now that we've identified them, we can consider whether they would have confounded any of the analyses.

The main concern is the reads with insertions or deletions. Because the initial filtering required that the 10 control bases all be perfectly matched, most of these were removed, and the 10^7 recovered reads we analyzed only included about 1500 with insertions or deletions that misaligned the core. That's too few to have misled the initial analysis, but it is a concern for the analyses of possible contamination and sequencing errors, and for the analysis of interaction effects. The postdoc has now finished checking for effects on the interaction analysis (none) and still needs to check for contamination and error effects.

A troll raises a semi-valid concern

2012-01-09T22:02:00.000-08:00

An anonymous troll posted this comment on a previous post:

im curious about how much time the DNA you prepared has spent in solution since you prepped it — supposing there were C-Ar bonds in the DNA at the time of lysis, has so much time passed now that, given the hydrolysis rates that were experimentally determined and recently reported in this JACS communication (offensive link deleted), I'm worried that theres no hope of seeing positive signal even if there was one to begin with. Any thoughts about this issue? Perhaps keeping the DNA stored as a dry pellet and avoiding any encounters with water until the last minute would be the way to go.

First, a few corrections: Arsenic is As, not Ar. The DNA backbone bonds are diester bonds, so the As is bound to oxygen (O), not carbon (C). It's of course not possible to purify DNA while avoiding any encounters with water.

It's indeed theoretically possible that the GFAJ-1 DNA from arsenic-grown cells originally contained As in its backbone, and that the As bonds were destabilized once the DNA was purified away from the proteins and other cellular components. If the chemists are correct, the As bonds would all hydrolyze within less than a second. The AsO4 liberated from the DNA would then be seen just as background in the CsCl gradient, or lost entirely in subsequent purification steps.

We would have to assume that the GFAJ-1 cells contain some powerful unknown-to-science DNA-binding proteins or other structures that stabilize the As bonds in the aqueous environment of the cell. These proteins would be removed during the initial DNA purification steps (SDS-lysis and phenol-extraction). Wolfe-Simon et al. did not report taking any precautions to prevent hydrolysis, so if their DNA prep really contained As bonds these must have been stable for at least the day it would have taken them to do the initial extractions, run the agarose gel, and cut out the DNA-containing gel slices. We don't need to count the time needed to send the gel slices to the LLNL lab at Livermore for Nano-SIMS analysis because the entire slices were analyzed (the DNA was not purified away from them).

Loss of As from the backbone would change the structure of the DNA. It would now be As-free, but would contain a single-strand break at each site that previously had an As. If a substantial fraction of the backbone was As, the DNA would be extensively degraded.

One way to check for hydrolysis of As bonds would be to run the DNAs in a gel, both intact and after separating the strands by boiling. Normal DNA is stable for months, so we could compare the fragment lengths of the DNAs from As-grown and P-grown cells, both immediately after purification and after extended storage in aqueous solution. Here's a diagram of what we'd expect to see in an agarose gel. The upper orange band in the lefthand lanes is the long fragments of double-stranded DNA present in my DNA preps immediately after purification (see gel photo here), and the broader bands on the right are how single-stranded DNA would appear.

Later: Comment send by email:

Since the cell’s interior is aqueous, then it would seem reasonable that there has to be something preventing the spontaneous hydrolysis in vivo. As you point out, some form of stabilizing protein would seem the most likely candidate. If this protein is going to work, then it could (for the purposes of this argument) remain attached throughout the purification process. If so, then you might find As in the purified DNA, and should also find traces of protein. I don’t know if this would be enough to alter the 260:280 ratio, but DNAse digestion followed by SDS-PAGE might detect something. Definitely a hotdog experiment, but once the DNA is purified, not that difficult.

SDS-phenol extractions are pretty harsh; for a protein to remain with the DNA it would need to have been covalently bound, which would certainly have complicated such cellular processes as DNA replication and transcription. And if the DNA contained a significant amount of As, the DNA would then contain enough protein that it wouldn't migrate properly in the gel but would stick as a blob of gunk in the well at the top of the gel. It also would band differently in a CsCl gradient.

