Our application to NIH is rated unscored.
This means that it was ranked in the bottom half of all submissions, based on the evaluations by the reviewers who read it before the Review Panel meeting, so it was not discussed at the meeting. The only reviews we'll get will be those prepared by these reviewers before the meeting (we'll get them in a couple of months). There's probably not much point in working on a resubmission till then.
Later: And as we're only allowed one resubmission, we need to make sure it's as strong as possible. Better to take our time and do it right.
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
Back to thinking about purine regulation of sxy translation
I spent part of yesterday reading over posts from earlier this spring about how purine nucleotides and the PurR repressor might contribute to competence regulation. I then started editing an old post to make it clearer, but now I'm elevating those edits to 'New Post' status here. I'm continuing to edit them to make my thinking as clear as possible (more edits July 19).
I think it's important to consider the interdependent effects of at least two factors that affect nucleotide pools (1) the extracellular and intracellular concentrations of purine precursors, especially guanine and hypoxanthine, which control the repressing activity of the PurR repressor, and (2) the intracellular concentrations of purine nucleotides (ATP and GTP) that are available for transcription. Another issue is the cell-to-cell variation in the amounts of cAMP, sxy mRNA and Sxy protein due to random fluctuations in transcription of the sxy gene, translation of sxy mRNA and the activities of various catalytic enzymes.
Here are the paragraphs I've been working on:
So what am I hypothesizing? When wildtype cells grow to high density in sBHI, cAMP levels become high in most or all cells because the phosphotransferase system senses that preferred sugars are scarce and activates adenylate cyclase. This cAMP activates CRP, causing high transcription of sxy, but most of the sxy transcripts are not translated because their mRNA has folded into an inhibitory secondary structure. In a small fraction of the cells in high-density cultures, this structure either doesn't form or doesn't prevent translation of sxy mRNA, whether by chance or because random fluctuations in the supply of purine nucleotides have slowed transcription. The combination of high Sxy and high cAMP then causes these cells to express competence genes and become competent. However, most cells in these cultures don't express enough Sxy protein to turn on competence genes.
In these cells the PurR repressor is active and the purine-biosynthesis genes are off because of high cytoplasmic concentrations of guanine and hypoxanthine, obtained directly from the medium or by conversion of other purine precursors.
Competence levels are different in dense cultures of hypercompetent-sxy mutants and of purR mutants. In hypercompetent-sxy mutants the supply of nucleotides doesn't matter because the mutations have destabilized the sxy mRNA secondary structure, so all the cells with high cAMP express enough Sxy protein to become competent when the culture becomes dense. (Even in low-density cultures, many cells with these mutations express enough Sxy to become competent.) The opposite effect is seen in the purR mutant, where the constitutive activation of the purine biosynthetic genes keeps levels of purine nucleotides relatively high, so few cells translate enough Sxy to become competent at high density, even though high cAMP levels are causing sxy transcription. These effects might cancel out in the purR- hypercompetent-sxy double mutants, because the sxy transcripts are still efficiently translated regardless of levels of purine nucleotides, but the exact effect will depend on the relative strengths of the effects of the two mutations.
Continuing the hypothesis -- effects in MIV: When wildtype cells in low-density ('log-phase) cultures are abruptly transferred to the starvation medium MIV, the cells are suddenly cut off from the nucleotide precursors they've been getting from the culture medium. This causes a rapid fall in the supply of purine nucleotides AMP and GMP. These nucleotides cannot be immediately synthesized from scratch ('de novo') because the genes have until now been repressed by PurR. cAMP levels also rise sharply on transfer to MIV, inducing sxy transcription, and the shortage of purine nucleotides allows efficient sxy translation so that most cells become competent. The lack of purine precursors also inactivates PurR, so the de novo biosynthesis genes are turned on, at least partially replenishing the supply of AMP and GMP.
What would happen if AMP or GMP was added to the MIV? (I think these nucleotides are readily interconverted, so only one is needed.) Even though nucleotides must be converted to nucleosides (have their phosphates removed) for transport across the cell membrane, I hypothesize that they have a more direct effect on nucleotide pools than do simpler base precursors such as inosine, guanine, or hypoxanthine, which require extensive biochemical processing to be converted to nucleotides. Thus supplementing MIV with AMP or GMP maintains their cytoplasmic concentrations above the threshold, preventing translation of sxy mRNA and thus the development of competence in most cells.
What about mutants? Hypercompetent-sxy mutants in MIV are expected to experience the same fall of nucleotides and rise of cAMP, and their insensitivity to nucleotide supply won't matter when the supply is already depleted. This explains why they reach the same level of competence in MIV as wildtype cells.
What about MIV competence in the purR knockout? The purR- cells have already made the enzymes for synthesizing purine nucleotides de novo, and our original thinking was that this would maintain high enough AMP and GMP concentrations to prevent translation of sxy mRNA when the external supply of precursors was removed. Thus we expected these cells to respond to MIV like wildtype cells do to high density in sBHI, with only a small fraction of cells becoming competent. But this prediction wasn't met; the purR cells became just as competent in MIV as wildtype cells. An alternative hypothesis is that de novo synthesis of purine nucleotides in the purR mutant isn't enough to compensate for the sudden removal of the exogenous supply of precursors (i.e. the salvage pathways make a bigger contribution than the de novo pathway). It's also possible that the de novo pathway is subject to feedback regulation that limits its contribution when salvage is active, even though the genes are fully expressed.
What to do? I can't immediately start using the purR and purH knockouts to test these ideas, because these mutants aren't ready to use yet. Is there another way to test the hypothesis that de novo synthesis is less effective than salvage?
I think it's important to consider the interdependent effects of at least two factors that affect nucleotide pools (1) the extracellular and intracellular concentrations of purine precursors, especially guanine and hypoxanthine, which control the repressing activity of the PurR repressor, and (2) the intracellular concentrations of purine nucleotides (ATP and GTP) that are available for transcription. Another issue is the cell-to-cell variation in the amounts of cAMP, sxy mRNA and Sxy protein due to random fluctuations in transcription of the sxy gene, translation of sxy mRNA and the activities of various catalytic enzymes.
Here are the paragraphs I've been working on:
So what am I hypothesizing? When wildtype cells grow to high density in sBHI, cAMP levels become high in most or all cells because the phosphotransferase system senses that preferred sugars are scarce and activates adenylate cyclase. This cAMP activates CRP, causing high transcription of sxy, but most of the sxy transcripts are not translated because their mRNA has folded into an inhibitory secondary structure. In a small fraction of the cells in high-density cultures, this structure either doesn't form or doesn't prevent translation of sxy mRNA, whether by chance or because random fluctuations in the supply of purine nucleotides have slowed transcription. The combination of high Sxy and high cAMP then causes these cells to express competence genes and become competent. However, most cells in these cultures don't express enough Sxy protein to turn on competence genes.
In these cells the PurR repressor is active and the purine-biosynthesis genes are off because of high cytoplasmic concentrations of guanine and hypoxanthine, obtained directly from the medium or by conversion of other purine precursors.
Competence levels are different in dense cultures of hypercompetent-sxy mutants and of purR mutants. In hypercompetent-sxy mutants the supply of nucleotides doesn't matter because the mutations have destabilized the sxy mRNA secondary structure, so all the cells with high cAMP express enough Sxy protein to become competent when the culture becomes dense. (Even in low-density cultures, many cells with these mutations express enough Sxy to become competent.) The opposite effect is seen in the purR mutant, where the constitutive activation of the purine biosynthetic genes keeps levels of purine nucleotides relatively high, so few cells translate enough Sxy to become competent at high density, even though high cAMP levels are causing sxy transcription. These effects might cancel out in the purR- hypercompetent-sxy double mutants, because the sxy transcripts are still efficiently translated regardless of levels of purine nucleotides, but the exact effect will depend on the relative strengths of the effects of the two mutations.
Continuing the hypothesis -- effects in MIV: When wildtype cells in low-density ('log-phase) cultures are abruptly transferred to the starvation medium MIV, the cells are suddenly cut off from the nucleotide precursors they've been getting from the culture medium. This causes a rapid fall in the supply of purine nucleotides AMP and GMP. These nucleotides cannot be immediately synthesized from scratch ('de novo') because the genes have until now been repressed by PurR. cAMP levels also rise sharply on transfer to MIV, inducing sxy transcription, and the shortage of purine nucleotides allows efficient sxy translation so that most cells become competent. The lack of purine precursors also inactivates PurR, so the de novo biosynthesis genes are turned on, at least partially replenishing the supply of AMP and GMP.
What would happen if AMP or GMP was added to the MIV? (I think these nucleotides are readily interconverted, so only one is needed.) Even though nucleotides must be converted to nucleosides (have their phosphates removed) for transport across the cell membrane, I hypothesize that they have a more direct effect on nucleotide pools than do simpler base precursors such as inosine, guanine, or hypoxanthine, which require extensive biochemical processing to be converted to nucleotides. Thus supplementing MIV with AMP or GMP maintains their cytoplasmic concentrations above the threshold, preventing translation of sxy mRNA and thus the development of competence in most cells.
What about mutants? Hypercompetent-sxy mutants in MIV are expected to experience the same fall of nucleotides and rise of cAMP, and their insensitivity to nucleotide supply won't matter when the supply is already depleted. This explains why they reach the same level of competence in MIV as wildtype cells.
What about MIV competence in the purR knockout? The purR- cells have already made the enzymes for synthesizing purine nucleotides de novo, and our original thinking was that this would maintain high enough AMP and GMP concentrations to prevent translation of sxy mRNA when the external supply of precursors was removed. Thus we expected these cells to respond to MIV like wildtype cells do to high density in sBHI, with only a small fraction of cells becoming competent. But this prediction wasn't met; the purR cells became just as competent in MIV as wildtype cells. An alternative hypothesis is that de novo synthesis of purine nucleotides in the purR mutant isn't enough to compensate for the sudden removal of the exogenous supply of precursors (i.e. the salvage pathways make a bigger contribution than the de novo pathway). It's also possible that the de novo pathway is subject to feedback regulation that limits its contribution when salvage is active, even though the genes are fully expressed.
What to do? I can't immediately start using the purR and purH knockouts to test these ideas, because these mutants aren't ready to use yet. Is there another way to test the hypothesis that de novo synthesis is less effective than salvage?
Cells behaving badly
I'm still having problems with the E. coli cells that contain the DNA construct for the advanced tweezers experiments. The cells grew OK overnight on an LB+Amp50 agar plate but poorly in what I later realized was LB+Amp200 broth (too much ampicillin). I did a plasmid prep from these cells but got little or no plasmid (a faint smear only). Yesterday the postdoc inoculated another colony for me, into LB+Amp50, but this also stopped growing at a low density, and a plasmid prep gave no plasmid at all.
The E. coli cells are strain DH5alpha, which should be quite vigorous, and the plasmid is a derivative of pGEM and so should have a high copy number, even with a 12 kb insert. Perhaps the cells aren't what they're supposed to be. Maybe there's something wrong with my medium (and the postdoc's medium). Perhaps there was something wrong with my original LB+Amp plate. Perhaps the plasmid replicates poorly because it's so big. Perhaps the plasmid is present in the cells but lost during purification, because the miniprep kit doesn't give good recovery for DNAs bigger than ~10 kb.
Update: this morning i inoculated colonies from the LB+Amp plate into plain LB and LB+Amp50. The cells in plain LB seem to be growing faster, so maybe the plasmid is toxic. I'd better go back and reread the thesis chapter of the M.Sc. student who made it.
The E. coli cells are strain DH5alpha, which should be quite vigorous, and the plasmid is a derivative of pGEM and so should have a high copy number, even with a 12 kb insert. Perhaps the cells aren't what they're supposed to be. Maybe there's something wrong with my medium (and the postdoc's medium). Perhaps there was something wrong with my original LB+Amp plate. Perhaps the plasmid replicates poorly because it's so big. Perhaps the plasmid is present in the cells but lost during purification, because the miniprep kit doesn't give good recovery for DNAs bigger than ~10 kb.
Update: this morning i inoculated colonies from the LB+Amp plate into plain LB and LB+Amp50. The cells in plain LB seem to be growing faster, so maybe the plasmid is toxic. I'd better go back and reread the thesis chapter of the M.Sc. student who made it.
Success, and maybe another reason why beads clump
I did a test of bead clumping. I started with my prep of chromosomal DNA that had been cut with XhoI and biotin-tagged at both ends of the fragments. I then cut this DNA with EcoRI, which will convert many of the fragments into shorter fragments with biotin-tags at only one end. Then I incubated this DNA with beads overnight, using either a very concentrated mixture (20 µl each of beads and DNA, in a total volume of 100 µl) or a 50-fold more dilute mixture (40 µl each of beads and DNA, in a total volume of 10 ml). The next morning I found almost no clumping, even less than at the beginning of the incubation.
But I had made one other change, in addition to the EcoRI digestion, which caused them to be much more vigorously mixed. Previously I had been placing the tubes containing the mixtures into the slots on our roller wheel, where they would rotate around their long axes with their bottoms always slightly lower than their tops. This allowed some of the beads to eventually settle at the bottom of the tube. This time I taped the tubes onto the sides of the wheel so they would rotate around their short axes, being turned completely upside down and then right side up with every rotation of the wheel. I don't know if this improved mixing also contributed to the elimination of clumps.
I had done these incubations with both magnetic Dynabeads and polystyrene beads. About half the Dynabeads went missing somewhere along the line (or maybe I accidentally put in less than I had intended). I tested using a hemocytometer to directly count the number of beads in a defined volume. This worked well, even though it's designed for cells much bigger than my beads, and I now know that I have about 2 x 10^7 of each type of bead, coated with DNA. But I don't yet know how much DNA is on the beads, and I'll have to use up a substantial fraction of the beads for the Picogreen assay to measure this.
I also grew up the strain carrying the DNA construct plasmid. It grew fine on a LB+Amp (ampicillin) plate, but not in LB +Amp broth. But I now suspect I may have put too much Amp into the broth.
But I had made one other change, in addition to the EcoRI digestion, which caused them to be much more vigorously mixed. Previously I had been placing the tubes containing the mixtures into the slots on our roller wheel, where they would rotate around their long axes with their bottoms always slightly lower than their tops. This allowed some of the beads to eventually settle at the bottom of the tube. This time I taped the tubes onto the sides of the wheel so they would rotate around their short axes, being turned completely upside down and then right side up with every rotation of the wheel. I don't know if this improved mixing also contributed to the elimination of clumps.
I had done these incubations with both magnetic Dynabeads and polystyrene beads. About half the Dynabeads went missing somewhere along the line (or maybe I accidentally put in less than I had intended). I tested using a hemocytometer to directly count the number of beads in a defined volume. This worked well, even though it's designed for cells much bigger than my beads, and I now know that I have about 2 x 10^7 of each type of bead, coated with DNA. But I don't yet know how much DNA is on the beads, and I'll have to use up a substantial fraction of the beads for the Picogreen assay to measure this.
I also grew up the strain carrying the DNA construct plasmid. It grew fine on a LB+Amp (ampicillin) plate, but not in LB +Amp broth. But I now suspect I may have put too much Amp into the broth.
