Field of Science

Planning the GTA work

My goal for the rest of my time in Andrew Lang's GTA lab  is to gather data that constrains estimates of the efficiency of GTA transduction.  I have lots of ideas but they're not very well organized, and I keep getting distracted by the minutiae of GTA biology (and our general ignorance of same).  So this post is an attempt to get a sensible plan written out.

The bottom line for efficiency is how many transductants are generated for each cell that produces GTA and then dies. This depends on many factors, so I'm going to try to break down the steps and evaluate their limitations.

Here are some of the questions I'd like to know the answers to.  (Some of these questions overlap with others, and some of them are addressed by data we already have.)

  1. How many functional GTA particles does a cell produce under 'normal' conditions?  
  2. Are lots of defective particles produced too?
  3. Do individual cells of 'overproducer' mutants produce more GTA particles than normal cells, or is overproduction due only to more cells being producers? 
  4. How stable are GTA particles in the cultures where they are produced?
  5. How stable are GTA particles in more dilute solutions?
  6. Do GTA particles bind to free capsule or to cell-envelope components released by lysed cells?
  7. Do cells in producer cultures bind the GTA particles produced by other cells and take up ands recombine their DNA? 
  8. How good are recipient cells at finding GTA particles when cells and particles are scarce?
  9. Do cells die if they are exposed to too high a concentration of GTA particles?
Experiments I'm going to do:
  1. Measure stability of GTA titers in culture filtrates stored at room temperature.
  2. Measure growth of wildtype and overproducer strains by plating dilutions and counting colonies, in addition to measuring culture density by its turbidity.  At the same time measure accumulation of GTA (see this post and this post).
  3. Compete an overproducer mutant against its isogenic parent during growth under GTA-producing conditions, to estimate the cost of GTA production.  This is especially important since my most recent growth curves don't show much difference between ovverproducer and wildtype strains.  This requires that one strain carry an antibiotic resistance marker the other lacks, so I'm using GTA to transfer a kanamycin-resistance marker from a derivative of the 'wildtype' strain into its overproducer sibling.  Then I can do the competition both ways, starting with the kanR overproducer at low frequency in a background of kanS wildtype cells, or starting with the kanR wildtype at low frequency in a background of kanS overproducer.  I have all these strains now (just confirming that the kanR overproducer does overproduce GTA), so I can start the experiment as soon as I grow up the cultures.  I should also Do a complete growthtiter the amounts fo GTA produced, since the goal is to get the ration of GTA produced to cells died.
  4. Do the same competitions, but between an overproducer and a no-GTA mutant, or between wildtype and no-GTA mutant
  5. Add marked GTA to a producer culture (to multiple different producer cultures) to see how efficiently the cells take up  GTA.  The producer strains are all rifR, so this needs a GTA prep carrying a different marker.  I've made a GTA filtrate that transduces kanR, but this transduction is very inefficient compared to rifR, no doubt partly because the kanR is a big insertion, not a point mutation.
  6. To get an independent antibiotic resistance point mutation, I've just started selecting for a spontaneous mutation giving resistance to streptomycin, by plating GTA-producer strains on streptomycin plates.  Mutations giving strR are common and this selection has been successful for R. capsulatus in the past.
  7. Do a GTA-producer time course analysis that distinguishes between GTA production and GTA accumulation.  Experiments to date have just assayed the amount of GTA in the culture at different times, and there are unexplained peculiarities about the results (see this post:

Scheduling complication:  I'm here until August 12, but I'll be tied up with visitors for part of the time, next week and for the last two weeks of July.  Because R. capsulatus grows slowly, I need to wait two days to see the result of each experiment.  

I could do the first 'quick-and-dirty' version of the competition experiment now, starting the cell mixtures tomorrow (Friday) morning and growing them for only 24 or 48 hr, taking time point samples at t=0, t=24 and t=48 (Sunday morning).  Then I could count the colonies on Tuesday morning.  Will I also measure the amount of GTA in each mixture, by its ability to transduce rifR and kanR?

Why doesn't all the GTA get taken up?

I've been modelling the production and uptake of GTA particles in a culture, hoping to understand the cause of the surprising GTA-accumulation curve I described in the previous post.  But this has led me to a more fundamental surprise.
Only a very small fraction of the cells in a GTA+ culture produce GTA particles and lyse, and all the other cells are able to bind GTA particles and take up their DNA.  So why doesn't all the new GTA quickly get taken up by all the surviving cells?
Here are the basic principles I've been assuming, based on what's in the literature:  GTA production:  Cells in exponential growth don't produce GTA.  The GTA genes are turned on as the culture density gets high and growth slows.  Once the culture reaches its stationary-phase density GTA production stops.  GTA uptake:  Cells in exponential growth express the capsule genes at a low level and bind GTA particles with moderate efficiency.  The capsule genes are turned up when culture density reaches a quorum-sensing threshold, and ability to bind GTA particles gradually increases.  Stationary phase cells bind GTA particles efficiently.  GTA decay: BTA particles are moderately unstable, so they fall apart with some unknown probability.