How dumb do (a couple of) our students think we are?

2012-01-08T16:52:00.000-08:00

Email I just received from a couple of undergraduates:

HI Dr.Rosie Redfield,

Here's a chance to represent UBC's Zoology Science Department.

The annual Science Week Events Committee , organized by the Science Undergraduate Society, would like to invite you to join our event on Thursday, January 26th, 2012 at 12:30-1:45.

As you may know, Science Week, is a week-long (form January 23rd - January 27th), multi-events celebration which allows students to show off their UBC pride while rewarding them for their first term achievements. So far we have jell-O wrestling, jeopardy, and a scavenger hunt. Although, we realized something was missing...(*1)

This year, we have added a new event to our venue, called the "Professor Pageant". UBC Professors, like yourself, will be participating in the pageant and showcasing their popularity, attitude and talents (*2). Only one competitor will come out on top, however, each participant will go home with a special customized award.

We would like to invite you to join us because you are deemed awesome (yes, awesome) by an overwhelming consensus among students (*3). This is an opportunity that you do not want to miss! (*4)

We would love to hear from you and provide more details. Also, if you would like to suggest any of your colleagues (open to all faculties), please provide their name so we could can invite them accordingly!

Sincerely

(Names withheld to protect the naive.)

(*1): Maybe it's "science"?

(*2): Ooh, why didn't anyone tell me that's what I'm valued for?

(*3) Somehow these students neglected to complete their evaluations of my teaching.

(*4) Oh yes I do!

More power of CsCl gradients

2012-01-03T12:19:00.000-08:00

The grad student pointed out to me by email that I'd overlooked one big advantage of using CsCl gradients to clean up the DNA. He's not analyzing only the fractions that contain DNA, but all the fractions from the gradient. This allows him to detect where in the gradient any arsenic is, and thus lets him distinguish whether the arsenic is bound to the DNA or independent of it. So even a moderate level of arsenic contamination wouldn't be a problem.

Getting ready for some arsenic data

2012-01-01T15:42:00.000-08:00

Any day now I hope to receive some preliminary results from the mass spectrometry test for arsenic in GFAJ-1 DNA. In preparation I though I should at least attempt to understand the control data that the grad student doing the work sent me a couple of weeks ago. But I got sidetracked by the easier task of understanding some control CsCl-gradient data he also sent. This is a pre-analysis step, used to further purify the DNA before the analysis.

What he did: He ran control DNA (from cells grown with lots of phosphate and no arsenate) in two CsCl gradients and collected fractions (~ 100 µl fractions from gradients with total volumes of 1 or 2 ml). He then measured the volume and DNA concentration of each fraction. This showed a nice DNA peak in each gradient (green is the 1 ml gradient, red is the 2 ml gradient). He then pooled the high-DNA fractions of each gradient, desalted them to remove the CsCl, and digested the DNAs in preparation for mass spectrometry (LC-MS).

What I've done: Arithmetic to calculate how much purification these gradients would have accomplished.

The green data: 6778 ng of DNA (89% of the total DNA recovered) is in four fractions with a total volume of 300 µl (37% of the volume recovered). This means that the concentrations of soluble contaminants not bound to the DNA will have been reduced to about 40% of what they were.

The red data: 5135.3 ng of DNA (68% of the total DNA recovered) is in two fractions with a total volume of 310 µl (17% of the total volume recovered). This means that the concentration of soluble contaminants will have been reduced to about 25%.

Hmm, that's not very efficient purification. Larger gradient volumes and longer spins might help. And of course the desalting step should have removed much more of the soluble contaminants.

But this arithmetic may not matter much. The real advantage of the CsCl step is not that it's removing soluble contaminants. Instead, it's fractionating on completely independent principles than the other steps we use, and so it is expected to reduce or remove contaminants that the other methods might not remove. It should remove contaminants that might have coprecipitated with the DNA when it was spooled out of 70% ethanol, and ones that might elute with the DNA in the desalting column because they're insoluble and soluble under the same combinations of conditions as DNA (we typically have to treat these conditions as 'secret sauce' because the manufacturers of the desalting columns don't like to reveal how they work).