Why beads clump, and ways to prevent or minimize it
My preps of beads+DNA (biotin-tagged DNA linked to streptavidin-coated polystyrene or magnetic beads) typically contain dense clumps of beads - sometimes more of the beads are in these clumps than are solitary or in small groups. This is a big problem, partly because the tweezers work requires isolated beads and partly because I can't accurately measure the properties of the preps.
It's pretty clear that the clumping is caused by the biotin-tagged DNA. Adding DNase I to the preps breaks up the clumps, and there are no big clumps in preps of washed beads (no DNA) or if the beads are incubated with DNA that hasn't been biotin-tagged. The likely cause is that, if the tagging reaction worked well, both ends of each DNA fragment should have biotin tags. Such fragments can then bind to two beads, crosslinking them, and may also create DNA loops that entrap other DNA fragments.
I can see several ways to reduce this problem, illustrated in the figure.
1. One strategy is to use a defined DNA substrate rather than restriction-digested chromosomal DNA, one that can be biotin-tagged at only one end. A 14 kb plasmid designed for such experiments is available, and the biophysics grad student who worked on this project modified added a H. influenzae uptake sequence to it. It's in our freezer. All I need to do is digest with XhoI, biotin-tag the ends, and then digest with EcoRI to snip off the tag at the USS end. I won't need to purify the long fragment, as the snipped-off bit shouldn't interfere with anything. The EcoRI site is only 6 bp from the Xho site; luckily the ever-useful New England Biolabs catalog tells me that EcoRI doesn't mind cutting very close to an end. Because this fragment contains only a single uptake sequence rather than the ~1 uptake sequence per kb of chromosomal DNA, it might not be as useful for the preliminary experiments whose goal is just to get cells to bind to DNA on beads and begin uptake.
2. Another strategy is to use chromosomal DNA but reduce the number of fragments that are biotinylated at both ends, by cutting with a second restriction enzyme, one that doesn't give the appropriate DNA overhangs for the biotinylation reaction. The first enzyme should be one that cuts infrequently, giving fragments that are mostly larger than, say, 20 kb. The second enzyme should be one that cuts more frequently. Because restriction sites are randomly positioned on chromosomal DNA. this second digest would cut many but probably not all of the doubly-tagged fragments, so I'd need to find the best balance between leaving some uncut and cutting too often, producing very short singly tagged fragments and many untagged fragments. The presence of untagged fragments is probably not a serious problem, given the high affinity of streptavidin for biotin, but DNA uptake may not be easy to detect if the fragments are too short.
3. The third strategy is technically simplest, although it's the last one I thought of. I've been incubating the DNA with the beads in a small volume. For most kinds of reactions, using a high concentration of reactants is good because it increases the frequency of interactions. But streptavidin-biotin interactions have such strong affinities that this may not be an issue, especially because I can let the reactions proceed for a long time. On the other hand, once a doubly tagged DNA fragment has bound a bead at one end, using a dilute mixture will increase the chance that the biotin at the fragment's other end will bind to the same bead before it encounters a different bead.
4. (not shown) As a stopgap solution I can take the bead+DNA preps that I have and let the big clumps settle out.
It's pretty clear that the clumping is caused by the biotin-tagged DNA. Adding DNase I to the preps breaks up the clumps, and there are no big clumps in preps of washed beads (no DNA) or if the beads are incubated with DNA that hasn't been biotin-tagged. The likely cause is that, if the tagging reaction worked well, both ends of each DNA fragment should have biotin tags. Such fragments can then bind to two beads, crosslinking them, and may also create DNA loops that entrap other DNA fragments.
I can see several ways to reduce this problem, illustrated in the figure.
1. One strategy is to use a defined DNA substrate rather than restriction-digested chromosomal DNA, one that can be biotin-tagged at only one end. A 14 kb plasmid designed for such experiments is available, and the biophysics grad student who worked on this project modified added a H. influenzae uptake sequence to it. It's in our freezer. All I need to do is digest with XhoI, biotin-tag the ends, and then digest with EcoRI to snip off the tag at the USS end. I won't need to purify the long fragment, as the snipped-off bit shouldn't interfere with anything. The EcoRI site is only 6 bp from the Xho site; luckily the ever-useful New England Biolabs catalog tells me that EcoRI doesn't mind cutting very close to an end. Because this fragment contains only a single uptake sequence rather than the ~1 uptake sequence per kb of chromosomal DNA, it might not be as useful for the preliminary experiments whose goal is just to get cells to bind to DNA on beads and begin uptake.
2. Another strategy is to use chromosomal DNA but reduce the number of fragments that are biotinylated at both ends, by cutting with a second restriction enzyme, one that doesn't give the appropriate DNA overhangs for the biotinylation reaction. The first enzyme should be one that cuts infrequently, giving fragments that are mostly larger than, say, 20 kb. The second enzyme should be one that cuts more frequently. Because restriction sites are randomly positioned on chromosomal DNA. this second digest would cut many but probably not all of the doubly-tagged fragments, so I'd need to find the best balance between leaving some uncut and cutting too often, producing very short singly tagged fragments and many untagged fragments. The presence of untagged fragments is probably not a serious problem, given the high affinity of streptavidin for biotin, but DNA uptake may not be easy to detect if the fragments are too short.3. The third strategy is technically simplest, although it's the last one I thought of. I've been incubating the DNA with the beads in a small volume. For most kinds of reactions, using a high concentration of reactants is good because it increases the frequency of interactions. But streptavidin-biotin interactions have such strong affinities that this may not be an issue, especially because I can let the reactions proceed for a long time. On the other hand, once a doubly tagged DNA fragment has bound a bead at one end, using a dilute mixture will increase the chance that the biotin at the fragment's other end will bind to the same bead before it encounters a different bead.
4. (not shown) As a stopgap solution I can take the bead+DNA preps that I have and let the big clumps settle out.
CIHR results
We got the score and reviews for our CIHR grant proposal on DNA uptake this morning.
Score: 4.37
Rank: #10 out of 47 proposals reviewed
Percentile: 21.28% (yes, they report four significant figures)
Chance of funding: Borderline
We'll find out whether it's funded in a few weeks.
(If you want to read the proposal, it's here.)
Score: 4.37
Rank: #10 out of 47 proposals reviewed
Percentile: 21.28% (yes, they report four significant figures)
Chance of funding: Borderline
We'll find out whether it's funded in a few weeks.
(If you want to read the proposal, it's here.)
DNA is stably attached to beads
OK, after some delay due to putting the microtiter plate wrong-way-around into the plate-scanning machine, I have the results of the DNA assays of my bead+DNA preps before and after a final wash.
I think the diagram is pretty self-explanatory. Most important result: Almost all of of the DNA is firmly attached to the beads. Even for the magnetic Dynabeads (columns on the right), the volume of DNA-containing supernatant remaining with the washed beads contributes no more than 1% of the total DNA in the -after-wash prep. The three preps on the right have fewer but larger fragments than the three preps on the left.
Surprisingly, the Picogreen assay detected consistently more DNA on the beads after washing than before washing. This is probably an artefact of something. The beads were all originally in TE (10 mM Tris, 1 mM EDTA), and were washed in TE overnight and resuspended in TE. Might the overnight rolling at 37 °C have better dispersed the DNA so the Picogreen could access it easier? This seems unlikely to be the explanation as, because of my putting-the plate-in-backwards error, all the samples were mixed with Picogreen and then sat for 24 hr before being read.
Because the tweezers work doesn't need millions of beads, these preps should be enough to go on with. This characterization also gives me confidence that my biotin-tagging and bead-attaching procedures work reliably.
Next steps: Repeat the transformation and cell-attachment assays. But first, clarify the bead-clumping situation.
I think the diagram is pretty self-explanatory. Most important result: Almost all of of the DNA is firmly attached to the beads. Even for the magnetic Dynabeads (columns on the right), the volume of DNA-containing supernatant remaining with the washed beads contributes no more than 1% of the total DNA in the -after-wash prep. The three preps on the right have fewer but larger fragments than the three preps on the left.
Surprisingly, the Picogreen assay detected consistently more DNA on the beads after washing than before washing. This is probably an artefact of something. The beads were all originally in TE (10 mM Tris, 1 mM EDTA), and were washed in TE overnight and resuspended in TE. Might the overnight rolling at 37 °C have better dispersed the DNA so the Picogreen could access it easier? This seems unlikely to be the explanation as, because of my putting-the plate-in-backwards error, all the samples were mixed with Picogreen and then sat for 24 hr before being read.
Because the tweezers work doesn't need millions of beads, these preps should be enough to go on with. This characterization also gives me confidence that my biotin-tagging and bead-attaching procedures work reliably.
Next steps: Repeat the transformation and cell-attachment assays. But first, clarify the bead-clumping situation.
How much washing is enough?
Today I'm testing my preps of beads with DNA bound to them by streptavidin-biotin linkages. I want to know how much DNA is on the beads, and to be sure that this DNA is held by a streptavidin-biotin bond at one or both ends, rather than being passively trapped by other bead-DNA linkages.
I had four preps I'm made previously, two to EcoRI-cut DNA and two to XhoI-cut DNA; the average fragment sizes are about 6 kb and about 12 kb respectively. I also have two new preps of a mixture of both DNAs, bound to magnetic Dynabeads (either 2.8 µ M280 beads or 1 µ T1 beads. All of the preps were washed three times with about 100:1 volumes of TE, and resuspended in 100 µl of TE. DNA concentrations have only been measured for one of the EcoRI and one of the XhoI preps.
The goal is to find out whether the intervening incubation and an additional thorough washing (rotating overnight in 10 volumes of TE) will remove more DNA from the beads.
So last night I measured the volume of each bead+DNA prep and took a 5 µl sample, to be assayed today for DNA concentration using the Picogreen assay. The beads+DNA I did assay had about 250 ng DNA/ml, so this 5 µl was chosen to give enough sensitivity without using up too much of my preps. Then I added 9 volumes of TE to each prep and put the tubes on the roller wheel in the 37°C incubator overnight. Now I am going to pellet the beads (centrifuge or magnet), remove the supernatant, and resuspend the beads at their original concentration (= original volume - 5 µl). Then I'll take 5 µl of the beads+DNA and 25 µl of each supernatant for Picogreen assay.
For assay standards I have lambda DNA at 1000, 100, 10 and 1 ng/µl, and lambda DNA mixed with beads at known concentrations (to check that the beads don't interfere with the assay).
I hope to find that most of the DNA is still associated with the beads.
I had four preps I'm made previously, two to EcoRI-cut DNA and two to XhoI-cut DNA; the average fragment sizes are about 6 kb and about 12 kb respectively. I also have two new preps of a mixture of both DNAs, bound to magnetic Dynabeads (either 2.8 µ M280 beads or 1 µ T1 beads. All of the preps were washed three times with about 100:1 volumes of TE, and resuspended in 100 µl of TE. DNA concentrations have only been measured for one of the EcoRI and one of the XhoI preps.
The goal is to find out whether the intervening incubation and an additional thorough washing (rotating overnight in 10 volumes of TE) will remove more DNA from the beads.
So last night I measured the volume of each bead+DNA prep and took a 5 µl sample, to be assayed today for DNA concentration using the Picogreen assay. The beads+DNA I did assay had about 250 ng DNA/ml, so this 5 µl was chosen to give enough sensitivity without using up too much of my preps. Then I added 9 volumes of TE to each prep and put the tubes on the roller wheel in the 37°C incubator overnight. Now I am going to pellet the beads (centrifuge or magnet), remove the supernatant, and resuspend the beads at their original concentration (= original volume - 5 µl). Then I'll take 5 µl of the beads+DNA and 25 µl of each supernatant for Picogreen assay.
For assay standards I have lambda DNA at 1000, 100, 10 and 1 ng/µl, and lambda DNA mixed with beads at known concentrations (to check that the beads don't interfere with the assay).
I hope to find that most of the DNA is still associated with the beads.
Ionic strength?
My physicist collaborator asked me the ionic strength of the MIV medium I'll be using for my optical tweezers experiments. This is significant because the ions in the medium may shield the DNA from surface charges on the glass slide and cover slip, and so reduce unwanted binding. I knew roughly what ionic strength is, but I had to look up the formula in Wikipedia. I then looked up the recipe for 'solution 21'(the main component of MIV), looked up the molecular weights, and did the arithmetic. The ionic strength of MIV is 0.5 M. I think this is high enough to give lots of electrostatic shielding.
While I was at it I also calculated the weight/volume concentration of MIV; it's about 3.5% (amino acids and salts), which I think is not high enough to interfere with the refraction needed by the tweezers.
While I was at it I also calculated the weight/volume concentration of MIV; it's about 3.5% (amino acids and salts), which I think is not high enough to interfere with the refraction needed by the tweezers.
Did the cells attach to the DNA-coated beads?
Yesterday I scored the results of Wednesday's test. I had incubated competent H. influenzae cells with magnetic beads, using both DNA-coated beads and control beads that had gone through the same treatment without DNA. Then I had washed the unattached cells away from the beads, taking samples for plating at every step so I could track the progress of the three washes. I added DNase I to the beads after the last wash, so I'd be counting the total number of attached cells rather than the number of beads that had cells on them. Then I plated all the samples, and later counted the colonies.
There were ten times more cells on the DNA-coated beads than on the control beads (about 10^5 vs 10^4). This is good, but because I used about 10^8 beads, it means that, at best, only about one bead in 1000 had a cell attached. I hadn't measured how much DNA was on these beads, lazily hoping that they had as much DNA as the similarly treated polystyrene beads. Now I need to repeat the experiment with better-characterized beads.
There were ten times more cells on the DNA-coated beads than on the control beads (about 10^5 vs 10^4). This is good, but because I used about 10^8 beads, it means that, at best, only about one bead in 1000 had a cell attached. I hadn't measured how much DNA was on these beads, lazily hoping that they had as much DNA as the similarly treated polystyrene beads. Now I need to repeat the experiment with better-characterized beads.
Progress without tweezers
Yesterday I went to the biophysics lab, both to attend their weekly group meeting (another very good practice talk by a grad student) and to finally test whether I can see the cells under their microscopes (yes I could).
Their ordinary scope is a Zeiss that has darkfield but not phase contrast. We couldn't get the darkfield set up properly. Having now read up a bit on how to set up darkfield, I realize this was probably because we didn't have the 'stop' disk for the 20X objective, and didn't have a special darkfield condenser for the 100X objective. But the RA showed me how to get OK contrast by shutting down the condenser diaphragm to a pinhole, and with this I was able to see not only B. subtilis cells but the much smaller H. influenzae cells. This let me check that the cells were attached to the coverslip of the chamber before I put the slide with the chamber into the tweezers apparatus.
The laser component of the apparatus hadn't been realigned yet, but that didn't matter as I just wanted to use the visible-light component to find out whether I could see the cells. If I hadn't been able to see the cells I might have had to abandon the whole project. I guess I wasn't proceeding entirely out of optimistic ignorance, because I did know that the biophysics grad student who started this project a few years ago had been able to see H. influenzae cells in the apparatus he was using. I couldn't get the cells into very clear focus under the 70X water-immersion lens of the tweezers apparatus, I think because the optics were a bit out of alignment. The cells also weren't very conspicuous because they aren't very refractile (unlike the polystyrene beads). But I could easily see that they were there.
From now on I'm going to mark the center of the coverslip with an X using a fine-point Sharpie before I assemble each chamber. This X will give me a easy target to focus on, right in the plane of the attached cells. I did this today, with some more careful tests of cell attachment, and it worked very well.