Let's put some numbers to this:

  1. Assume that 1% of cells produce GTA over the course of the permissive stage.
  2. Assume that each producer cell produces 100 particles and then dies.
  3. Assume that each non-producer cell can take up 1 GTA particle.

Result:  All the GTA particles are taken up.  The concentration of GTA particles in the medium falls to zero.

In reality, assumption 1 is likely to be an overestimate, and assumption 3 an underestimate.  I'm going to do some experiments to see if I can clarify what's going on.

Marc Solioz's 1975 PhD thesis on GTA

PhD students, don't assume that your thesis will moulder unread in the library.  More than 40 years after he submitted it, I'm reading Marc Solioz's PhD thesis (The Gene Transfer Agent of Rhodopseudomonas capsulata).  I want to understand the kinetics of GTA production, and his is the only good data I can find.

Here's what he reported:

A. Stability of and transduction by GTA in various solutions:  He tested a wide range of solutions.  In these studies he didn't try to distinguish between conditions that stabilize GTA for storage and conditions that maximize its ability to attach to cells and inject its DNA.  It's happiest in 1mM each of Na+, Mfg+ and Ca++.  This can be buffered with 10 mM Tris, with or without gelatin or BSA (no effect).  It's destabilized by 10% gycerol, even for freezing.  GTA preps made by filtering culture supernatants should be diluted at least 10-fold to reduce the destabilizing effect of the medium constituents.

B. Inactivation by other factors:  GTA's stability is not affected by temperatures up to 50°C.  Keeping it on ice is not better than room temperature, and there was no difference between partially purified and purified stocks.  It's inactivated by proteases but not RNase or DNase.  It's not inactivated by ether or chloroform, or by phospholipases, consistent with the absence of any membrane.

C. Inactivation by UV:  UV damages DNA so it is expected to inactivate the transducing activity of GTA particles.  To control for experimental variation (a big concern with UV experiments), he compared GTA inactivation to inactivation of phage T2 UV'd together in the same solution.  The action spectra are the same for GTA and T2, but GTA inactivation requires much higher doses, consistent with the small amount of DNA in each particle.

D. Conditions and kinetics of GTA production:   1. Production kinetics: This is the same surprising result (Solioz's term) I showed in the previous post. Cells were grown photosynthetically/anaerobically in a yeast extract + peptone medium.  The dashed line approximates the combined growth curves seen in the four independent experiments, but it's in 'arbitrary units' (I think on a log scale) so I have to infer the cell densities from how my cells grow.

He reports that the initial peak and drop were consistently seen across all his experiments, but that sometimes the drop was not followed by the final high-GTA stage.  He saw a similar pattern using a strain that does not absorb GTA (strain H9), so the changes in GTA titre are not due to changes in the removal of GTA particles from the medium.   However this conclusion is weakened by the description of strain H9 in the methods, which just says 'does not act as a recipient of GTA, with no reference'.)  Other tests he did could not rule out effects of transient inhibitory/inactivating factors in the culture supernatant.

2. Effects of growth conditions on production.  Defined medium RCV gave low titres of GTA.  Yields with different concentrations of yeast extract and/or peptone were variuable, apparently depending even on the batch no. of ingredient used.  Variation sin culture growth rate and final density did not correlate with GTA titres.

3. Isolation of mutants:  He attempted to isolate an overproducer mutant but failed.  The original producer strain B10 carried two phages, so he made a derivative strain, SB1003, that was cured of the phages and carried the convenient RifR point mutation.   This new strain is the one I have been using as the standard donor; it's good to know its provenance.

4. Radiolabelling:  He put in a lot of work to find a way to radioactively label GTA.  This was used to guide the purification studies.

E. Purification of GTA particles:  This is a long section that's not of much interest to me.  He tested a wide range fo the available biochemical techniques used for purification of organelles, phage and molecules.

F. Characterization of the nucleic acid:  He used the single-strand-specific nuclease S1 to show that the DNA in GTA particles is double-stranded.  He used CsCl ultracentrifugation to estimate its base composition as 65% G+C, the same as that of the R. capsulatus genome.  Repeating this analysis with heat-denatured DNA confirmed that the DNA is linear, not closed-circular like plasmid DNA.  Banding in a CsSO4 gradient showed that it is not extensively modified.  In sucrose gradients it co-sedimented with SV40 DNA, suggesting a size of 3.6 x 10^6 Daltons.  How big is this in base pairs or kb, you ask - about 5.5 kb.  He says it would be better to run the DNA in an agarose gel, but this emerging technology wasn't available to him yet.