In both gradients, contaminants that weren't soluble in the CsCl solution will probably have either pelleted to the bottom of the tube or risen to the top of the tube, depending on their density relative to the CsCl. (As I recall from my undergrad experiences, RNA pellets but proteins rise.) Whether these would have recontaminated the fractions as they were collected depends on how the collection was done. Fractions collected as drops from the bottom of the tube would have avoided contaminants that had risen to the top but would have encountered any pelleted contaminants on their way out.

Do we need to also consider contaminants that might have banded at a specific density in the gradient? The centrifugation is powerful enough to cause the heavy Cs+ ions to move down in the tube, might it also affect the distribution of other ions? What does Wikipedia say? (Ah, the correct term is 'isopycnic centrifugation'.) Nothing about other ions. CsCl gradients have typically been used to separate DNAs with different base compositions from each other (e.g. nuclear DNA from mitochondrial or plastid DNA); I don't know if anyone ever used them to separate DNA from soluble contaminants.

Bottom line: If the LC-MS data shows arsenic in the DNA, we can polish up these DNA purification steps. If it doesn't, we won't need to bother.

Must stop analyzing data....

2012-01-01T14:10:00.000-08:00

The postdoc is back and we are driving each other nuts with ideas for more analyses (both of us), more analyses (him), and requests to stop analyzing the bloody data and finish writing the damned paper (me). Just now I found myself thinking "If only we had less data...".

One simple control analysis we really did need was using his USS-scoring matrices to score some simulated genomes (random-sequence strings of the same length and base composition as the H. influenzae genome). These are controls for the analysis I wrote about here. He's done these now, and they nicely show that both scoring motifs see the bulk of the genome as no different from random sequence, and that the ~200 high-scoring positions they both find are not found in random-sequence 'genomes'.

What to discuss in the Discussion?

2011-12-28T17:28:00.000-08:00

The postdoc gets back from his Christmas break tomorrow, and I plan to have a revised version of his uptake-bias manuscript waiting for him. I've been working through the last part of the Results, and it's looking pretty good. Not finished of course, as there are still two analyses to add. One is the experimental test of the positional interactions predicted by his analysis, comparing uptake of DNA fragments containing single or double mismatches from the consensus USS. I think he's going to do this experiment as soon as he gets back. The other is analysis of out-of-alignment uptake sequences in fragments that were taken up despite lacking a good in-alignment uptake sequence. This issue arises because some fragments contained small insertions or deletions that caused their uptake sequence to be misaligned in the original analysis, and some other fragments may have substitutions that created an uptake sequence at a new location in the fragment. There aren't very many of these out-of-alignment uptake sequences but sorting them out helps clarify some other issues. He's already done the analysis (at least most of it) but we still need to incorporate it into the Results.

The bigger problem is what to say in the Discussion section. Right now it's a shambles, with lots of interesting points and good sentences and well-written paragraphs all jumbled together. I need to get a better perspective on this - to think about what actually should be discussed.

One place to start is the Introduction. Issues we raised there should be addressed and ideally resolved in the Discussion. Unfortunately, our lovely work doesn't really resolve these.

Well, so much for working on the Discussion... I'm now back to asking myself (and, in absentia, the postdoc) questions about how we interpret his analysis of interaction effects: Do interaction effects explain the discrepancy between genomic motif and uptake motif? Do interaction effects support the hypothesis that uptake bias is intrinsic to the mechanism of uptake, and not the effect of a single dedicated recognition protein? And more questions I've added to his lovely figure, so I remember to ask him them tomorrow.

And now, I've got two manuscript reviews (both overdue) and a book review to do.

The postdoc's new analysis saves us work

2011-12-21T20:57:00.001-08:00

Just before he left for a brief Christmas vacation the postdoc did a detailed analysis of the genomic uptake sequences identified by (i) the genomic USS motif identified by the GibbsMotif Sampler and (ii) the DNA uptake motif identified by his sequencing experiment. The two motifs look quite different, and if we applied them both to the same long random DNA sequences we expect that they would pick out different sub-sequences that more-or-less correspond to the motif. But what will they identify in the H. influenzae genome?