Tomorrow I'm going to make more competent B. subtilis cells, and continue testing cell attachment issues, especially how to prevent the beads from sticking to the cover slip.
Their ordinary scope is a Zeiss that has darkfield but not phase contrast. We couldn't get the darkfield set up properly. Having now read up a bit on how to set up darkfield, I realize this was probably because we didn't have the 'stop' disk for the 20X objective, and didn't have a special darkfield condenser for the 100X objective. But the RA showed me how to get OK contrast by shutting down the condenser diaphragm to a pinhole, and with this I was able to see not only B. subtilis cells but the much smaller H. influenzae cells. This let me check that the cells were attached to the coverslip of the chamber before I put the slide with the chamber into the tweezers apparatus.
The laser component of the apparatus hadn't been realigned yet, but that didn't matter as I just wanted to use the visible-light component to find out whether I could see the cells. If I hadn't been able to see the cells I might have had to abandon the whole project. I guess I wasn't proceeding entirely out of optimistic ignorance, because I did know that the biophysics grad student who started this project a few years ago had been able to see H. influenzae cells in the apparatus he was using. I couldn't get the cells into very clear focus under the 70X water-immersion lens of the tweezers apparatus, I think because the optics were a bit out of alignment. The cells also weren't very conspicuous because they aren't very refractile (unlike the polystyrene beads). But I could easily see that they were there.
From now on I'm going to mark the center of the coverslip with an X using a fine-point Sharpie before I assemble each chamber. This X will give me a easy target to focus on, right in the plane of the attached cells. I did this today, with some more careful tests of cell attachment, and it worked very well.
Tomorrow I'm going to make more competent B. subtilis cells, and continue testing cell attachment issues, especially how to prevent the beads from sticking to the cover slip.
Planning: Do competent cells bind to DNA-coated magnetic beads?
I want to test whether competent cells will bind to DNA that's attached to the surface of beads. I'll use magnetic beads for this, rather that the polystyrene beads I'll use with the tweezers, as I can easily separate them from cells. In principle I just need to mix cells and beads, pull out the beads, and plate them to count the number of beads that have attached cells But the details need some thought.
In fact I already gave this experiment some thought, but I was derailed by finding that the beads formed big clumps when mixed with competent cells and by running out of competent cells. I found that the BSA-coated ('MyOne') beads didn't clump so I'll use these, even though they're not the ones recommended for DNA work. And I've made new cells. But I didn't think much about some other details.
One is timing. I think I should allow only a short time for the cells to interact with the beads. This is more critical for B. subtilis, which can cut bound DNA and so might break lose of the beads. So I think I'll give the H. influenzae cells 5 min to find the DNA, and the B. subtilis cells only 1 min. Then I'll chill the tubes on ice before exposing them to the magnet.
Another issue is the absolute and relative concentrations of cells and beads. I think I should use similar numbers of cells and beads. My frozen competent H. influenzae cells are at about 10^9 cfu/ml. The frozen competent B. subtilis cells are more dilute, ~2x10^8/ml. The MyOne bead stock is even more concentrated, ~10^10 beads/ml, but I only have 1.0 ml and will use only a little bit of this.
So let's say I coat 10 µl of beads with DNA, and mix them with 100 µl of H. influenzae cells, and after incubation I dilute this to 1 ml with cold wash solution. Should I plate both the bead fraction and the supernatant ('side-natant'? call it the SN) fraction, and how many washes should I do? If all the cells are competent and all the beads are DNA-coated, then all the cells might bind to the beads. If most of the cells aren't competent, or if cells are inefficient at finding and binding to the beads, most of the cells will be in the SN, and plating it will probably not be sensitive enough to detect the small decrease.
Let's conservatively assume that only 10^6 cells bind to the 10^8 beads, and that about 5% of each SN is carried over with the beads in each wash. The first SN will still contain ~10^7 cells/ml, and the bead fraction will have 1.5x10^6 cells (~10^6 attached and ~5x10^5 free). I'll resuspend the beads in 1 ml, mix gently, and collect the bead fraction again. The second SN will contain about 5x10^5 cells/ml, and the beads will contain about 10^6 (~10^6 attached and 2.5x10^4 free). If I wash these beads a third time, the SN will contain about 2.5x10^4 cells/ml and the bead fraction will still contain about 10^6 cells.
So I could get away with doing only two washes, but doing a third would give more confidence that the cells are really attached. This assumes that the cells stay attached to the beads through the washings - this will require mixing them quite gently and keeping them cold, and hoping.
One other complication is that counting the cells on the beads uses up beads. But I don't want to count more than ~100 colonies, so I could just take 10 µl of each 1 ml resuspended bead fraction for dilution and plating.
In fact I already gave this experiment some thought, but I was derailed by finding that the beads formed big clumps when mixed with competent cells and by running out of competent cells. I found that the BSA-coated ('MyOne') beads didn't clump so I'll use these, even though they're not the ones recommended for DNA work. And I've made new cells. But I didn't think much about some other details.
One is timing. I think I should allow only a short time for the cells to interact with the beads. This is more critical for B. subtilis, which can cut bound DNA and so might break lose of the beads. So I think I'll give the H. influenzae cells 5 min to find the DNA, and the B. subtilis cells only 1 min. Then I'll chill the tubes on ice before exposing them to the magnet.
Another issue is the absolute and relative concentrations of cells and beads. I think I should use similar numbers of cells and beads. My frozen competent H. influenzae cells are at about 10^9 cfu/ml. The frozen competent B. subtilis cells are more dilute, ~2x10^8/ml. The MyOne bead stock is even more concentrated, ~10^10 beads/ml, but I only have 1.0 ml and will use only a little bit of this.
So let's say I coat 10 µl of beads with DNA, and mix them with 100 µl of H. influenzae cells, and after incubation I dilute this to 1 ml with cold wash solution. Should I plate both the bead fraction and the supernatant ('side-natant'? call it the SN) fraction, and how many washes should I do? If all the cells are competent and all the beads are DNA-coated, then all the cells might bind to the beads. If most of the cells aren't competent, or if cells are inefficient at finding and binding to the beads, most of the cells will be in the SN, and plating it will probably not be sensitive enough to detect the small decrease.
Let's conservatively assume that only 10^6 cells bind to the 10^8 beads, and that about 5% of each SN is carried over with the beads in each wash. The first SN will still contain ~10^7 cells/ml, and the bead fraction will have 1.5x10^6 cells (~10^6 attached and ~5x10^5 free). I'll resuspend the beads in 1 ml, mix gently, and collect the bead fraction again. The second SN will contain about 5x10^5 cells/ml, and the beads will contain about 10^6 (~10^6 attached and 2.5x10^4 free). If I wash these beads a third time, the SN will contain about 2.5x10^4 cells/ml and the bead fraction will still contain about 10^6 cells.
So I could get away with doing only two washes, but doing a third would give more confidence that the cells are really attached. This assumes that the cells stay attached to the beads through the washings - this will require mixing them quite gently and keeping them cold, and hoping.
One other complication is that counting the cells on the beads uses up beads. But I don't want to count more than ~100 colonies, so I could just take 10 µl of each 1 ml resuspended bead fraction for dilution and plating.
Cells attached to glass slides are not only viable but transformable
I bound competent cells (H. influenzae and B. subtilis, separately) onto cover slips in chambers, and then washed in medium containing genetically marked DNA. After 15 minutes at 37 °C I washed in some medium with DNase I and then medium containing 0.5% low-melt agarose, with and without either nalidixic acid (to select the H. influenzae nalR allele) or tryptophan (to select the B. subtilis trp+ allele). Then I sealed the chambers' ends with wax from a candle (clumsily) and incubated them overnight.
The results were a bit messy. The H. influenzae cells started out quite dense (1000-5000 per 40X field of view) and without nalidixic acid they grew into nice microcolonies. With nalidixic acid they instead formed filaments (i.e. grew without dividing, and then stopped growing). At the edges of the chamber some large and well-defined microcolonies were present, but these were absent from the central part of the chamber. So transformation is clearly happening, but I can't estimate a frequency. The distribution of the NalR microcolonies suggests that oxygen might affect resistance, but I don't know enough about how the drug acts (a gyrase inhibitor) to guess why that would be the case.
The B. subtilis results were worse. The chambers with and without tryptophan had similar numbers of cells. Both had substantially more cells than had originally been attached, so there must have been some growth. But there were no obvious microcolonies. I might have made an error with the medium. But many of the cells were moving around, so I suspect that I also need to use a higher concentration of low-melt agarose to block their strong motility.
On a separate topic, a reader suggested using polyethylene glycol (PEG) to block the poly-L-lysine coating and prevent beads from sticking to it once the cells had been bound. After checking that PEG isn't toxic to cells, I tried washing my chambers with a 1% solution of the PEG type we had on the shelf (PEG 3350, which I think is a moderate chain length for PEG), washing it out, and then washing in some beads and washing them out after 10 minutes. Result - no significant difference between PEG and no PEG in the numbers of beads bound to the coverslip. I could try a higher concentration of PEG, but maybe the problem is washing out the PEG before adding the beads. I'll try adding PEG to the beads as well as to the washing solution. If this does prevent beads sticking, I'll next need to test whether PEG inhibits transformation.
While I was at it I also tested whether the 16% glycerol that's mixed with the frozen competent cells inhibits cell binding to the coverslips. I had been conscientiously pelleting the thawed cells and resuspending them in glycerol-free medium before adding them to the chambers, but now I know that cells bind just fine in the presence of glycerol. The glycerol is subsequently washed out of the chamber along with the non-attached cells, so I won't bother with the centrifuging step any more.
And I tested alternatives to sealing the chamber ends with wax, which is difficult to apply and tends to form big lumps rather than a smooth layer. (I don't like the risk of getting wax on the microscope lens.) First I tried a better way of applying the melted wax - rather than using a glass Pasteur pipette with a rubber bulb, I tried using an old Pipetman p200 with a snipped-off tip. This gave better control, but the wax still formed a big lump when it met the glass. Parafilm didn't work - a film of liquid quickly seeped under the parafilm. Melting the parafilm with a heated spatula didn't help. I also tried using paint from a paint-Sharpie; this only sort-of worked, perhaps because it's a water-based paint. (But it was cool to look at the paint droplets under the microscope.)
The results were a bit messy. The H. influenzae cells started out quite dense (1000-5000 per 40X field of view) and without nalidixic acid they grew into nice microcolonies. With nalidixic acid they instead formed filaments (i.e. grew without dividing, and then stopped growing). At the edges of the chamber some large and well-defined microcolonies were present, but these were absent from the central part of the chamber. So transformation is clearly happening, but I can't estimate a frequency. The distribution of the NalR microcolonies suggests that oxygen might affect resistance, but I don't know enough about how the drug acts (a gyrase inhibitor) to guess why that would be the case.
The B. subtilis results were worse. The chambers with and without tryptophan had similar numbers of cells. Both had substantially more cells than had originally been attached, so there must have been some growth. But there were no obvious microcolonies. I might have made an error with the medium. But many of the cells were moving around, so I suspect that I also need to use a higher concentration of low-melt agarose to block their strong motility.
On a separate topic, a reader suggested using polyethylene glycol (PEG) to block the poly-L-lysine coating and prevent beads from sticking to it once the cells had been bound. After checking that PEG isn't toxic to cells, I tried washing my chambers with a 1% solution of the PEG type we had on the shelf (PEG 3350, which I think is a moderate chain length for PEG), washing it out, and then washing in some beads and washing them out after 10 minutes. Result - no significant difference between PEG and no PEG in the numbers of beads bound to the coverslip. I could try a higher concentration of PEG, but maybe the problem is washing out the PEG before adding the beads. I'll try adding PEG to the beads as well as to the washing solution. If this does prevent beads sticking, I'll next need to test whether PEG inhibits transformation.
While I was at it I also tested whether the 16% glycerol that's mixed with the frozen competent cells inhibits cell binding to the coverslips. I had been conscientiously pelleting the thawed cells and resuspending them in glycerol-free medium before adding them to the chambers, but now I know that cells bind just fine in the presence of glycerol. The glycerol is subsequently washed out of the chamber along with the non-attached cells, so I won't bother with the centrifuging step any more.
And I tested alternatives to sealing the chamber ends with wax, which is difficult to apply and tends to form big lumps rather than a smooth layer. (I don't like the risk of getting wax on the microscope lens.) First I tried a better way of applying the melted wax - rather than using a glass Pasteur pipette with a rubber bulb, I tried using an old Pipetman p200 with a snipped-off tip. This gave better control, but the wax still formed a big lump when it met the glass. Parafilm didn't work - a film of liquid quickly seeped under the parafilm. Melting the parafilm with a heated spatula didn't help. I also tried using paint from a paint-Sharpie; this only sort-of worked, perhaps because it's a water-based paint. (But it was cool to look at the paint droplets under the microscope.)
Tweezers-prep progress
1. I tested cell adhesion to the coverslips the RA had cleaned for me - it wasn't dramatically better or more consistent than either uncleaned or acid-washed coverslips (after all had been coated with poly-L-lysine using his spreading method. Overall plenty of cells are adhering, given that I'll only need a small number of cells adhered for each chamber. I think part of the trick was inverting the slides for 10 minutes before washing away the nonadherent cells, so the cells settle on the coverslip. I'm also giving the slides a few brisk taps during the washing, so that loose cells detach and wash away.
2. Incubating the chambers with concentrated BSA protein didn't significantly reduce the number of H. influenzae or B. subtilis cells that stick to the poly-L-lysine coated coverslips. Nor did it prevent the polystyrene beads from sticking to the coverslips. A Google search for treatments that might block/inhibit the poly-L-lysine surface revealed only that people are coupling poly-L-lysine and polyethylene glycol or other materials into 'block' copolymers, and testing the ability of poly-L-lysine to 'inhibit' cell growth and various enzymatic reactions. I didn't test BHI directly, but cells growing in BHI did adhere to the coverslips so I don't think it's much of an inhibitor either. I don't know how big a problem the unwanted sticking is going to be, but I'd welcome suggestions for other possible blocking agents to test. Just for fun I'm going to test milk. Not reconstituted powdered skim milk, but whatever's in the food fridge (2% milk, I think).
3. I made my big batch of competent cells and froze 34 tubes (0.5 ml), as well as 14 tubes of log-phase cells. But I haven't yet tested how competent they are.
4. Several tests showed that the H. influenzae cells that are stuck to the coverslips are otherwise healthy. In the first test I filled the chambers with sBHI plus low-melt agarose and incubated them overnight. I didn't seal the ends of the chambers but put the slides in a humidified box, but the chambers dried out completely overnight. But I could see that there had been lots of cell doublings before the medium dried up. So next I tested sealing the chamber ends with nail polish. The tweezers lab people use wax from a candle, because they worry that the nail polish might be toxic, but I didn't have a candle. The nail polish is indeed toxic (probably because acetone is quite water-soluble); the cells close to the ends of the chamber didn't divide at all. But the cells in the middle part of the chamber grew well, producing nice tight microcolonies by the end of the day. I brought in a candle, and today I'll see if I can improve my skill at applying melted wax to chamber ends.
5. I think today I'll also test whether the cells on the coverslips can be transformed, by washing in MAP7 DNA, and then DNaseI and then sBHI agarose with added novobiocin. Provided I have thousands of cells stuck to the coverslip, I should be able to find rare transformant colonies. I'll try this with B. subtilis too, selecting for Trp+.