G. Examination of GTA with the Electron Microscope:  He saw lots of tails, and empty heads, some with tails.  Apparently-full heads came in different sizes, from 150-600 Angstroms  in diameter (15-60 nm).  But he thinks much of this may be artefacts of the purification and EM-preparation procedures.

Summary of R. capsulatus Bioscreen growth curves

The previous post (GTA competition experiments) described the results of the follow-up set of R. capsulatus growth curves that I planned at the end of the previous experiment (R. capsulatus growth curves in RCV medium).  But it didn't pull together the results of all the Bioscreen growth curves, nor integrate them with what was previously known/thought).  So here goes:

First, what's already been reported about growth in liquid culture?  Not a lot.  The graphs below are all I could find.  (I asked my colleague here - he says he doesn't know of any others.)

GTA production: 

The only work that measured GTA production along with growth is Solioz et al. 1975, and their 'growth curve' is just a schematic.  The titers of GTA this shows are very peculiar.  The titer is very low while the culture is growing, and rises to about 3x10^4 just before culture density levels off.  But then it dips sharply, falling to about 10^3 over a few hours, and then rises again to its final stable level of about 4x10^5.

I don't understand how the titer can fall that quickly.  Where do the GTA particles go?  The titers are transformants to RifR or StrR, so the total number of active GTA particles per ml is about 1000-fold higher, so ~4x10^7 at he first peak, and 10^6 at the valley.  Perhaps there's an initial burst of GTA production that stops abruptly, and most of the released GTAs are quickly lost because they attach to the remaining cells.  There would be at least 10^8 cells at that stage so this could easily happen.  After a few hours the second wave of GTA production begins.  This produces at least 4x10^8 GTA particles that remain free (and possibly many that attach to cells and are not detected).
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">
Are the GTA titers from my last Bioscreen run comparable?  I got 780 RifR transductants per ml, from a culture that had about 10^9 cells/ml; this is about 20-fold lower than Solioz et al. reported, and about 3-fold lower than I saw in an earlier (not-Bioscreen) culture.   The difference may partly be due to the different culture conditions in the Bioscreen.
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">Effects of PO4:  
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">The Westbye graphs on the lower right come from a study of the effects of phosphate levels on GTA production.  This is in the defined medium RCV, either with its normal 10 mM PO4 or with only 0.5 mM PO4.  Low PO4 allowed higher GTA production.  Differences in PO4 did not affect the culture density of the normal strain SB1003, probably because less than 1% of the cells in a culture produce GTA, but low PO4 caused a drop in the density of the overproducer strain DE442, where up to 20% of cells are thought to produce GTA.  The phosphate effect is thought to be on release of GTA particles from the producer cells, not on GTA synthesis or on stability of parrticles in the medium.
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">In my Bioscreen runs I saw the effect of low PO4 on GTA levels, but no the predicted drop in culture density of DE442.   Instead both DE442 cultures levelled off at densities well below that of both SB1003 cultures.
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">
<10 100-fold="" 20-fold="" 3="" a="" about="" again="" and="" are="" as="" but="" cells="" exponentially="" falls="" final="" going="" growing="" hrs.="" is="" it="" its="" just="" level="" linear="" ml="" nbsp="" of="" on="" over="" p="" quickly="" rapidly="" rises="" scale...="" schematic="" the="" then="" to="" was="" while="" write="" x10="">

GTA competition experiments

I'm in St. John's for the 'summer'*, doing GTA-related experiments in Andrew Lang's lab at Memorial University of Newfoundland ('MUN').

The first experiments I'm going to do are growth competitions between GTA-producing strains and otherwise-identical non-producer strains created by deleting the GTA genes.  Because GTA production requires cell lysis, we predict that the non-producers should outcompete the producers.

While I was still in Vancouver I did detailed growth curves of the various strains.  Preliminary ones are described here, and I'll paste the graph from the latest ones below:

I wanted to check the effect of phosphate concentration in GTA production and culture growth, so I only used two strains, SB1003 (wildtype) and DE442 (a GTA overproducer).  I used two PO4 concentrations; 0.1 mM, which should allow high GTA production and reduced growth, and 10 mM, which should cause low GTA production and better growth.  The growth differences should be detectable only for DE442.  (I also used three different cell densities.  I'm only showing the results for cultures started at the highest density, but the others grew similarly with the expected delays.)