We expect the sequences picked out by the genomic USS motif to resemble the search motif (because that's how the motif was identified in the first place). But what will the uptake motif find? It's much simpler, so will it find mainly sequences that just have the four-base inner core GCGG motif?
The analysis is done by sliding the motif across the genome, at each position using the motif to calculate a score for the 32 bases lined up with the motif. This is done with each strand of the 1,830,138 bp genome, so a total of 3,660,276 scores are generated with each motif. The postdoc then plotted a histogram of the scores for each motif.

At this resolution it's no different than you would get for a random DNA sequence. But if we zoom in on the bottom right corner of each graph, we see little blips of about 2000 high-scoring positions. As expected, the sequences of the 1793 positions in the genomic-motif scoring blip give a motif that looks just like the genomic motif we searched with. Unexpectedly, the sequences of the 1892 positions found with the much simpler uptake motif also give a motif a lot like the genomic motif, much more complex than the uptake motif. In fact, the two searches found mostly the same positions; 1689 of the positions in each blip were also present in the other blip.

One of the explanations we were considering for the differences between the two motifs is that the Gibbs Motif Sampler might have unrecognized biases that caused the sequences it identified to not be properly representative of the sequences in the genome. (The most likely candidate is the way we specified the search frame for the Gibbs analysis.) We were going to test this possibility by simulating the evolution of some genomes using each of the motifs in turn, and then test whether Gibbs searches of these evolved genomes gave the original motifs.

But this new result tells us that this possibility is not the explanation for the discrepancy between the two motifs. The uptake sequences in the genome really do look like the full genomic motif, even though the bias of the uptake machinery only cares strongly about the four inner-core bases. I confess that I like this result partly because it saves me from having to run a bunch of USS-evolution simulations to generate sequences for Gibbs analysis.

We suggest three other explanations. First, the steps leading from uptake to recombination might have sequence biases, so that only sequences with the complex motif efficiently recombine. Second, there might be functional constraints on the sequences after they've recombined, so that the complex ones are more likely to become fixed in the population. Although it's certainly likely that some sequence biases and functional constraints do exist, to me it seems very unlikely that they would generate such a complex motif. Thus I prefer our final possibility, that the uptake motif produced by the data is incomplete because it neglects the effects of interactions between the different positions that contributed to uptake (that is, because it incorrectly assumes that each base in the motif acts independently of the others).

We then go on to describe the interaction analysis we've done and the tests we've made (well, the postdoc's about to make) using defined sequences.

The postdoc's DNA uptake paper (the never-ending saga)

2011-12-19T19:46:00.000-08:00

The postdoc and I are back at it yet again, working on his paper about the sequence specificity of DNA uptake. I'm beginning to think there's something pathologically wrong, either with us or with this piece of research, because we never seem to get closer to finishing it. Instead, we just keep discovering more analyses that need to be done. (The part that's done gets better and better, but we seem to be no closer to submission.)

This time it's that we need a more rigorous comparison of the uptake-specificity motif his data has produced with the old 'genomic' motif we derived by analyzing the genome with the Gibbs Motif Sampler. Both motifs consist of numbers representing the probability of finding each of the four bases (A, G, C T) at each position in a 32 bp segment. We've been saying and writing that, although these motifs have the same consensus, they are very different in the importances they ascribe to different positions. We have a list of four possible explanations for the differences, but before we discuss these we need to test whether the motifs actually pick out different subsets of the genome. Maybe all of the ~2500 sequences that would be found by searching for the genomic motif would also be found by searching for the less-constraining uptake motif. If so, we might then focus on what other sequences the uptake motif found, or, if it didn't find any, why not.

No NovR colonies at all?

2011-12-19T18:27:00.000-08:00

Yesterday I made 8 preps of competent cells, all to further our phenotyping of our new competence-gene mutants. Four of them needed to have transformation assays done, three were to be frozen for later DNA-uptake assays by the postdoc*, and one was a knockout of the competence-regulator sxy, to be used as a negative control in the uptake assays.