2. Incubating the chambers with concentrated BSA protein didn't significantly reduce the number of H. influenzae or B. subtilis cells that stick to the poly-L-lysine coated coverslips. Nor did it prevent the polystyrene beads from sticking to the coverslips. A Google search for treatments that might block/inhibit the poly-L-lysine surface revealed only that people are coupling poly-L-lysine and polyethylene glycol or other materials into 'block' copolymers, and testing the ability of poly-L-lysine to 'inhibit' cell growth and various enzymatic reactions. I didn't test BHI directly, but cells growing in BHI did adhere to the coverslips so I don't think it's much of an inhibitor either. I don't know how big a problem the unwanted sticking is going to be, but I'd welcome suggestions for other possible blocking agents to test. Just for fun I'm going to test milk. Not reconstituted powdered skim milk, but whatever's in the food fridge (2% milk, I think).
3. I made my big batch of competent cells and froze 34 tubes (0.5 ml), as well as 14 tubes of log-phase cells. But I haven't yet tested how competent they are.
4. Several tests showed that the H. influenzae cells that are stuck to the coverslips are otherwise healthy. In the first test I filled the chambers with sBHI plus low-melt agarose and incubated them overnight. I didn't seal the ends of the chambers but put the slides in a humidified box, but the chambers dried out completely overnight. But I could see that there had been lots of cell doublings before the medium dried up. So next I tested sealing the chamber ends with nail polish. The tweezers lab people use wax from a candle, because they worry that the nail polish might be toxic, but I didn't have a candle. The nail polish is indeed toxic (probably because acetone is quite water-soluble); the cells close to the ends of the chamber didn't divide at all. But the cells in the middle part of the chamber grew well, producing nice tight microcolonies by the end of the day. I brought in a candle, and today I'll see if I can improve my skill at applying melted wax to chamber ends.
5. I think today I'll also test whether the cells on the coverslips can be transformed, by washing in MAP7 DNA, and then DNaseI and then sBHI agarose with added novobiocin. Provided I have thousands of cells stuck to the coverslip, I should be able to find rare transformant colonies. I'll try this with B. subtilis too, selecting for Trp+.
Uptake sequence manuscript done, back to the tweezers experiments
Yesterday I finished revising all the components of our manuscript on uptake sequence evolution, and sent them to the co-author who's handling the resubmission. it was already a good manuscript, but now it's very very good, so if the editor and reviewers don't like it we'll just sent it somewhere else.
I've been working on the components of the optical tweezers experiments, with modest progress. The tweezers apparatus still had problems when I was there the other day (it wasn't aligned properly?), but I have lots of other parts to work on. Here's a list:
I've been working on the components of the optical tweezers experiments, with modest progress. The tweezers apparatus still had problems when I was there the other day (it wasn't aligned properly?), but I have lots of other parts to work on. Here's a list:
- The RA of the tweezers lab showed me a way to pre-clean coverslips, and I need to test the ones he prepared to see if they give more consistent results than the ones I've been using. (Coat his and mine with poly-L-lysine, make into chambers, test binding of competent cells.)
- I need to test whether washing treated coverslips with culture medium (BHI) or protein (BSA) blocks cell and bead binding, because once I have cells bound to the surface I don't want beads to stick to it.
- I need to make a big batch of competent cells and freeze them in 0.5 ml aliquots, because the cells I made before were contaminated. (The contaminants were flagellated cells that otherwise looked just like H. influenzae - they were the spinning wiggling cells I described in my last post.)
- I need to test whether cells that have stuck to a coverslip are still able to grow. To do this, my plan is to replace the medium in the chamber with medium containing low-melt agarose, and once this has set incubate the chamber at 37 °C for a few hours and then examine the cells under the microscope, looking for clusters of cells (microcolonies). As a control I'll first try this with cell-free chambers and agarose medium that already contains growing cells.
- For today I'm just going to work with H. influenzae, but I should also test these factors with B. subtilis cells.
Haemophilus doesn't have flagellae!
I'm still testing treatments to get cells to stick to coverslips (with more success):
I used more poly-L-lysine this time, but the important thing I realized was that I need to tap the slides briskly to loosen the cells that are just sitting on the surface but not attached,a nd to rinse a lot of medium through the chamber. This let me see that there were usually substantially more B. subtilis cells attached to the coverslip than to the glass slide.
So then I tested whether my competent H. influenzae cells would also bind to the coverslips. They did, better than the B. subtilis cells. I could see that many of the cells were only attached at one end, with the other end moving in the medium. But what was surprising was how vigorously the free parts of the cells were moving around. Not just bouncing back and forth (can Brownian motion be that vigorous?), but sometimes spinning around almost like cells tethered by their flagella (see this movie, which I think is Salmonella but might be E. coli).
But H. influenzae definitely doesn't have flagella. So I'm plating the cells I have onto LB and BHI plates with and without hemin and NAD, to check that some flagellated imposter hasn't snuck in to my culture. (It would have to make colonies that look just like H. influenzae colonies, and be sensitive to novobiocin...)
I used more poly-L-lysine this time, but the important thing I realized was that I need to tap the slides briskly to loosen the cells that are just sitting on the surface but not attached,a nd to rinse a lot of medium through the chamber. This let me see that there were usually substantially more B. subtilis cells attached to the coverslip than to the glass slide.
So then I tested whether my competent H. influenzae cells would also bind to the coverslips. They did, better than the B. subtilis cells. I could see that many of the cells were only attached at one end, with the other end moving in the medium. But what was surprising was how vigorously the free parts of the cells were moving around. Not just bouncing back and forth (can Brownian motion be that vigorous?), but sometimes spinning around almost like cells tethered by their flagella (see this movie, which I think is Salmonella but might be E. coli).
But H. influenzae definitely doesn't have flagella. So I'm plating the cells I have onto LB and BHI plates with and without hemin and NAD, to check that some flagellated imposter hasn't snuck in to my culture. (It would have to make colonies that look just like H. influenzae colonies, and be sensitive to novobiocin...)
Sticking cells to coverslips (one step forward, two steps back?)
I coated some coverslips with poly-L-lysine, using the rub-until-dry method I was shown. I then assembled the coverslips into 'chambers' like those described here, and tested whether competent B. subtilis cells would stick to them.
I wanted to use chambers for these tests because that's what I'll be using with the optical tweezers. Another benefit turned out to be that I could compare how well cells stuck to the coated coverslip surfaces to how well they stuck to the untreated surfaces of the glass slides that form the bottoms of the chambers.
I let the cells sit in the chamber for 5-10 minutes at room temperature, and then washed them out by flowing about 10 volumes of medium through the chamber, introducing it in tiny drops from a pipette tip at one end and absorbing the flow-through with a square of blotting paper at the other end. Then I looked at the chamber under the microscope, comparing the cells on its upper and lower surfaces.
In most tests I saw little or no difference between the treated and untreated surfaces; quite a few cells were stuck on both. The largest volume of poly-L-lysine solution I used did give some patches where many cell stuck, but these were at the edges of the coverslip, where the rub-until-dry treatment hadn't reached well. Cell density (on both surfaces) was also generally higher at the edges of the chambers - I think this may just v=be because the washing is less effective at the edges.
I also tested my DNA-coated beads. They didn't stick any better than the cells did, which is good. But there were lots of clumps of beads - perhaps I didn't vortex them well.
Changes/improvements:
In a previous experiment I had better results with coverslips that had been presoaked in acid alcohol, so I'll try this again.
I'll try higher concentrations of poly-L-lysine.
I'll try using more dilute cells.
I'll try H. influenzae cells as well as B. subtilis cells, because their surfaces have very different chemistries.
I'll try incubating the chambers+cells upside down before washing the cells out, so the cells will settle on the coverslip rather than the slide. Though, for the tweezers experiments, it doesn't matter which surface the cells are stuck on.
I'll try marking a reference spot on the coverslip, so I can track whether cells present after the first wash are removed by the second one.
I'll reread the papers that did tweezers studies of competent cells, and email their authors for advice.
I wanted to use chambers for these tests because that's what I'll be using with the optical tweezers. Another benefit turned out to be that I could compare how well cells stuck to the coated coverslip surfaces to how well they stuck to the untreated surfaces of the glass slides that form the bottoms of the chambers.
I let the cells sit in the chamber for 5-10 minutes at room temperature, and then washed them out by flowing about 10 volumes of medium through the chamber, introducing it in tiny drops from a pipette tip at one end and absorbing the flow-through with a square of blotting paper at the other end. Then I looked at the chamber under the microscope, comparing the cells on its upper and lower surfaces.
In most tests I saw little or no difference between the treated and untreated surfaces; quite a few cells were stuck on both. The largest volume of poly-L-lysine solution I used did give some patches where many cell stuck, but these were at the edges of the coverslip, where the rub-until-dry treatment hadn't reached well. Cell density (on both surfaces) was also generally higher at the edges of the chambers - I think this may just v=be because the washing is less effective at the edges.
I also tested my DNA-coated beads. They didn't stick any better than the cells did, which is good. But there were lots of clumps of beads - perhaps I didn't vortex them well.
Changes/improvements:
In a previous experiment I had better results with coverslips that had been presoaked in acid alcohol, so I'll try this again.
I'll try higher concentrations of poly-L-lysine.
I'll try using more dilute cells.
I'll try H. influenzae cells as well as B. subtilis cells, because their surfaces have very different chemistries.
I'll try incubating the chambers+cells upside down before washing the cells out, so the cells will settle on the coverslip rather than the slide. Though, for the tweezers experiments, it doesn't matter which surface the cells are stuck on.
I'll try marking a reference spot on the coverslip, so I can track whether cells present after the first wash are removed by the second one.
I'll reread the papers that did tweezers studies of competent cells, and email their authors for advice.
Transformation by DNA on beads
Yesterday I incubated some of my competent H. influenzae cells with DNA carrying a novobiocin-resistance gene. One tube got the normal prep of chromosomal NovR DNA. Two others got DNA that had been cut with one of two restriction enzymes (either EcoRI, average fragment size 6 kb or XhoI, average fragment size 12 kb) and then had biotin incorporated at the ends of the fragments, and three others got streptavidin-coated polystyrene beads with the biotinylated DNA bound to them.
I had calculated that the first batch of DNA+beads had about 250 ng of DNA per ml, so I used the same volumes of beads for the others, and diluted the non-bead DNAs to 250 ng/ml before using the same volumes. I incubated the DNA plus cells for 15 minutes and then plated the cells on Nov agar, and (diluted) on plain agar. Today I counted the colonies and calculated the transformation frequencies.
Point 1. Cutting the DNA with XhoI reduces its transforming ability by 4-fold, but cutting with EcoRI reduces it by about 100-fold. So I checked where these two enzymes cut relative to the NovR (gyrB) gene. EcoRI cuts inside the gene, but XhoI only cuts at sites 1 kb and 3 kb on either side of it.
Point 2. DNA attached to beads transforms! This wasn't a sure thing, for lots of reasons.
Point 3. DNA attached to beads transforms 10-30-fold worse than the same amount of free DNA. This is not surprising because most of the DNA on the beads will be inaccessible because it's tangled up with or behind other DNA fragments.
Now I'll celebrate by spreading some poly-L-lysine on some cover slips, using the new method I was taught yesterday, so tomorrow I can test whether cells bind to these and whether they stay alive after binding.
I had calculated that the first batch of DNA+beads had about 250 ng of DNA per ml, so I used the same volumes of beads for the others, and diluted the non-bead DNAs to 250 ng/ml before using the same volumes. I incubated the DNA plus cells for 15 minutes and then plated the cells on Nov agar, and (diluted) on plain agar. Today I counted the colonies and calculated the transformation frequencies.
Point 2. DNA attached to beads transforms! This wasn't a sure thing, for lots of reasons.
Point 3. DNA attached to beads transforms 10-30-fold worse than the same amount of free DNA. This is not surprising because most of the DNA on the beads will be inaccessible because it's tangled up with or behind other DNA fragments.
Now I'll celebrate by spreading some poly-L-lysine on some cover slips, using the new method I was taught yesterday, so tomorrow I can test whether cells bind to these and whether they stay alive after binding.
Tweezers progress
Yesterday I took my new batches of frozen competent cells (B. subtilis and H. influenzae) across town to the biophysics lab, so I'll have cells there to test without needing to make fresh ones. While I was there I learned a better way to coat cover slips with poly-L-lysine, which I hope will give more reproducible attachment of competent cells. And I spent more time making microscope-slide chambers and learning about the tweezers apparatus (mostly standing by while the expert grad student made adjustments to the optics and electronics).
I also attended a seminar about DNA bending. Short fragments of double-stranded DNA (~100 bp) have been reported to circularize much more efficiently than predicted by their expected persistence length (paper by Cloutier and Widom), and one proposed explanation is the formation of tiny 'bubbles' in the DNA structure – places where several base pairs have separated although the DNA backbones remain intact. Even though such short single-stranded regions are expected to be very transient they can have a big impact on the probability that the ends of the DNA will meet, allowing DNA ligase to join them and circularize the DNA.
I also attended a seminar about DNA bending. Short fragments of double-stranded DNA (~100 bp) have been reported to circularize much more efficiently than predicted by their expected persistence length (paper by Cloutier and Widom), and one proposed explanation is the formation of tiny 'bubbles' in the DNA structure – places where several base pairs have separated although the DNA backbones remain intact. Even though such short single-stranded regions are expected to be very transient they can have a big impact on the probability that the ends of the DNA will meet, allowing DNA ligase to join them and circularize the DNA.
Blocking with BSA makes a big difference
The streptavidin-coated magnetic beads that hadn't been blocked with BSA aggregated into big clumps when mixed with cells, but the treated beads remained separate. The clumping was independent of whether the beads had been incubated with biotinylated DNA, so I think the clumping was because most cells stuck to more than one bead and most beads stuck to more than one cell.
So then I checked the streptavidin-coated polystyrene beads I'll use for the tweezers experiments. Luckily adding cells didn't make them clump.
Now I'm making fresh batches of competent H. influenzae and B. subtilis to freeze, so I'll have consistent cells to work with.
So then I checked the streptavidin-coated polystyrene beads I'll use for the tweezers experiments. Luckily adding cells didn't make them clump.
Now I'm making fresh batches of competent H. influenzae and B. subtilis to freeze, so I'll have consistent cells to work with.
Do cells bind to DNA stuck on beads?
I need to plan the experiment(s) where I test whether competent cells bind to DNA on magnetic beads.
I have 1 ml each of two types of 1µ and 2.8µ magnetic 'dynabeads'. The beads have been coated with streptavidin, and one batch of each size has then been blocked with BSA, which apparently reduces the surface charge and makes them better for binding protein but not so good for nucleic acids and I think more prone to clumping in high-salt buffers. I don't know how the different surfaces will affect non-specific cell binding, something I want to avoid (Invitrogen recommends including 0.01-0.1% Tween 20 to reduce non-specific binding, where that won't interfere with the assay). And I don't know how the surface properties of these beads compare to the properties of the polystyrene beads I'll be using for the tweezers experiments. (I need to keep reminding myself that I'm only using the magnetic beads to check whether cells will bind to DNA on beads, so I shouldn't waste a lot of time optimizing the assay.)