I measured GTA production at two times, by removing cultures from some wells, filtering out the cells, and using the cell-free supernatants to transduce an RifS strain to RifR.

The results are below.  (The upper graph is plotted on a linear scale, and the lower graph is the same data plotted on a log scale, for easier comparison of growth rates.)  The growth curves are very similar to those from a previous experiment (RR#1438) where I didn't measure GTA production.

The GTA production happened as expected.  SB1003 produced no significant GTA in high PO4, and a modest amount (780 transductants per ml) in low PO4.  DE442 produced lots more GTA under all conditions, but about 4-fold more in low PO4 than in high PO4.

On the linear scale the two strains appear to have very similar exponential growth rates, but the log scale reveals that DE442 (the GTA overproducer) is slower in exponential growth.  DE442 also reaches a lower final densities (SB1003, OD ~ 1 - 1.08; DE442 OD ~ 0.7).

The growth differences are unlikely to be directly due to the lysis required by GTA production, because the GTA differences caused by the different PO4 levels do not correlate with OD differences.

DE442 is not isogenic with SB1003; it carries a mutation that blocks synthesis of the red accessory pigment.  Could DE4432’s pigment phenotype be responsible for its poorer growth?  These were aerobic cultures in a dark room, so the growth difference is not a direct consequence of differences in photosynthesis.

I don't think it would be straightforward to transfer the ‘overproducer’ mutation into the SB1003 background, since typical transduction frequencies are less than 1/1000, and we have no way to select for overproducer colonies against the background of normal colonies.  If the pigment difference causes the growth difference, we could transfer the wildtype pigment allele into DE442 or the mutant allele into SB1003.  I wonder how the parent strain of DE442 (Y262, I think) grows.

* It's definitely not summer yet here.  Icy winds anywhere near the coast, and several thin snowfalls in the last few days. I remain hopeful, because most of the trees are finally getting their leaves, and the spring bulbs are blooming.

Wait, there's a much simpler explanation! (For CRISPR-Cas, not for GTA)

I'm in Halifax for a couple of weeks, visiting Ford Doolittle and his philosophical colleagues,  We've spent much of the time considering the extent to which CRISPR-Cas systems can or should be considered 'Lamarckian'.  I started with the simplistic perspective that of course it is, because an acquired character (immunity to future phage or plasmid infection) becomes inherited because the Cas proteins insert short phage- or plasmid-derived DNA sequences as a CRISPR 'spacer' into the chromosome.

Here's a very detailed diagram I made of the evolutionary events (mutation and selection) affecting CRISPR-Cas systems (click to embiggen):

We ended up concluding that 'directed mutation' was a better perspective.

But, once our ideas started settling down, this detailed diagram got me thinking about how uncertain and far in the future the 'immunity to future infection' benefit is.  That's a problem for CRISPR-Cas evolution, since this uncertainty greatly weakens the selection maintaining and refining the system.  Iv selection is too weak, the system shouldn't be maintained at all.

A more urgent problem is that the cell needs to survive the immediate infection/invasion before it has any chance of benefiting from the long-term immunity.  This becomes especially important if the bias against potentially-lethal self-spacers arises because the cell contains many copies of the invader genome.

But the cell does have a very nice mechanism to clear the invader, because it has just created an invader-specific spacer in its CRISPR array.  Transcribing this new spacer would give it many copies of an invader-specific crRNA with which Cas9 can destroy all the copies of the invader genome.

So here's my new hypothesis:
The primary function of CRISPR-Cas systems is the detection and immediate destruction of phage and /or plasmid DNA.  Benefits from immunity to future infection are relatively unimportant. 
Things I need to find out:

 Is this a new idea?  I don't remember seeing it anywhere, but if any reader knows of a prior proposal please let me know in the comments or via Twitter (@rosieredfield).

Is relevant data available?  The basic experiment is, in principle at least, quite simple.  Do cells with an intact CRISPR-Cas system survive phage infection better than cells with a defective system?  Do they become transformed less efficiently by plasmids?  These tests would be most sensitively done under sub-optimal infection conditions.

How is transcription of the Cas genes and CRISPR array regulated?  In particular, how efficiently is the CRISPR array transcribed and processed immediately after a new spacer has been added?  In the context of my GTA-as-CRISPR-vaccine ideas (see this post from a few months ago) I'd been looking for reports that new CRISPR spacers can be immediately transcribed, creating crRNAs that can immediately attack the original invader.  I didn't find any solid data, but neither did I find anything that ruled this out. 