I didn't include a wild-type positive control strain for my transformation assays, because I've done this lots of times before. But I did assay the sxy mutant, just to confirm that I had the right strain. The assay is simple: mix 1 ml of competent cells (~10^9 cells) with 1 µg of NovR chromosomal DNA, incubate for 15 min, add 10 µg DNase I, incubate for 5 min, dilute and plate on plain sBHI agar and on sBHI agar with 2.5 µg novobiocin/ml.

Most of the strains I assayed had been tested before, but some with not-very-consistent results, and I was expecting to see a wide range of transformation frequencies (maximum about 5 x 10^-3 and minimum less than 10^-8 (the detection limit)). Because I wasn't sure what I would find, I made a point of plating 100 µl of undiluted culture from each assay. BUT, there were absolutely no NovR colonies on any of the plates.

So I'm pretty sure I screwed something up. But what? I used the same DNA stock tube I've used many times before, and I definitely remember putting 3 µl of DNA into each assay tube. I made fresh sBHI + novobiocin plates using pre-made BHI agar,, and I definitely remember adding the hemin (4 ml), NAD (80 µl) and novobiocin (40 µl) to the melted agar before I poured the plates. The DNaseI should be fine; I've used this tube before. And the cells aren't dead, as the plain sBHI plates had the expected numbers of colonies. Oh how I wish I'd included the positive control! Luckily I froze one tube of competent culture of each of the strains I transformed, 'just in case', so I can redo the transformations without having to make now competent preps.

To check if I somehow screwed up the agar plates despite my 'definite' memory, I've streaked a test known NovR strain on them, with and without more NAD or hemin. Before I go home I should set up some overnight cultures of the strains I'm going to test tomorrow... Wait, will I have time to do this tomorrow? It takes time and planning to get the cells into the right growth stage for the competence treatment, and then a couple of hours for competence development and the transformation assays. I have a meeting in the middle of the day, and then we're going to finalize the grades for the big genetics course... Yes, sure, I can always work late.

* The postdoc is getting interesting results from these assays. First, all of the mutants he's tested, even those lacking genes thought to be essential for DNA uptake bind/take up at least fourfold more DNA than the noncompetent log-phase cells he's using as a negative control. The competent sxy- cells I've just made won't induce any competence genes in this medium, so the amount of DNA they associate with will tell us whether this binding is just a property of cells that have been incubated in the starvation medium, or represents induction of some competence genes.

For a couple of the mutants, he's found much higher DNA association levels than we expected. One mutant should lack the secretin pore through which the pseudopilus contacts the DNA; the other lacks PilF2, an 'accessory' pilin that's absolutely required for transformation, presumably because it's absolutely required for DNA uptake. The next step is to repeat the DNA uptake assays, this time comparing treatments with and without DNase I in the wash step. If the mutant cells are binding to DNA but not taking it up, the DNase I should remove the DNA.

UPDATE: My novobiocin plates had no NovR colonies because I had forgotten to add the required hemin supplement to the agar! How embarrassing - I haven't made that mistake in years.

Felisa Wolfe-Simon's poster at the Dec. 2011 AGU meeting

2011-12-16T06:19:00.000-08:00

I just found the Abstract for a poster presented by Felisa Wolfe-Simon at this month's American Geophysical Union Annual Meeting.

TITLE: Characterizations of intracellular arsenic in a bacterium
SESSION TYPE: Poster
SESSION TITLE: B51G. Life Under Stress: How Do Microbes Cope?
AUTHORS: Felisa Wolfe-Simon, Steven M. Yannone, John A. Tainer^. Life Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States.
ABSTRACT: Life requires a key set of chemical elements to sustain growth. Yet, a growing body of literature suggests that microbes can alter their nutritional requirements based on the availability of these chemical elements. Under limiting conditions for one element microbes have been shown to utilize a variety of other elements to serve similar functions often (but not always) in similar molecular structures. Well-characterized elemental exchanges include manganese for iron, tungsten for molybdenum and sulfur for phosphorus or oxygen. These exchanges can be found in a wide variety of biomolecules ranging from protein to lipids and DNA. Recent evidence suggested that arsenic, as arsenate or As(V), was taken up and incorporated into the cellular material of the bacterium GFAJ-1. The evidence was interpreted to support As(V) acting in an analogous role to phosphate. We will therefore discuss our ongoing efforts to characterize intracellular arsenate and how it may partition among the cellular fractions of the microbial isolate GFAJ-1 when exposed to As(V) in the presence of various levels of phosphate. Under high As(V) conditions, cells express a dramatically different proteome than when grown given only phosphate. Ongoing studies on the diversity and potential role of proteins and metabolites produced in the presence of As(V) will be reported. These investigations promise to inform the role and additional metabolic potential for As in biology. Arsenic assimilation into biomolecules contributes to the expanding set of chemical elements utilized by microbes in unusual environmental niches.