I can separate the beads from 150 µl of liquid by simply drawing the mixture up in a pipette tip, holding the tip in the nanobead magnet rack for 10 seconds, and slowly expelling the liquid; almost all the beads remain behind on the side of the tip. I can then resuspend the beads by drawing clean buffer up into the tip, away from the magnet.
So: First mix one aliquot of beads with DNA (Invitrogen says to do this in TE + 1 M NaCl). To start I'll just use one size of beads, arbitrarily the 1 µ ones. I know that the stocks contain 10 mg beads per ml, but I don't know how many beads this is. I was going to find that out, by diluting some beads and looking at them under the microscope (we have a hemocytometer). But here's a rough calculation: If the density of the beads is a bit higher than that of water, then a single bead 1 µ in diameter will have the same mass as a 1 µ cube of water, which is 10^-9 mg. So 10 mg of beads/ml is ~10^10 beads/ml. As I did with the polystyrene beads, I'll wash the beads several times to remove unbound DNA. I don't think I need to check that DNA has bound by using the sensitive fluorescence assay - I'll leave that to do if I don't see a difference in cells associated with beads with and without DNA.
Then I'll thaw some frozen competent H. influenzae cells, wash away the glycerol and resuspend them in BHI. Then I'll mix them with DNA-treated beads and control (no DNA) beads, at a concentration of, say, ~ 5x10^8 cells and beads per ml. I'll incubate cells plus beads briefly (1 min at 37°C?) and then wash the beads three times, saving the eluate/supernatant/whatever it should be called. Then I'll plate the beads (several dilutions) and the eluates. I'll also look at the beads under the microscope (maybe do that first).
I have 1 ml each of two types of 1µ and 2.8µ magnetic 'dynabeads'. The beads have been coated with streptavidin, and one batch of each size has then been blocked with BSA, which apparently reduces the surface charge and makes them better for binding protein but not so good for nucleic acids and I think more prone to clumping in high-salt buffers. I don't know how the different surfaces will affect non-specific cell binding, something I want to avoid (Invitrogen recommends including 0.01-0.1% Tween 20 to reduce non-specific binding, where that won't interfere with the assay). And I don't know how the surface properties of these beads compare to the properties of the polystyrene beads I'll be using for the tweezers experiments. (I need to keep reminding myself that I'm only using the magnetic beads to check whether cells will bind to DNA on beads, so I shouldn't waste a lot of time optimizing the assay.)
I can separate the beads from 150 µl of liquid by simply drawing the mixture up in a pipette tip, holding the tip in the nanobead magnet rack for 10 seconds, and slowly expelling the liquid; almost all the beads remain behind on the side of the tip. I can then resuspend the beads by drawing clean buffer up into the tip, away from the magnet.
So: First mix one aliquot of beads with DNA (Invitrogen says to do this in TE + 1 M NaCl). To start I'll just use one size of beads, arbitrarily the 1 µ ones. I know that the stocks contain 10 mg beads per ml, but I don't know how many beads this is. I was going to find that out, by diluting some beads and looking at them under the microscope (we have a hemocytometer). But here's a rough calculation: If the density of the beads is a bit higher than that of water, then a single bead 1 µ in diameter will have the same mass as a 1 µ cube of water, which is 10^-9 mg. So 10 mg of beads/ml is ~10^10 beads/ml. As I did with the polystyrene beads, I'll wash the beads several times to remove unbound DNA. I don't think I need to check that DNA has bound by using the sensitive fluorescence assay - I'll leave that to do if I don't see a difference in cells associated with beads with and without DNA.
Then I'll thaw some frozen competent H. influenzae cells, wash away the glycerol and resuspend them in BHI. Then I'll mix them with DNA-treated beads and control (no DNA) beads, at a concentration of, say, ~ 5x10^8 cells and beads per ml. I'll incubate cells plus beads briefly (1 min at 37°C?) and then wash the beads three times, saving the eluate/supernatant/whatever it should be called. Then I'll plate the beads (several dilutions) and the eluates. I'll also look at the beads under the microscope (maybe do that first).
- Make washing buffer, wash beads, resuspend in BHI, in MIV and in PBS.
- Check under microscope to see if they're clumping.
- Incubate beads with DNA, wash well.Thaw and wash cells. Resuspend in BHI or MIV.
- Mix cells with beads, incubate briefly and wash.
- Assess binding by plating and/or microscopy.
Revisions almost done
Today I finally finished what I very much hope is the last major revision of the uptake-sequence variation manuscript. I've rewritten half of the Introduction and all of the Discussion, using my new non-adversarial, everyone-wins framework. I've pulled together all the data for the supplementary table that serves only to show how quantitative our results are. I've redrawn one figure with new data (same results as the old data). I've gone back through the Responses to Reviewers, changing the responses to match what we've now improved. I've started to draft a cover letter to the Editor.
Then I emailed everything to both my coauthors and to the three people in the lab (postdoc, RA and visiting researcher), asking for only essential changes and polishing of the writing. With luck it will be resubmitted sometime next week, and with even more luck the critical reviewer will find our changes acceptable.
Now I've run out of excuses for not cleaning up my office.
Then I emailed everything to both my coauthors and to the three people in the lab (postdoc, RA and visiting researcher), asking for only essential changes and polishing of the writing. With luck it will be resubmitted sometime next week, and with even more luck the critical reviewer will find our changes acceptable.
Now I've run out of excuses for not cleaning up my office.
Framing the uptake sequence problem (Intro and Discussion)
I think the last post may have been a bit incoherent, but it led me to a new perspective on the problems posed by uptake sequences, one that I think gives a much better frame for the manuscript.
INTRODUCTION (new frame):
(After introducing uptake sequences and uptake biases...)
Why bacteria take up DNA is controversial, and presence in two bacterial groups of DNA uptake sequences and their associated uptake biases pose problems for both major hypotheses.
It's generally assumed that bacteria take up DNA to get benefits from homologous genetic recombination, and that uptake sequences plus biases are a mate-choice adaptation to maximize these benefits by excluding DNAs that are not from close relatives. Although this is intuitively appealing, it is evolutionarily problematic, both because it requires simultaneous evolution of bias in the uptake machinery and genomic sequences matching this bias, and because the genomic sequences can only be 'selected' after the cell carrying them is dead. (There's also the bigger problem that the presumed benefits of recombination are expected to be, on average, very small or nonexistent.)
The alternative hypothesis is that bacteria take up DNA as a source of nutrients (initially nucleotides but also carbon, nitrogen and phosphate), for which the very existence of uptake sequences plus bias is counterintuitive. If DNA in the environment is valued only as nucleotides on a string, all DNAs should be equally useful. Although the sequence bias might play a mechanistic role in DNA uptake (such biases are typical of proteins that bind DNA, even ones whose functions are sequence independent), the high density of the preferred sequences in the genome is perplexing.
The phenomenon of molecular drive may resolve the worst of these problems for both hypotheses, by providing a hypothesis-neutral explanation for uptake sequence abundance.
(Explain molecular drive here.)
If molecular drive is indeed an inevitable consequence of biased DNA uptake and homologous recombination, its action may remove the biggest obstacles for both hypotheses. Below we use a computer simulation of genome evolution to test its requirements and consequences.
METHODS
RESULTS
DISCUSSION (new frame):
Summarize the findings. They are robust.
What's been gained: Proponents of the mate-choice hypothesis now need only explain how natural selection for the benefits of recombination would favour uptake specificity in the genes encoding the uptake machinery - the corresponding uptake sequences will inevitably accumulate in the genome as the specificity strengthens. Proponents of the DNA=food hypothesis need only explain how sequence bias would evolve for mechanistic benefits in DNA uptake; uptake sequences in the genome can be ignored.
The above paragraph is really too adversarial a perspective. It's now much clearer what information is needed to explain uptake specificity. First we need a much more detailed characterization of the real uptake biases (Neisserial and Pasteurellacean). Second we need to know what role uptake bias plays in the process of uptake in each organism. Does a dedicated cell-surface protein pre-screen DNA fragments for uptake sequences before uptake is initiated? Do uptake sequences provide structural flexibility for DNA bending or kinking during initiation of uptake? Do uptake sequences play any role after initiation? Do they affect DNA synapsis or other stages of recombination?
We also need new explicit models of the evolutionary forces that would act on uptake genes and preferred sequences. Can selection for genetic benefits of recombination be strong enough to cause evolution of uptake bias? Or, vice versa, can exclusion of unrelated DNAs reduce the costs of DNA uptake? Analysis of protein sequences in genomes with and without uptake sequences suggests that their evolutionary costs are small, but a theoretical framework for this is lacking. Because the model presented in this paper tracks only a single focal genome, it is not suitable for investigating effects on organismal fitness (whether due to the costs of uptake sequences or to the genetic benefits of recombination).
INTRODUCTION (new frame):
(After introducing uptake sequences and uptake biases...)
Why bacteria take up DNA is controversial, and presence in two bacterial groups of DNA uptake sequences and their associated uptake biases pose problems for both major hypotheses.
It's generally assumed that bacteria take up DNA to get benefits from homologous genetic recombination, and that uptake sequences plus biases are a mate-choice adaptation to maximize these benefits by excluding DNAs that are not from close relatives. Although this is intuitively appealing, it is evolutionarily problematic, both because it requires simultaneous evolution of bias in the uptake machinery and genomic sequences matching this bias, and because the genomic sequences can only be 'selected' after the cell carrying them is dead. (There's also the bigger problem that the presumed benefits of recombination are expected to be, on average, very small or nonexistent.)
The alternative hypothesis is that bacteria take up DNA as a source of nutrients (initially nucleotides but also carbon, nitrogen and phosphate), for which the very existence of uptake sequences plus bias is counterintuitive. If DNA in the environment is valued only as nucleotides on a string, all DNAs should be equally useful. Although the sequence bias might play a mechanistic role in DNA uptake (such biases are typical of proteins that bind DNA, even ones whose functions are sequence independent), the high density of the preferred sequences in the genome is perplexing.
The phenomenon of molecular drive may resolve the worst of these problems for both hypotheses, by providing a hypothesis-neutral explanation for uptake sequence abundance.
(Explain molecular drive here.)
If molecular drive is indeed an inevitable consequence of biased DNA uptake and homologous recombination, its action may remove the biggest obstacles for both hypotheses. Below we use a computer simulation of genome evolution to test its requirements and consequences.
METHODS
RESULTS
DISCUSSION (new frame):
Summarize the findings. They are robust.
What's been gained: Proponents of the mate-choice hypothesis now need only explain how natural selection for the benefits of recombination would favour uptake specificity in the genes encoding the uptake machinery - the corresponding uptake sequences will inevitably accumulate in the genome as the specificity strengthens. Proponents of the DNA=food hypothesis need only explain how sequence bias would evolve for mechanistic benefits in DNA uptake; uptake sequences in the genome can be ignored.
The above paragraph is really too adversarial a perspective. It's now much clearer what information is needed to explain uptake specificity. First we need a much more detailed characterization of the real uptake biases (Neisserial and Pasteurellacean). Second we need to know what role uptake bias plays in the process of uptake in each organism. Does a dedicated cell-surface protein pre-screen DNA fragments for uptake sequences before uptake is initiated? Do uptake sequences provide structural flexibility for DNA bending or kinking during initiation of uptake? Do uptake sequences play any role after initiation? Do they affect DNA synapsis or other stages of recombination?
We also need new explicit models of the evolutionary forces that would act on uptake genes and preferred sequences. Can selection for genetic benefits of recombination be strong enough to cause evolution of uptake bias? Or, vice versa, can exclusion of unrelated DNAs reduce the costs of DNA uptake? Analysis of protein sequences in genomes with and without uptake sequences suggests that their evolutionary costs are small, but a theoretical framework for this is lacking. Because the model presented in this paper tracks only a single focal genome, it is not suitable for investigating effects on organismal fitness (whether due to the costs of uptake sequences or to the genetic benefits of recombination).
Discussion draft
(Here I'm just trying to lay out the philosophical framework of the Discussion.)
Uptake sequences might seem to be a minor trivial problem (only two relatively unimportant bacterial groups), but they have important implications for the evolution of sex. The key question is, are uptake sequences (= sequence-biased DNA uptake plus abundant preferred sequences in the genome) evidence of selection for benefits of homologous recombination?
Start with recap of what needs to be explained. It's difficult to disentangle the hypotheses: Are bias and accumulation the two interdependent components of an adaptation to promote uptake of fragments that can contribute to beneficial recombination, as often assumed? The big problems with this are the weakness of the hypothesized benefits of recombination in current models of the evolution of sex, and the difficulty of selecting for sequences that only act after the owner is dead. Alternatively, each component (uptake bias and uptake sequences in the genome) could have a separate explanation; for example, uptake sequences might have a cellular function, and sequence bias might play a mechanistic role in uptake.
But an acceptable model for bacterial uptake sequences must explain the very strong correspondence between the sequences that are preferred by the uptake machinery and those that are overrepresented in the genome. Given an abundant sequence with a cellular function, might bias favouring it evolve because of genetic benefits of homologous recombination (mate choice), or because of mechanistic benefits of evolving high-affinity DNA-binding proteins specializing in a commonly available sequence? Conversely, given an intrinsically biased uptake machinery due to mechanistic constraints or the need for high-affinity DNA binding, might the preferred sequences accumulate in the genome regardless of recombination benefits?
This last is the simplest hypothesis to test. Our results show accumulation of uptake sequences like those in real genomes, provided only that DNA uptake is strongly biased and homologous recombination sometimes occurs. The results don't require that the homologous recombination provide any genetic benefits. However, the effects we have found will act regardless of whether the benefit to the cell comes from the DNA's nucleotides or its genetic information. They also don't need selection for homologous recombination, as that occurs anyway in repair-capable cells
So now we have shown that, provided DNA is taken up and bias exists, uptake sequences will accumulate. If DNA uptake is selected because it provides food, then uptake sequences have not evolved as markers of sequence homology, and we expect to find that uptake is biased for non-genetic reasons. If DNA is instead taken up for its genetic information, we need to study how the accumulation of uptake sequences affects the genetic benefits, and how the benefits can affect the evolution of the gene causing the bias..
Our finding that uptake sequences accumulate without selection removes one of the problems with the uptake-bias for sex model (that of needing to simultaneously select for bias and uptake sequences. Instead now just need to test whether uptake bias will evolve (gradually) because of benefits of homologous recombination, with uptake sequence accumulation following passively along. Our model of uptake sequence evolution is not designed to evaluate this because it is not population-based (it tracks only a single focal genome). (Can I mention here my new modeling of the effect of selection at one position?)
An alternative approach is to investigate the role bias plays in the uptake mechanism. Finding that bias is created by a mechanism-independent protein that pre-screens sequences for uptake sequences would support the hypothesis that bias exists to promote homologous recombination. Conversely, finding that bias makes a mechanistic contribution to the process of uptake would be consistent with non-genetic functions of DNA uptake. Our present focus is on properly characterizing the true uptake bias of the H. influenzae uptake system, identifying the gene or genes responsible for the bias, and finding out the role of sequence biases in the mechanism of uptake
Uptake sequences might seem to be a minor trivial problem (only two relatively unimportant bacterial groups), but they have important implications for the evolution of sex. The key question is, are uptake sequences (= sequence-biased DNA uptake plus abundant preferred sequences in the genome) evidence of selection for benefits of homologous recombination?