R. capsulatus growth curves in RCV medium

My upstairs GTA colleague and I were surprised that the Bioscreen growth curves in the previous post didn't show a dip in OD600 of the GTA-overproducer strain like that seen in manual (non-automated) growth curves.  This dip is thought to be caused by the lysis of GTA-producing cells as GTA production peaks when cells hit stationary phase.

We thought part of the problem might be that I used the standard YPS medium which is based on modest concentrations of yeast extract and peptone.  The clearest/most-recent published demonstration that GTA-producing cultures used RCV, a simpler 'defined' medium based on malate, and showed that the apparent lysis occurred in medium with 0.5 mM PO4 but not in medium with 10 mM PO4.

So I redid the growth curves for all 6 strains, using both high-P and low-P versions of RCV (kindly supplied by my upstairs colleague).  The results are not inconsistent with the Westbye results, but they're not at all compelling.  None of the strains decreases in OD600

The problem is that there's quite a bit of between-strain variation in growth and in the stability of the stationary phase OD.  (Within each strain the replicate wells give very similar results, with one exception.)

The graph below shows growth in the high-phosphate medium.  The main graph shows OD600 on a log scale (appropriate to exponential growth), and all the strains appear to stably reach similar densities.  But the inset shows the same data on a linear scale, which makes the variation look more significant.  The overproducer strain stops growing abruptly at OD600 = 0.7 a lower density than the other strains.

Here's the cells in the low-phosphate medium.  There's an initial drop in OD600, over the first 10 hours, but then all the strains grow steadily except strain YW1, where the individual wells grew at different rates for no apparent reason.  Again the linear-scale inset shows the substantial variation at stationary phase.  The overproducer DE442 again stops growing, this time at OD600 = 0.8, and now its OD falls by about 20% over the next 40 hours.

I really don't feel comfortable drawing any solid conclusions from this one experiment, especially since there's a blip in many of the growth curves at a point where I stopped and restarted the runs to add more time when I realized that 3 days wasn't going to be long enough.  Even though the shaking only stopped for 2-3 minutes, and the trays of cells remained in their holder with the lid closed, most of the strains had an abrupt change in OD600.  (You can see the blips at hour 63.)

Plan:  Do the run again.  This time I'll pre-grow the cells into log phase in high-P and low-P RCV.  medium (the upstairs colleague has offered me enough medium to do this).  And I'll plan on pausing the run at key times to take samples that I can assay for GTA production.

What can we learn from growth curves?

Here's the results of the Bioscreen growth curves I ran for Rhodobacter capsulatus strains:

Each dot is the mean OD600 of 15 replicate wells, each containing 300 µl of culture, with ODs read every 20 minutes for 45 hours.  The cultures all grew up at about the same times, but I've shifted the X-axes so the curves don't overlap.  OD values below about 0.015 are not significantly above the backround absorption of the culture medium. The Y-axis is a log scale, so when doubling time is constant the dots will fall in a straight line.

I did these runs 'just-in-case', because I'm going to be working with Rhodobacter capsulatus at Memorial University in Newfoundland for the next few months (on sabbatical leave) and thought they probably wouldn't have a convenient Bioscreen that I could use.

Now I need to figure out what we learn from these, and whether I should do any more experiments before I leave UBC.

The simplest expectation is that once the cells have adjusted to the medium (after 'lag phase') they will grow at a constant rate until they run out of nutrients or experience other bad consequences of high cell density (little oxygen, accumulation of toxic byproducts).  But all of these cultures instead exhibit 'diauxy', a mid-growth shift from one resource to another.  We see  this as a brief slowing or even cessation of growth at about OD=0.05 (orange shaded band), after which growth resumes, often at a different rate.  The pause occurs because the cells need time to adjust their metabolism to a change they've caused in the medium, such as exhaustion of one nutrient or new availability of another. 

I don't know enough about R. capsulatus metabolism to speculate about what the change might be, but it might affect production of Gene Transfer Agent particles.  The pause isn't due to lysis of GTA-producing cells, because it's not changed in the ∆∆ strains, which have deletions of the GTA gene cluster and lysis gene.

SB1003, B10 and YW1 are all 'wildtype' strains, I think.  Strain YW1 grows much slower than the others, although it still speeds up after the growth pause, and it reaches a slightly lower final density.

Strain DE442 carries a mutation that causes over-expression of the GTA genes and over-production of GTA particles.  Growth curves in a 2013 paper found that this strain had a substantial drop in OD once growth ceased, thought to be due to lytic release of GTA particles, but no drop is seen in the Bioscreen culture.  That work used a low-phosphate version of a different medium, RCV.  But an earlier paper found strong lysis with the same complex medium I used (YPS), and low lysis with the high-phosphate (10 mM) standard RCV medium.