The work it describes is new, as it was done in John Tainer's lab at Lawrence Berkeley. Unfortunately there's not much meat. That's not surprising, since poster abstracts typically have to be submitted months in advance and the deadline for AGU seems to have been August 4. I can't find any tweets or other information about this poster - did anyone see it?

Some control results! (Don't get excited, it's just a control...)

2011-12-14T17:59:00.000-08:00

My collaborators have taken pity on me and sent me some of their control analysis data. This is mass-spectrometry analysis of a control DNA sample I sent several months ago.

The GFAJ-1 cells this DNA was purified from were grown in medium without arsenic, so we don't expect to find any arsenic in the DNA. This DNA was just used to test the methods they will use, but it also provides some measure of the purity of the DNA I sent them. That's because this DNA hasn't been put through the CsCl-density-gradient purifications step that they'll use for the new DNA samples I sent a few weeks ago.

I'm going to have to put in a bit of work before I have any idea of what I'm looking at. But I'm not complaining - this is exactly the part of science I like best.

What's happening with the GFAJ-1 DNA

2011-12-14T13:43:00.000-08:00

I've just gotten an update from my collaborators. They're still polishing up their purification methods, and waiting for the mass spectrometer machine-time that will open up when other researchers take time off over Christmas.

So with luck we'll have a preliminary answer in a few weeks. Then we'll have to get busy generating the final data and writing our paper!

Growth of GFAJ-1 in arsenate

2011-11-25T17:33:00.000-08:00

I've now tested whether the growth of GFAJ-1 is indeed stimulated by arsenate, as was suggested by the yields of my DNA-prep cultures. This time I was very careful to keep the ionic strength constant, giving each culture tube the same volume of varying mixtures of 1 M NaAsO4 and 1 M NaCl. The arsenic concentrations ranged from 5 µM to 60 mM in 2-fold or 2.5-fold steps (also 0 mM).

To make sure that all culture tubes started with the same medium and the same density of cells, I mixed up a big batch of no-phosphate medium, added cells (2 x 10^5 cfu/ml), and divided the culture into thirds. I then added NaPO4 to two of these parts, to give 3 µM and 1500 µM (in addition to whatever phosphate might be contaminating the medium), and added 5 ml of each part to each of 15 screw-cap galss tubes to which I had already added the appropriate NaAsO4/NaCl mixture. So I had 45 tubes in all, 15 with no added phosphate, 15 with 3 µM, and 15 with 1500 µM.

I incubated the tubes at 28 °C with gentle rocking, and checked the optical density after 24 and 48 hr.

Conclusion: Arsenate stimulated growth, but didn't affect the final densities of the cultures. The stimulation is not because the arsenate is contaminated with phosphate, because the effect was strong only in the cultures with 1500 µM added phosphate, and because it didn't affect final density in the phosphate-limited cultures.

There's still much more variation in final culture density than I'd like to see. This might be due to minor differences in trace contaminants in the tubes, although they were all last used for similar cultures and all thoroughly washed the same way. One solution would be to use only new tubes, but these tubes are not cheap and I don't want to take money from our transformation work.

I'm not going to do any more work on this - not going to do experiments to find out why arsenate stimulates growth, unless the mass spec shows that there really is arsenic in the DNA of arsenate-grown cells. The growth stimulation I'm seeing isn't a replication of Wolfe-Simon et al's report that their cultures grew with arsenate but not without it, but it might reflect the same biological process.