Start with recap of what needs to be explained. It's difficult to disentangle the hypotheses: Are bias and accumulation the two interdependent components of an adaptation to promote uptake of fragments that can contribute to beneficial recombination, as often assumed? The big problems with this are the weakness of the hypothesized benefits of recombination in current models of the evolution of sex, and the difficulty of selecting for sequences that only act after the owner is dead. Alternatively, each component (uptake bias and uptake sequences in the genome) could have a separate explanation; for example, uptake sequences might have a cellular function, and sequence bias might play a mechanistic role in uptake.
But an acceptable model for bacterial uptake sequences must explain the very strong correspondence between the sequences that are preferred by the uptake machinery and those that are overrepresented in the genome. Given an abundant sequence with a cellular function, might bias favouring it evolve because of genetic benefits of homologous recombination (mate choice), or because of mechanistic benefits of evolving high-affinity DNA-binding proteins specializing in a commonly available sequence? Conversely, given an intrinsically biased uptake machinery due to mechanistic constraints or the need for high-affinity DNA binding, might the preferred sequences accumulate in the genome regardless of recombination benefits?
This last is the simplest hypothesis to test. Our results show accumulation of uptake sequences like those in real genomes, provided only that DNA uptake is strongly biased and homologous recombination sometimes occurs. The results don't require that the homologous recombination provide any genetic benefits. However, the effects we have found will act regardless of whether the benefit to the cell comes from the DNA's nucleotides or its genetic information. They also don't need selection for homologous recombination, as that occurs anyway in repair-capable cells
So now we have shown that, provided DNA is taken up and bias exists, uptake sequences will accumulate. If DNA uptake is selected because it provides food, then uptake sequences have not evolved as markers of sequence homology, and we expect to find that uptake is biased for non-genetic reasons. If DNA is instead taken up for its genetic information, we need to study how the accumulation of uptake sequences affects the genetic benefits, and how the benefits can affect the evolution of the gene causing the bias..
Our finding that uptake sequences accumulate without selection removes one of the problems with the uptake-bias for sex model (that of needing to simultaneously select for bias and uptake sequences. Instead now just need to test whether uptake bias will evolve (gradually) because of benefits of homologous recombination, with uptake sequence accumulation following passively along. Our model of uptake sequence evolution is not designed to evaluate this because it is not population-based (it tracks only a single focal genome). (Can I mention here my new modeling of the effect of selection at one position?)
An alternative approach is to investigate the role bias plays in the uptake mechanism. Finding that bias is created by a mechanism-independent protein that pre-screens sequences for uptake sequences would support the hypothesis that bias exists to promote homologous recombination. Conversely, finding that bias makes a mechanistic contribution to the process of uptake would be consistent with non-genetic functions of DNA uptake. Our present focus is on properly characterizing the true uptake bias of the H. influenzae uptake system, identifying the gene or genes responsible for the bias, and finding out the role of sequence biases in the mechanism of uptake
Still (STILL!) working on the uptake sequence variation manuscript
(If the damned thing takes much longer we're going to have to ask the Editor for an extension!)
It's my fault - I'm stalled at fixing up the Discussion.
The problematic reviewer felt that the model didn't make testable predictions, so I wanted to include a brief discussion of how it could be used to evaluate more complex hypotheses about uptake sequence evolution. Unfortunately, a proper test that includes selection will require the model to follow a population of cells or genomes, rather than a single focal genome. Such a model would necessarily be more complicated than ours, and if it was set up like ours the run times might be prohibitively long. (Of course a clever programmer might find ways to streamline it without losing scientific relevance.)
But I thought of a simple test of whether uptake sequences accumulate near to positions that are under fitness selection in the diverging sibs of the focal genome. Modifying the program to do this took only about 12 lines of code, but getting this code to work properly took me most of a day (spent chasing curly brackets and finding out the correct way to use the Perl 'substr' function).
This new version of the program includes selection at position 5000 of the evolving genome. As before, each DNA fragment in the environment is first scored for sequences matching the uptake motif, and the resulting score determines its probability of recombining with the focal genome. But now, if the fragment overlaps position 5000, it is also checked for its base at that position. Fragments with an A at position 5000 keep their original uptake-sequence score for the recombination step, but fragments with the other bases have their scores reduced by a factor of 0.7, 0.4 or 0.1 (for G, C and T respectively). This is intended to simulate poor survival of cells with these bases. If the focal genome has an A it will have a near-normal recombination around position 5000 (except for the 1/100 fragments that carry mutations to less favoured bases), but if it has one of the other bases it will have reduced recombination except for the higher recombination of fragments carrying mutations there). Recurrent mutation at position 5000 (in the focal genome and in the divergent fragments) may create a recurrent benefit of recombination, and if uptake sequences promote beneficial recombination, might select for uptake sequences close to position 5000. On the other hand, if recombination more often brings in harmful mutations, uptake sequences close to the selected position might be selected against.
So I examined the final locations of uptake sequences in genomes from a bunch of runs that started either with 10 kb random-sequence genomes or with 20 kb genomes pre-seeded with uptake sequences (one very close to position 5000). In the random-sequence genomes there were just as many uptake sequences around position 5000 as anywhere else, and in the pre-seeded genomes the uptake sequence at position 4982 was no more and no less stable than any other uptake sequence.
This isn't a very good test, in lots of ways (in fact it's quite awful), but I think it will show the Editor that the model is indeed testable, and that we have made a reasonable effort to satisfy the reviewer. In the manuscript's Discussion I'll describe it in less detail than I have above, and I won't present any data ("Redfield, unpublished"). And I'll explain that a proper test that incorporates selection for beneficial alleles will require a population-based version of the model.
[I also still have to assemble some new data into a replacement for one of the figures, and to go back over the latest changes one more time before sending them to my coauthors on last time.]
It's my fault - I'm stalled at fixing up the Discussion.
The problematic reviewer felt that the model didn't make testable predictions, so I wanted to include a brief discussion of how it could be used to evaluate more complex hypotheses about uptake sequence evolution. Unfortunately, a proper test that includes selection will require the model to follow a population of cells or genomes, rather than a single focal genome. Such a model would necessarily be more complicated than ours, and if it was set up like ours the run times might be prohibitively long. (Of course a clever programmer might find ways to streamline it without losing scientific relevance.)
But I thought of a simple test of whether uptake sequences accumulate near to positions that are under fitness selection in the diverging sibs of the focal genome. Modifying the program to do this took only about 12 lines of code, but getting this code to work properly took me most of a day (spent chasing curly brackets and finding out the correct way to use the Perl 'substr' function).
This new version of the program includes selection at position 5000 of the evolving genome. As before, each DNA fragment in the environment is first scored for sequences matching the uptake motif, and the resulting score determines its probability of recombining with the focal genome. But now, if the fragment overlaps position 5000, it is also checked for its base at that position. Fragments with an A at position 5000 keep their original uptake-sequence score for the recombination step, but fragments with the other bases have their scores reduced by a factor of 0.7, 0.4 or 0.1 (for G, C and T respectively). This is intended to simulate poor survival of cells with these bases. If the focal genome has an A it will have a near-normal recombination around position 5000 (except for the 1/100 fragments that carry mutations to less favoured bases), but if it has one of the other bases it will have reduced recombination except for the higher recombination of fragments carrying mutations there). Recurrent mutation at position 5000 (in the focal genome and in the divergent fragments) may create a recurrent benefit of recombination, and if uptake sequences promote beneficial recombination, might select for uptake sequences close to position 5000. On the other hand, if recombination more often brings in harmful mutations, uptake sequences close to the selected position might be selected against.
So I examined the final locations of uptake sequences in genomes from a bunch of runs that started either with 10 kb random-sequence genomes or with 20 kb genomes pre-seeded with uptake sequences (one very close to position 5000). In the random-sequence genomes there were just as many uptake sequences around position 5000 as anywhere else, and in the pre-seeded genomes the uptake sequence at position 4982 was no more and no less stable than any other uptake sequence.
This isn't a very good test, in lots of ways (in fact it's quite awful), but I think it will show the Editor that the model is indeed testable, and that we have made a reasonable effort to satisfy the reviewer. In the manuscript's Discussion I'll describe it in less detail than I have above, and I won't present any data ("Redfield, unpublished"). And I'll explain that a proper test that incorporates selection for beneficial alleles will require a population-based version of the model.
[I also still have to assemble some new data into a replacement for one of the figures, and to go back over the latest changes one more time before sending them to my coauthors on last time.]
Some comments about my Evo-WIBO talk plan from a reader:
...what I'm really curious about is the sense I get that you feel a phenotype must be some sort of evolutionary goal (i.e., why would we have an a priori expectation that enzymes would evolve to accomplish homologous recombination?) Gender doesn't seem to pop onto the Natural History landscape full blown and ready to be appreciated. So why should HR? I really like the notion that HR might proceed from a DNA replication and repair background.I didn't mean to imply that phenotype is a goal, neither generally or with respect to homologous recombination (HR). But most other microbiologists and molecular biologists have been assuming but not rigorously evaluating) that HR exists because of selection for its sometimes-beneficial consequences.
And is it not possible that natural competence is currently an orphan process that exists for food uptake but was once a piece of a primordial sex process that developed further in other lineages but was cast aside in bacteria? (photosynthesis may have been cast aside in oomycetes in favor of parasitism).That seems backwards to me, because selection for the food benefit is so straightforward and selection for sex so problematic.
To me, HR has to be more beneficial than horizontal gene transfer for a lineage to find it worth the trouble. When organisms are extremely simple the selective disadvantage of maintaining DNA that isn't carrying its weight should lead to its elimination. The notion of an allele implies the existence of a gene - but a gene not in the sense of a capable ORF but in the sense of two or more ORFs in a population that perform the same function in manner that the environment will influence and that selection can act on. If said variant ORFs come to be in the same cell, then HR can go to work on them.Homologous recombination isn't really any 'trouble', to the extent that it happens as an accidental (i.e. unselected) consequence of enzymes selected for their effects on DNA replication and repair and of accidental transfer of DNA fragments by genetic parasites or of DNA uptake for food. And in bacteria there's very little evidence that it ever occurs any other way.
The value of taking a different tack on a problem is prescient. And physics offers a host of tools and a philosophical background that could really help. To me the challenge of the 15 minute presentation is to illustrate how having data that describe the physical process of DNA uptake should allow mathematical model development for the process which then allows development of testable hypotheses. There are "big organism" examples of this approach bearing fruit.I don't think that the phenomenon of natural competence needs mathematical models at all (nor do any other of the phenomena that sometimes lead to to recombination in bacteria). My point for the talk is that many hypotheses can be directly evaluated by more thorough investigation of the phenomena in question.
Defending 'functional design' analysis at Evo-WIBO
In a couple of weeks I'll be giving a short talk at the regional Evo-WIBO meeting. My title is What's an evolutionary biologist doing in a physics lab?. I think I'm going to combine a description of my specific scientific question (the physical properties of DNA uptake by Haemophilus influenzae) with a rehash of the defense of 'functional design' that I made in a post last month. So I might subtitle the talk A defense of functional design analysis.
I'll only have 15 minutes including question time, so I'll need to keep it simple.
I'll only have 15 minutes including question time, so I'll need to keep it simple.
- The simple answer is, I'm measuring the physical properties of DNA uptake by the bacterium Haemophilus influenzae. I'll show you how this is done at the end of my talk, with a nice explanatory animation.
- Why is this of evolutionary interest? Because it's one of the final pieces of the Do bacteria have sex? puzzle.
- Why aren't I using more evolution-style approaches, behaving like a proper evolutionary biologist? How will knowing physical forces answer evolutionary questions? Shouldn't I be using the comparative method? Since these are bacteria, why aren't I doing Rich Lenski-style lab evolution experiments?
- A defense of 'functional design' analysis:
- Understanding 'natural history' (the stamp collecting side of biology?) is fundamental to investigating evolutionary forces. Before we try to explain how natural selection has acted on any phenotype or behaviour, we first need a solid understanding of what the phenotype or behaviour is.
- First a big-organism example: The head-nodding lizards. We can use the usual methods of natural history. What does it do, when does it do it, what are the typical outcomes?
- Next, a bacterial example: For bacteria, we need to use the methods of molecular biology. Consider RecBCD (3 proteins that work together). How was it discovered, what was its function initially thought to be? What was later learned about the phenomenon (not by evolutionary biologists). Molecular biologists often treat both 'functions' as equivalently important. How should evolutionary biologists think about it (consider relative strengths of selective forces).
- Similar history of thinking about nearly all the genes that contribute to homologous recombination in bacteria. The molecular biology isn't my work, but I spell out the implications for evolutionary biologists.
- Main conclusion: Many (and perhaps all) bacteria don't have 'sex'; that is, they don't have any genes that evolved to promote homologous recombination with alleles from other cells of the same or closely related species. Two of the three processes that move DNA from one cell to another are caused by genetic parasites, and the genes responsible for the physical recombination all have important functions in DNA replication and repair. True of E. coli.
- I say 'perhaps all' because the function of one of the three processes that move DNA is still controversial. That's natural competence
How best to test binding of competent cells to DNA on beads?
Now I have lots of biotinylated DNA, and a well-tested procedure for binding DNA fragments to streptavidin-coated styrene beads, I'm ready to test whether competent bacterial cells (B. subtilis or H. influenzae) will bind to the DNA on the beads.
How to do this isn't straightforward. One problem is that the beads are about the same size and density as the cells (B. subtilis cells a bit bigger, H. influenzae cells a bit smaller), so once mixed they can't be easily separated. That means I have no way to wash unbound beads away from cells, or unbound cells away from beads. Another problem is that B. subtilis cells are known to cut DNA fragments as part of the uptake process, and in principle this might terminate uptake. Though maybe not, as the cutting is part of the process that initiates uptake across the inner membrane. H. influenzae cells don't cut DNA.
I could just mix competent cells and DNA-coated beads, both at low densities, on a microscope slide and watch for them sticking to each other. Alternatively, we have some streptavidin-coated paramagnetic beads I could use - this would allow me to pull out the beads and see if cells had stuck to them. But these 50 nm (super-tiny) beads, too small to see individually, so I'd have to plate them to see if there were cells there. We might also have some micron-sized ones; I'll look around.
OK, I found our 'starter kit' of 1 and 2 micron paramagnetic beads. The only problem is, we were too cheap to pay the $150 for the starter version of the magic magnetic rack that holds microfuge tubes against magnets so the beads stick to the side and the liquid can be removed. So I tested various magnets from around the lab, and all of them pulled the rusty-brown beads to the side of the tube in a couple of minutes. Doing this in a way that holds the tube steady so I can remove the liquid...not yet.
I'm away for a few days, but when I get back I may send an email out asking if anyone in the building has a Dynabeads rack I could borrow for a little while.
How to do this isn't straightforward. One problem is that the beads are about the same size and density as the cells (B. subtilis cells a bit bigger, H. influenzae cells a bit smaller), so once mixed they can't be easily separated. That means I have no way to wash unbound beads away from cells, or unbound cells away from beads. Another problem is that B. subtilis cells are known to cut DNA fragments as part of the uptake process, and in principle this might terminate uptake. Though maybe not, as the cutting is part of the process that initiates uptake across the inner membrane. H. influenzae cells don't cut DNA.