The lab upstairs has both low-phosphate and high-phosphate versions of the RCV medium, so I'm going to repeat the time course with both.

growth time courses

In a few weeks I'll be headed for the Maritimes, for the final part of my sabbatical work on Gene Transfer Agent.  But before I leave here I want to run some detailed growth time courses on GTA-producing strains, taking advantage of the BioScreen machine belonging to the lab next door.

I'll first do a trial run with all the strains I have,  to check the basic growth kinetics under the Bioscreen growth conditions.  Then I'll see if I can combine the growth measurements with testing for the amounts of GTA produced.

Phage plaqueing still sucks - what to do now?

I feel like I've been sucked down a hole of trying to get consistently countable plaques from the Rhodobacter capsulatus phage I'm testing.  After seven weeks of plaqueing with various combinations of strains and agar concentrations and cell densities, I'm no closer to having a well-behaved phage I can use to test the GTA-as-vaccine hypothesis.

Along the way I've eliminated various sub-hypotheses:

1.  The plaques are tiny/faint/blurry/invisible because the phage capsids have long fibers that reduce diffusion through the top agar:  Test - use increasingly dilute top agar.  Top agar us usually 0.75% agar; I've taken this down to 0.3% (the lowest concentration that's still stable enough to handle). The first time I got somewhat larger plaques, but this was not reproducible.

2.  The plaques are tiny/faint/blurry/invisible because GTA gene products contribute to phage production:  Test:  Plaque phage on a GTA overproducer strain.  Result:  On the first try, plaques on the overproducer seemed larger.  But this was not reproducible.

3.  The plaques are tiny/faint/blurry/invisible because the GTA-as-vaccine hypothesis is true:  (Plaques can't grow because rapid diffusion of GTA particles allows surrounding cells to become CRISPR-resistant to the phage before the phage gets to them.)  Tested by plaqueing the phage on cells deleted for the entire GTA operon and for the separate endolysin.  Result:  Plaques on these '∆∆' strains are just as lousy (maybe more lousy) than on the GTA-producer parents.

4.  Variant (large) plaques contain mutations that increase infectivity or diffusion:  I made new lysates from a couple of big plaques that spontaneously appeared among the tiny plaques, but these lysates still gave tiny or no plaques

I know that the phage lysates do infect and kill the cells, and do produce progeny phages.   When I put a spot of sufficiently-concentrated phage onto a lawn, all the cells die, and when I make a lysate with this 'clear' top agar, I get way more phage then I put in.  Can I use the lysate to test the GTA-as-vaccine hypothesis even though I don't have countable plaques?

What would I do?  Here's an earlier blog post where I laid out a crude plan and a list of all the things I'd need to find out before actually doing the experiment that would test the hypothesis:

Luckily, after I wrote the above I made another grand attempt at titering the phages on the various strains.  Well, I made a sloppy attempt, learned from at and made a better attempt, which more-or-less worked. 

Basic test:  Pour lawns of the test strains, using cells concentrated from 400 µl of culture, in 1.5 ml of 0.4% top agar.  Put 10 µl spots of different dilutions of phage lysates onto these lawns, let the liquid absorb, and check the next day.  Yesterday I did this using photosynthetically grown cells (supposed to make better lawns) and today I've repeated it using cells grown aerobically in the dark.  Here's yesterday's result for one of the two phage and one of the six strains:

The central clear spot is undiluted lysate, and the other spots are 10-fold dilutions of that.  For undiluted, 10^-1, 10^-2 and 1-^-3, the spot is clear (all the cells have been lysed).  The 10^-4 spot still has patches of non-lysed lawn, and the 10^-5 and 10^-6 spots have distinguishable plaques.  Two of the four healthy strains gave countable plaques (27 and 29), which is nicely consistent.

I'll wait for tomorrow's results before proceeding.

Phage phrustration

Aacckk!  I've spent more than a month trying to get decent R. capsulatus phage plaques on R. capsulatus lawns.  Still no consistent success.  In one experiment I had much better plaques on cells of strain DE442 (a GTA overproducer), but that did not replicate.  I suspect that non-tiny plaques depend on exactly the right balance of the cells' physiological state, their density, the agar concentration, the culture medium, and other factors I haven't attempted to vary.

Yesterday I was almost ready to do a UV-irradiation experiment to generate mutant phages that make larger plaques, starting with two lysates I'd grown up from single plaques that were much larger than the rest.  But the dilution-series plates I was titering these lysates on grew up with very similar numbers of (mostly tiny) plaques, suggesting that phage contamination had crept into lysates with unexpectedly very low titers. 