I could just mix competent cells and DNA-coated beads, both at low densities, on a microscope slide and watch for them sticking to each other. Alternatively, we have some streptavidin-coated paramagnetic beads I could use - this would allow me to pull out the beads and see if cells had stuck to them. But these 50 nm (super-tiny) beads, too small to see individually, so I'd have to plate them to see if there were cells there. We might also have some micron-sized ones; I'll look around.
OK, I found our 'starter kit' of 1 and 2 micron paramagnetic beads. The only problem is, we were too cheap to pay the $150 for the starter version of the magic magnetic rack that holds microfuge tubes against magnets so the beads stick to the side and the liquid can be removed. So I tested various magnets from around the lab, and all of them pulled the rusty-brown beads to the side of the tube in a couple of minutes. Doing this in a way that holds the tube steady so I can remove the liquid...not yet.
I'm away for a few days, but when I get back I may send an email out asking if anyone in the building has a Dynabeads rack I could borrow for a little while.
How much DNA is on the beads?
The NanoDrop tech support person said that the styrene beads wouldn't hurt the NanoDrop spec, and agreed that light scattering might be a problem. It was, and that combined with the detection threshold of the Nanodrop meant that my measurements didn't give any evidence of DNA on my beads. So today I used the PicoGreen assay to look for DNA on the beads. It's much more sensitive, and not bothered much by light scattering due to the beads.
But first I should describe what my samples were and how I made them. I incubated some 1.26 µ streptavidin-coated beads with a diluted solution of my biotin-labeled DNA, diluted because I didn't want different beads binding to the two ends of a fragment, and I didn't want steric interference by the DNA on the beads. I incubated the beads with the DNA for 30 minutes, gently mixing at 37C on our roller wheel. Then I pelleted the beads, and washed them twice with 1.0 ml of TE, each time rolling the beads plus TE for 10 minutes, and resuspended the washed beads in 100 µl TE (call these Beads1). I also added another aliquot of beads to the DNA solution I'd already used with the first beads, and put these beads through the same incubation, washing and resuspension steps (call these Beads2).
Beads1 and Beads2 had very similar DNA concentrations, about 250 ng/ml. This isn't very much DNA (but see below), but because they're the same I know that the low binding isn't because my biotin-labeling failed. If the labeling had been the problem, then Beads1 would have had little DNA because they had bound up all the labeled DNA in the tube, and Beads2 would have had much less DNA. (I could check this by incubating more beads with the same DNA sample.) Instead the low labeling may be because of the amount of strepavidin on the beads, or its reduced accessibility once bound DNA fragments are getting in the way of other DNA fragments.
So how much DNA is this per bead? Here's a very back-of-the-envelope calculation: The bead concentration in the resuspended Beads1 and Beads2 preps is about 0.1%, assuming that no beads were lost in the washing steps. Let's consider 1 ml of Beads1 (or Beads2), just because it makes the arithmetic clearer. With 0.1% beads, 1 ml of Beads1 solution is about 1 µl of packed beads (and yes, that's about how big the pellets appeared). The beads are about 1.25 µ in diameter, and 1 µl is a cube that's 1000 µ on each side, so 1 µl of packed beads is a cube with about 800 beads per side, or about 5x10^8 beads. At 250 ng/ml, the same ml of Beads1 contains about 250x10^9 kb of DNA (using Rosie's universal constant of 10^18 kb/gram of DNA). The average fragment size of EcoRI-cut H. influenzae DNA is about 6 kb, so this is about 42x10^9 fragments. I conclude that the average bead has about 85 DNA fragments bound to it. That's pretty reasonable for my experiments, so I can go ahead and use these beads and this DNA to test cells binding to DNA on beads.
I also measured the DNA concentrations in the two washes from each aliquot of beads. The first washes had about 20 ng/ml DNA, and the second washes had fluorescences not significantly higher than background, so I know that the signals from Beads1 and Beads2 were due to bound DNA.
One control I didn't do was to make a standard curve using known amounts of DNA mixed with 0.1% beads. I should try this tomorrow. I've also saved the samples I measured, and I'll also try reading them again tomorrow using the high-sensitivity setting of the plate scanner. (Later - I was wrong; there is no high-sensitivity setting.) That's if I can figure out how to do this; the scanner software is very non-intuitive, and so far I've spent most of my time trying to find files I thought I'd saved.
But first I should describe what my samples were and how I made them. I incubated some 1.26 µ streptavidin-coated beads with a diluted solution of my biotin-labeled DNA, diluted because I didn't want different beads binding to the two ends of a fragment, and I didn't want steric interference by the DNA on the beads. I incubated the beads with the DNA for 30 minutes, gently mixing at 37C on our roller wheel. Then I pelleted the beads, and washed them twice with 1.0 ml of TE, each time rolling the beads plus TE for 10 minutes, and resuspended the washed beads in 100 µl TE (call these Beads1). I also added another aliquot of beads to the DNA solution I'd already used with the first beads, and put these beads through the same incubation, washing and resuspension steps (call these Beads2).
Beads1 and Beads2 had very similar DNA concentrations, about 250 ng/ml. This isn't very much DNA (but see below), but because they're the same I know that the low binding isn't because my biotin-labeling failed. If the labeling had been the problem, then Beads1 would have had little DNA because they had bound up all the labeled DNA in the tube, and Beads2 would have had much less DNA. (I could check this by incubating more beads with the same DNA sample.) Instead the low labeling may be because of the amount of strepavidin on the beads, or its reduced accessibility once bound DNA fragments are getting in the way of other DNA fragments.
So how much DNA is this per bead? Here's a very back-of-the-envelope calculation: The bead concentration in the resuspended Beads1 and Beads2 preps is about 0.1%, assuming that no beads were lost in the washing steps. Let's consider 1 ml of Beads1 (or Beads2), just because it makes the arithmetic clearer. With 0.1% beads, 1 ml of Beads1 solution is about 1 µl of packed beads (and yes, that's about how big the pellets appeared). The beads are about 1.25 µ in diameter, and 1 µl is a cube that's 1000 µ on each side, so 1 µl of packed beads is a cube with about 800 beads per side, or about 5x10^8 beads. At 250 ng/ml, the same ml of Beads1 contains about 250x10^9 kb of DNA (using Rosie's universal constant of 10^18 kb/gram of DNA). The average fragment size of EcoRI-cut H. influenzae DNA is about 6 kb, so this is about 42x10^9 fragments. I conclude that the average bead has about 85 DNA fragments bound to it. That's pretty reasonable for my experiments, so I can go ahead and use these beads and this DNA to test cells binding to DNA on beads.
I also measured the DNA concentrations in the two washes from each aliquot of beads. The first washes had about 20 ng/ml DNA, and the second washes had fluorescences not significantly higher than background, so I know that the signals from Beads1 and Beads2 were due to bound DNA.
One control I didn't do was to make a standard curve using known amounts of DNA mixed with 0.1% beads. I should try this tomorrow. I've also saved the samples I measured, and I'll also try reading them again tomorrow using the high-sensitivity setting of the plate scanner. (Later - I was wrong; there is no high-sensitivity setting.) That's if I can figure out how to do this; the scanner software is very non-intuitive, and so far I've spent most of my time trying to find files I thought I'd saved.
At the bench today
Today I did two reactions that labeled the ends of digested chromosomal DNA with biotin (one of EcoRI-digested DNA and one of XhoI-digested DNA). The next step is to clean up the DNA, to get rid of the Klenow polymerase and the EcoRI/XhoI and the unincorporated nucleotides. it's especially important to get rid of ALL of the unincorporated biotin-dUTP, because this will otherwise bind to the streptavidin-coated beads and prevent the biotinylated DNA from binding to them.
Because I want to wash the DNA well to get rid of the biotin-dUTP, a column cleanup is best. We have new cheap cleanup columns from a company called Epoch, to replace the relatively expensive Sigma genelute columns we've been using. (These in turn replaced very expensive columns from Qiagen.) What makes the Epoch columns such good value is that instead of charging for little bottles of salty water like the other companies (their 'secret sauce' buffers), Epoch just provides the recipes so users can make their own buffers.
The RA had already tested the new columns with a PCR cleanup, but I needed to test them with large fragments of chromosomal DNA, because DNA fragments bigger than 10-20 kb tend to stick to this kind of column. Bottom line: both columns release almost all the DNA fragments smaller than 20 kb.
I also wanted to compare the overall recovery of DNA from the columns, especially if they were heavily loaded with a lot of DNA (their stated binding capacities were either 10 µg or 20 µg, depending on which document I read). But my DNA wasn't as concentrated as I thought, so the most DNA I put on a column was probably about 14 µg. Recoveries were good, >80% even with more than 10 µg on the column.
So now I have about 35 µg of biotin-tagged EcoRI-cut chromosomal DNA, and about 25 µg of biotin-tagged XhoI-cut chromosomal DNA. The next step is to measure binding of this DNA to the streptavidin-coated beads. I can do this accurately now that the RA has shown me how to use Picogreen to measure very low DNA concentrations, and I've checked that beads don't interfere with these measurements. I wasn't sure if I could put samples containing beads onto the NanoDrop spec, but I just read their explanation of how it works and I don't see any problem. Maybe I'll email their Customer Service people just to be sure, as the NanoDrop we use belongs to the lab next door.
Because I want to wash the DNA well to get rid of the biotin-dUTP, a column cleanup is best. We have new cheap cleanup columns from a company called Epoch, to replace the relatively expensive Sigma genelute columns we've been using. (These in turn replaced very expensive columns from Qiagen.) What makes the Epoch columns such good value is that instead of charging for little bottles of salty water like the other companies (their 'secret sauce' buffers), Epoch just provides the recipes so users can make their own buffers.
The RA had already tested the new columns with a PCR cleanup, but I needed to test them with large fragments of chromosomal DNA, because DNA fragments bigger than 10-20 kb tend to stick to this kind of column. Bottom line: both columns release almost all the DNA fragments smaller than 20 kb.
I also wanted to compare the overall recovery of DNA from the columns, especially if they were heavily loaded with a lot of DNA (their stated binding capacities were either 10 µg or 20 µg, depending on which document I read). But my DNA wasn't as concentrated as I thought, so the most DNA I put on a column was probably about 14 µg. Recoveries were good, >80% even with more than 10 µg on the column.
So now I have about 35 µg of biotin-tagged EcoRI-cut chromosomal DNA, and about 25 µg of biotin-tagged XhoI-cut chromosomal DNA. The next step is to measure binding of this DNA to the streptavidin-coated beads. I can do this accurately now that the RA has shown me how to use Picogreen to measure very low DNA concentrations, and I've checked that beads don't interfere with these measurements. I wasn't sure if I could put samples containing beads onto the NanoDrop spec, but I just read their explanation of how it works and I don't see any problem. Maybe I'll email their Customer Service people just to be sure, as the NanoDrop we use belongs to the lab next door.
What's up with the manuscript about uptake sequence variation?
We're revising it, though not drastically. One of the reviewers didn't have many concerns, but the other was full of philosophical objections, which we're meeting with calm reason and more analysis.
One bit of data we'll now include is the density of uptake sequences in the equilibrium genomes we discuss. But when I went back to extract this data from the appropriate runs, I found that one run didn't have the data because it hadn't terminated when it was supposed to; there was a typo in the specified termination cycle (2000o0 rather than 200000), so it would have kept running forever if I hadn't stopped it.
And when I went to redo that run without the typo, I discovered that the set of 12 runs it belonged to had all had another error; instead of recombining 1000 fragments each cycle they had only recombined 100. Fixing this won't change the conclusions at all; the runs will just all converge on a modestly higher score. So I requeue'd all 12 runs, and then requeue'd them all again to terminate after 50,000 cycles rather than 200,000, because with ten times more recombination per cycle they may not need nearly as many cycles. I was thinking that having more recombination would let them run faster, but I forgot that, with more recombination, each cycle will take longer. Hmm, maybe I should even set them for only 20,000 cycles. I'll see how far they've gotten tomorrow morning.
One bit of data we'll now include is the density of uptake sequences in the equilibrium genomes we discuss. But when I went back to extract this data from the appropriate runs, I found that one run didn't have the data because it hadn't terminated when it was supposed to; there was a typo in the specified termination cycle (2000o0 rather than 200000), so it would have kept running forever if I hadn't stopped it.
And when I went to redo that run without the typo, I discovered that the set of 12 runs it belonged to had all had another error; instead of recombining 1000 fragments each cycle they had only recombined 100. Fixing this won't change the conclusions at all; the runs will just all converge on a modestly higher score. So I requeue'd all 12 runs, and then requeue'd them all again to terminate after 50,000 cycles rather than 200,000, because with ten times more recombination per cycle they may not need nearly as many cycles. I was thinking that having more recombination would let them run faster, but I forgot that, with more recombination, each cycle will take longer. Hmm, maybe I should even set them for only 20,000 cycles. I'll see how far they've gotten tomorrow morning.
Are the purR knockout mutants not really purR knockout mutants?
The meticulous RA thought it would be wise to use PCR to check the genotypes of the purR::kan knockout mutants I used for my time course last weekend. (I had already checked that they were both resistant to kanamycin.) So she designed and ordered some primers that would flank the insertion that was described in the notebook of the grad student who originally made the mutant, and did colony PCR on all four of the strains I had used.
Much to my surprise, all four strains produced bands of the size expected for purR+ cells (about 1.0 kb), and none of them produced bands of the size expected for the purR knockout (about 2.2 kb). Either there's something wrong with the PCR analysis (and she's very meticulous so I doubt that), or the strains aren't what we've been thinking they are.
I had made these strains by transforming cells with DNA I had isolated from cells grown from the old frozen stock of purR cells made by the grad student (at least, that's what I thought I was doing), and selecting for kanamycin-resistant transformants. Could I have used the wrong DNA? Or grown up the wrong cells from the freezer?
We know that the original cells made by the grad student had the correct mutation, both because he had carefully checked them out and because a technician had later thawed a vial and done a microarray analysis of RNA. This showed that the mutant dramatically overexpressed all the genes that were predicted to be repressed by PurR in wildtype cells.
So tonight I've streaked out more cells from the last freezer vial of the original purR knockout, and on Monday the RA will test them by PCR. I also located the DNA I had used for that transformation, so she can test that by PCR too. If these cells give the expected 2.2 kb band, we'll assume something went wrong with my transformation. If they give the 1.0 kb band, we'll carefully check out the new PCR primers and probably run a quantitative PCR of a PurR-repressed gene on RNA from the original mutant and from one of the new mutants (with wildtype cells as control). Or, because the RA has recombineering working well now, she might just remake the purR mutant with her new primers.
If the mutants I used for my time course turn out to not be purR-, I think we'd still be really interested to find out where their kanR cassette is, because we don't have any other mutants with this interesting phenotype. That can be done by cloning out the kanR cassette and flanking sequences (the old-fashioned way or by inverse PCR) and then sequencing the DNA on one or both sides of the cassette.
Much to my surprise, all four strains produced bands of the size expected for purR+ cells (about 1.0 kb), and none of them produced bands of the size expected for the purR knockout (about 2.2 kb). Either there's something wrong with the PCR analysis (and she's very meticulous so I doubt that), or the strains aren't what we've been thinking they are.