This week I've gotten a couple of good suggestions from visitors:

The first visitor suggested that I give up on the characterized/sequenced phages I've been working with and just isolate a better-behaved phage from R. capsulatus's natural environment.  I'd have to learn how to do this (get water

Another visitor suggested that maybe the problem is the correctness of my hypothesis about GTA transduction of phage DNA leading to CRISPR-mediated phage immunity in the GTA recipient.  That is, maybe the first cells to get infected in my lawns produce so many phage-DNA-carrying GTA particles that many of the neighbouring cells that would otherwise be lysed by the phage become immune to the phage before the plaque can form.

There's a simple way to test this - see if the phage form better plaques on a strain that doesn't produce GTA.  So tomorrow I'm getting some GTA mutants from the guy upstairs.

Phage plans - let's put natural selection to work!

I have a two-pronged plan to get a phage strain that gives good enough plaques for my GTA-as-vaccine experiments.

I obtained reasonable titers of two phages, 'Titan' and 'Saxon'.  I'll invest a couple of weeks to see if I can get better and more reproducible plaques with either of these.  The genome sequences of these phages are not closely related.

First, improve the plaquing conditions:  The researcher who isolated the phages recommends using for the lawn cells that have been grown photosynthetically to a high density,  He also suggested trying a lower top-agar concentration.  I'll play around with these and other variables to see if I can get better plaques.

Second, use artificial selection to get a better strain of phage:  I'll pick the few best-looking plaques of each of my two phages and plate the phage they contain in new lawns.  From those new lawns I'll again pick the best-looking plaques, and plate their phage in new lawns.  Etc.  Maybe I'll introduce a bit of UV mutagenesis along the way.

The first step will be to make fresh lysates of these phages.  The lawns I made before are too old, so I'll grow up some cells for lawns today and tomorrow I'll retiter the lysates.  On Friday I can pick plaques from these lawns and make plate lysates.  If there's a plate with near-confluent plaques I can use it directly to make a plate lysate.  (10^7 or 10^8 pfu/ml, and I have maybe 5 µl so at best I can get plates with 5 x 10^4 or 5 x 10^5 plaques.  The latter might be enough to get a good lysate.  There are small volumes (50-100 µl?) of the original lysates in the lab upstairs, so maybe I'll sue these.  Or Maybe I should save these until I've improved the plaquing conditions.

thin lawns, feeble or absent phage

My phage titering gave disappointing results.  Three of the five lysates gave no plaques at all, and the other two gave small indistinct plaques that couldn't be accurately counted or characterized.

I took some photos of the plaques I did see.  The top photo is a section of one lawn, with several thousand tiny indistinct plaques.  (The blurry markings are the label on the bottom of the plate.)  The second photo is a closeup  of an area on another lawn where I had spotted more-dilute phage, taken with my iPhone's Olloclip zoom lens.  A few tiny plaques are visible, maybe 3, maybe 5.

For comparison, here's what nice plaques look like.  These are plaques of the E. coli phage lambda (source)

I won't be able to use these R. capsulatus phage for my GTA-vaccine experiments unless I can get better plaques.  I'll need to know whether the phage makes turbid or clear plaques, and I'll need to be able to count it accurately.

I can try using another strain as the host.  These lawns were made with a culture of strain YW1, the strain that these phage were originally isolated on.  I have several other strains, though I don't know if they are closely related.

I can also try changing the plating conditions.  I followed the protocol that I obtained from people who have worked with these phage, but perhaps I could grow the cells to a different density, or incubate the plates at a different temperature.   I'll ask the experts for advice.

Titering my lysates

Planning today's work:

Titering the phage lysates should be a no-brainer, but it's been a long time since I worked with phage so I'd better think things through before I do it.

I have 15 µl of each of 5 phage stocks ('lysates').  The original titers (plaque-forming units/ml, pfu/ml) are written on the tubes - they range from 6x10^5 pfu/ml to 2x10^11 pfu/ml.  But the lysates are probably quite old (maybe 2 years, maybe more), so their titers may have dropped a lot.

I think I'll do one poured-lawn plate of each, using an amount of lysate that should be about 1000 pfu according to the original titer.  And I'll do a spotted-lawn plate for each phage, using undiluted lysate  and a range of dilutions.

I'll dilute the lysates in the same YPS medium I've grown the cells in.  It has calcium and magnesium added so should be fine for typical phage.

About bacterial lawns and phage plaques

This was going to be a post where I do the planning to titer my new lysates today, but it turned into an explanation of how microbiologists use plaques in lawns of bacteria to study phages.