I had made these strains by transforming cells with DNA I had isolated from cells grown from the old frozen stock of purR cells made by the grad student (at least, that's what I thought I was doing), and selecting for kanamycin-resistant transformants. Could I have used the wrong DNA? Or grown up the wrong cells from the freezer?
We know that the original cells made by the grad student had the correct mutation, both because he had carefully checked them out and because a technician had later thawed a vial and done a microarray analysis of RNA. This showed that the mutant dramatically overexpressed all the genes that were predicted to be repressed by PurR in wildtype cells.
So tonight I've streaked out more cells from the last freezer vial of the original purR knockout, and on Monday the RA will test them by PCR. I also located the DNA I had used for that transformation, so she can test that by PCR too. If these cells give the expected 2.2 kb band, we'll assume something went wrong with my transformation. If they give the 1.0 kb band, we'll carefully check out the new PCR primers and probably run a quantitative PCR of a PurR-repressed gene on RNA from the original mutant and from one of the new mutants (with wildtype cells as control). Or, because the RA has recombineering working well now, she might just remake the purR mutant with her new primers.
If the mutants I used for my time course turn out to not be purR-, I think we'd still be really interested to find out where their kanR cassette is, because we don't have any other mutants with this interesting phenotype. That can be done by cloning out the kanR cassette and flanking sequences (the old-fashioned way or by inverse PCR) and then sequencing the DNA on one or both sides of the cassette.
What on earth is 'constructive neutral evolution'?
Ford Doolittle gave a talk here today in the Biodiversity seminar series, which is attended by all the evolutionary biologists. It was titled 'Irremediable Complexity', and was promoting a concept originally published by Arlin Stoltzfus under the title 'On the possibility of constructive neutral evolution' (here, but probably behind a paywall). I haven't read it but it's been more influential than Ford said, cited 103 times.
Arlin's title is not at all self-explanatory; here's what I now think the words are intended to mean: 'Evolution' means 'a change over time in how a function is accomplished'. 'Constructive' means 'the change is that the function is accomplished in a more complex way'. And as a result of some helpful questions at the end, I now think that 'neutral' means 'the function itself is under stabilizing selection but not under adaptive/directional selection, and how it is accomplished (the change in complexity) is not under selection at all'. Ford didn't define 'complexity' until the question period; he then suggested that one measure of a function's complexity might be the number of components required for it.
The issue didn't seem very important to the evolutionary biologists in the audience, I think because they don't constantly deal with the just-so-story functions that molecular biologists typically ascribe to any complicating feature of a process. To many molecular biologists, every base pair in the genome, every intermolecular interaction, and every small RNA in the cell is the product of adaptive selection. There are no accidental interactions. Shit never just happens.
*On the other hand (not considered by Ford at all), mutations whose effects are made worse by the accidental interaction will be more efficiently eliminated by the stabilizing selection on the function. I don't think this can be said to reduce the complexity of the function, because the interaction was accidental and thus not included in the complexity count. I don't know if it would it create selection against the interaction.
**Psci Wavefunction has blogged about this concept in some detail, here and here. I confess that I haven't read these very long posts through, but perhaps now I will. (She asked an excellent question after the talk.)
*** Somewhere in his talk Ford was describing clade selection; using the example of how a propensity to speciate can cause a lineage to have many more species than other lineages. He said that more species means more individuals, but that's certainly not true.
Arlin's title is not at all self-explanatory; here's what I now think the words are intended to mean: 'Evolution' means 'a change over time in how a function is accomplished'. 'Constructive' means 'the change is that the function is accomplished in a more complex way'. And as a result of some helpful questions at the end, I now think that 'neutral' means 'the function itself is under stabilizing selection but not under adaptive/directional selection, and how it is accomplished (the change in complexity) is not under selection at all'. Ford didn't define 'complexity' until the question period; he then suggested that one measure of a function's complexity might be the number of components required for it.
Here's the executive summary:
Once organisms have evolved to have many components, some components will inevitably interact with others in 'accidental' ways that have, at least initially, not been shaped by selection. Once these accidental interactions exist, they will modify how selection acts on mutations that affect the function, sometimes making things worse but sometimes mitigating the effects of what would otherwise be deleterious mutations eliminated by selection*. These mitigating effects will weaken stabilizing selection on the function, sometimes allowing the mutations to be preserved (especially if populations are small). Preservation of the mutation effectively creates selection for maintenance of the formerly-unselected interaction. The function has become more 'complex (by Ford's definition), but there hasn't been any selection for the complexity. If mutations with these kinds of effects recur repeatedly, the function will become increasingly complex without having been in any way improved.As evidence that this type of complexity-building is common and important, Ford cited several molecular examples where a process has become ridiculously ('stupidly') complex but doesn't work any better that simpler versions. The RNA editing of trypanosomes is not well known but is a compelling example. So are introns and the spliceosomal machinery that lets eukaryotes cope with them. Simpler examples are the 'maturation proteins' that assist type I and II self-splicing introns. The ribosome itself may be a (not very stupid) example, where proteins have gradually taken over activities originally handled by the catalytic RNAs.
The issue didn't seem very important to the evolutionary biologists in the audience, I think because they don't constantly deal with the just-so-story functions that molecular biologists typically ascribe to any complicating feature of a process. To many molecular biologists, every base pair in the genome, every intermolecular interaction, and every small RNA in the cell is the product of adaptive selection. There are no accidental interactions. Shit never just happens.
*On the other hand (not considered by Ford at all), mutations whose effects are made worse by the accidental interaction will be more efficiently eliminated by the stabilizing selection on the function. I don't think this can be said to reduce the complexity of the function, because the interaction was accidental and thus not included in the complexity count. I don't know if it would it create selection against the interaction.
**Psci Wavefunction has blogged about this concept in some detail, here and here. I confess that I haven't read these very long posts through, but perhaps now I will. (She asked an excellent question after the talk.)
*** Somewhere in his talk Ford was describing clade selection; using the example of how a propensity to speciate can cause a lineage to have many more species than other lineages. He said that more species means more individuals, but that's certainly not true.
Contest to win an Ion Torrent DNA sequencer
The postdoc discovered that a new company called Ion Torrent is having a contest. The prizes are two of their new Personal Genome Machines.
The object of the Contest is 'to submit the best ideas for development of new applications for DNA sequencing'. We originally interpreted this as asking for the best research proposal using the machine (i.e. what we would do with the machine) but now I think it's asking for more general brilliant ideas for applications, not necessarily a project to be carried out by the winner.
On reading over the eligibility restrictions, I just discovered that entries can only come from people who live in the USA (but for some reason not people living in Arizona). That lets us out.
The object of the Contest is 'to submit the best ideas for development of new applications for DNA sequencing'. We originally interpreted this as asking for the best research proposal using the machine (i.e. what we would do with the machine) but now I think it's asking for more general brilliant ideas for applications, not necessarily a project to be carried out by the winner.
On reading over the eligibility restrictions, I just discovered that entries can only come from people who live in the USA (but for some reason not people living in Arizona). That lets us out.
Successful time course of competence development
Yesterday I did the failed time course again. The goal was still to replicate the earlier quick-and-dirty experiment that had suggested that knocking out the purine repressor prevented competence development in late-log cultures. This time the cells grew better, and the results are clear.
I had four strains: KW20 is wildtype, RR3005 has the purR knockout, RR699 has the sxy-1 hypercompetence mutation that we think should make competence induction less dependent on depletion of nucleotides, and RR1345 has both the purR and sxy-1 mutations. The graph below shows that the wildtype and sxy-1 strains grew at similar rates, and the two strains with the purR mutation grew a bit slower, perhaps because they were wasting resources on synthesizing nucleotides. (All four cultures stopped growing at about half the density they should reach with the best medium.)
The next graph shows the transformation frequencies of the four cultures at the same times. The wildtype cells (blue diamonds) showed the usual pattern, with very low transformation frequencies when cells were growing exponentially (first time point), and 1000-fold higher transformation when the culture became dense. The purR mutant (blue circles) also started out very low, but its transformation frequency remained low throughout growth, about 200-fold lower than its wildtype parent.
The sxy-1 mutant (green squares) also behaved normally. Its log-phase transformation frequency was >1000-fold higher than the wildtype strain, and it became about 50-fold more competent when the culture got dense. (Its final competence and that of the wildtype strain were both a bit lower than I normally see - I suspect this is due to the lower growth in the poorer medium.) The transformation frequencies of the purR sxy-1 double mutant (green triangles) were lower, but only about 3-9-fold.
So this experiment confirms both observations from the quick-and-dirty one. First, the purR mutation does prevent the competence development that normally occurs when cultures get dense. Since this mutation's major effect is to keep the purine biosynthesis pathway maximally active even in exponential growth, this suggests that running short of purines (purine nucleotides?) is the signal that normally induces competence when cultures get dense. The microarray analysis showed that wildtype cells at high density still have enough of the purine precursors hypoxanthine and inosine to keep PurR in repressing mode.
Second, the sxy-1 mutation makes cells much less sensitive to the competence-inhibiting effect of the purR mutation. The mutation causes hypercompetence by weakening the secondary structure of sxy mRNA, so this new result supports our hypothesis that the function of the secondary structure is to sense depletion of nucleotide (purine) pools. When the stem is weakened by mutation, it behaves as if nucleotides are depleted even when they're not, causing many cells make enough Sxy protein to become competent even in log phase. Some of the other sxy hypercompetent mutations have stronger effects (sxy-2 and maybe sxy-3), so I need to check if they are even less sensitive to the purR mutation.
I should also make a purR double mutant with the other kind of hypercompetence mutation. We know that some point mutations in murE, a gene responsible for one of the steps in cell wall synthesis, cause even stronger hypercompetence than mutations in sxy. But we have no idea how these mutations do this - we've ruled out most of the obvious explanations. (I would have thought I'd posted previously about this set of mutants, but I can't find anything by searching for 'peptidoglycan' or 'murE' or 'cell wall', so maybe I haven't. I'd better do a separate post about them.)
We already know that the mRNA secondary structure limits translation of the sxy mRNA into Sxy protein. In my mind, the simplest way for the secondary structure to sense depletion of nucleotide pools is the following: (1) Depleted pools slows the rate of mRNA elongation; (2) Because the two parts of the main stem are separated by ~100 nucleotides (I forget the actual number), slower elongation delays the formation of the secondary structure. (3) Because the ribosome binding site and start codon are in the region between these parts, this delay makes them more accessible and increases the initiation of translation. (4) once translation has started, the secondary structure can't form.
I would really like to complete the story by showing that the rate of transcription determines the efficiency of translation.
I had four strains: KW20 is wildtype, RR3005 has the purR knockout, RR699 has the sxy-1 hypercompetence mutation that we think should make competence induction less dependent on depletion of nucleotides, and RR1345 has both the purR and sxy-1 mutations. The graph below shows that the wildtype and sxy-1 strains grew at similar rates, and the two strains with the purR mutation grew a bit slower, perhaps because they were wasting resources on synthesizing nucleotides. (All four cultures stopped growing at about half the density they should reach with the best medium.)
The sxy-1 mutant (green squares) also behaved normally. Its log-phase transformation frequency was >1000-fold higher than the wildtype strain, and it became about 50-fold more competent when the culture got dense. (Its final competence and that of the wildtype strain were both a bit lower than I normally see - I suspect this is due to the lower growth in the poorer medium.) The transformation frequencies of the purR sxy-1 double mutant (green triangles) were lower, but only about 3-9-fold.
Second, the sxy-1 mutation makes cells much less sensitive to the competence-inhibiting effect of the purR mutation. The mutation causes hypercompetence by weakening the secondary structure of sxy mRNA, so this new result supports our hypothesis that the function of the secondary structure is to sense depletion of nucleotide (purine) pools. When the stem is weakened by mutation, it behaves as if nucleotides are depleted even when they're not, causing many cells make enough Sxy protein to become competent even in log phase. Some of the other sxy hypercompetent mutations have stronger effects (sxy-2 and maybe sxy-3), so I need to check if they are even less sensitive to the purR mutation.
I should also make a purR double mutant with the other kind of hypercompetence mutation. We know that some point mutations in murE, a gene responsible for one of the steps in cell wall synthesis, cause even stronger hypercompetence than mutations in sxy. But we have no idea how these mutations do this - we've ruled out most of the obvious explanations. (I would have thought I'd posted previously about this set of mutants, but I can't find anything by searching for 'peptidoglycan' or 'murE' or 'cell wall', so maybe I haven't. I'd better do a separate post about them.)
We already know that the mRNA secondary structure limits translation of the sxy mRNA into Sxy protein. In my mind, the simplest way for the secondary structure to sense depletion of nucleotide pools is the following: (1) Depleted pools slows the rate of mRNA elongation; (2) Because the two parts of the main stem are separated by ~100 nucleotides (I forget the actual number), slower elongation delays the formation of the secondary structure. (3) Because the ribosome binding site and start codon are in the region between these parts, this delay makes them more accessible and increases the initiation of translation. (4) once translation has started, the secondary structure can't form.
I would really like to complete the story by showing that the rate of transcription determines the efficiency of translation.
Cells behaving badly (is it the medium's fault?)
Yesterday I did a time course experiment, to see how the 'spontaneous' development of competence in the rich medium sBHI differed between wildtype cells and cells with either the sxy-1 hypercompetence mutation, the purR::kan knockout mutation, or both. But I had to give up halfway through because the cells stopped growing.
The graph above shows the densities of the four cultures as a function of time, with the purple line showing what I had expected based on the many previous time courses I've done. (I deliberately started with cultures at slightly different densities, to space the sampling out a bit.) They were all growing in medium from the same bottle, and they all stopped growing at about the same time.
Just before I had taken the first samples I had diluted all the cultures in medium from a new bottle, one that had been prepared on a different day, so I wondered if there had been something wrong with this batch, or if I might have forgotten to add one of the needed supplements to it (NAD). I quickly added more NAD to each flask (at time = ~150 minutes), but that didn't boost growth.
Then I tested several different batches of medium, including the the remaining of the second bottle I had used, as well as two different batches of NAD. Unfortunately the first bottle I had used was the last bottle of its batch, so I couldn't test it. I inoculated each with the same amount of cells, and let them all grow overnight. The second graph shows that there are substantial differences between different batches of medium, and that none of them gives the amount of cell growth I'd expect from previous time courses (labeled as 'years ago' because I haven't done one recently).
I don't think the problem is just how long the bottles of medium had been sitting on the shelf, as the components are typically quite stable. Instead I'm wondering if we might be using medium from a different supplier. In the past I'd noticed substantial differences in how well different brands of BHI supported cell growth, and had sworn to use only the best (Difco), but I know we were recently given some BHI from another supplier. Tomorrow I'll repeat the time course, with the March 1 medium.
Just before I had taken the first samples I had diluted all the cultures in medium from a new bottle, one that had been prepared on a different day, so I wondered if there had been something wrong with this batch, or if I might have forgotten to add one of the needed supplements to it (NAD). I quickly added more NAD to each flask (at time = ~150 minutes), but that didn't boost growth.
I don't think the problem is just how long the bottles of medium had been sitting on the shelf, as the components are typically quite stable. Instead I'm wondering if we might be using medium from a different supplier. In the past I'd noticed substantial differences in how well different brands of BHI supported cell growth, and had sworn to use only the best (Difco), but I know we were recently given some BHI from another supplier. Tomorrow I'll repeat the time course, with the March 1 medium.
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