Wait, what's a 'lawn' and what's a 'plaque'?  A lawn is a thin layer of confluent bacterial growth, usually created by mixing a relatively large number of cells (≥10^6) with liquid agar solution ('top agar' or 'soft agar' and pouring the mixture onto the surface of a nutrient-containing agar plate. The top agar is usually at  0.5-0.75%, about half the concentration used for a normal solid plate.  The cells can't move around in the agar, and they grow to high density using the nutrients that diffuse upward from the bottom layer.

If a few of the initial cells were infected with a phage, the phage they release when they die will infect neighbouring cells and kill them, creating a cell-free zone called a 'plaque'.

Here is a detailed drawing of what's happening as a plaque forms:

Initially there's just one infected cell, and sparse uninfected cells in its neighbourhood.  When this cell lyses, the phages it releases can readily diffuse through the agar and infect nearby cells.  While this is happening, the uninfected cells are growing and dividing.  When the newly infected cells lyse, the phage they release add to the local population and infect more cells.  The phage continue to diffuse away, but soon the neighbouring cells become so dense that they stop growing and the phage can no longer replicate in them.  The cells are too big to diffuse through the agar like the phage, so lysis leaves a circular cell-free space called a plaque.  Typical plaques are 1-2 mm across so easy to see with the naked eye.

By counting the number of plaques that form in a lawn of bacteria, we know how many infectious phages were present in the mixture we poured on the plate.  This is the standard way to measure the number of phages (well, the number of 'plaque forming units', PFUs) in a preparation of phage (a 'lysate').

Regular poured-lawn method:  Cells and diluted phage are incubated together in a small volume of liquid (broth or phage-dilution solution) for long enough that most of the phage have attached to the cells.  Then hot liquid top agar is added to the tube and the contents are quickly mixed and poured onto an agar plate of whatever medium best supports lawn growth and plaque formation.  (Quickly so the mixture cools before the cells are damaged.)  The top agar quickly sets, and the plate is incubated overnight at an appropriate temperature for bacterial growth and phage plaque formation.

Spot-titer method:  Cells are quickly mixed with hot top agar (no phage) and the mixture is poured onto agar plates and left to set.  Sometimes the plates can be prepared days in advance, if the cells are happy sitting in the fridge. 10 µl dilutions of phage are then spotted onto the surface and the plates are incubated overnight as before.  If you're gentle you can even streak a drop of lysate across the lawn as you would streak cells on a normal plate, allowing you to grow well-isolated plaques without the nuisance of diluting your lysate.

Other ways we can use lawns and plaques:

Isolating phage from a single plaque:  Often you want to start an experiment with a genetically pure phage lysate that you grew up from a single plaque.  If plaques are well-separated (remember that the phage continue to diffuse out after the plaque forms and the lawn stops growing) you can use a Pasteur pipette to punch out the plaque away from the surrounding agar.  If this plaque is put into a small volume of phage-dilution solution, the many thousands of phages it contains will diffuse out over a few hours (or less) and the phage-containing liquid can be used in your experiment, or to prepare a new lysate whose phage all derive from the one that originated the plaque.

Plate lysates:  Lysates can be prepared in broth, by adding phage to growing cells, watching for the time when the culture clears because most of the cells have lysed, and pelleting out the cell debris.  This is a bit fussy to do, since clearing depends on having the right proportions of phage and cells.  A simpler method is to prepare a 'plate lysate', as follows.  Mix the liquid from a picked plaque or a small amount of a lysate with cells and top agar, and pour a lawn.  You want enough phage that the resulting plaques will be 'confluent' - will overlap just enough that very little intact lawn remains.  Once the plaques have formed, overlay the top agar with 5 ml of phage-dilution solution and leave for a few hours or overnight.  Half of the phage will diffuse into the liquid, and in the morning you just have to collect the liquid and add a few drops of chloroform to kill any cells.  These lysates usually have very high titers, because the cells in a lawn can grow to much higher density than those in a liquid culture.

Phage-resistant colonies:  
Sometimes, the area around an initially infected cell includes a cell that is genetically resistant to the phage due to a new mutation that blocks phage attachment or reproduction.  Such as cell (green in the diagram below) will be able to grow within the  area of spreading phage, and its descendants will form a visible colony within the plaque.

Turbid plaques:  One other phenomenon deserves mention, and that's the 'turbid' plaques formed when a 'temperate' phage infects a lawn.  Temperate phages are those that have a mechanism to enter a dormant state in host cells, where the phage genome is passively replicated by the cellular machinery, usually because it is integrated into the cell's chromosome.  Cells with such dormant phages ('lysogens', orange in the diagram below) are resistant to infection by external phages.  When a temperate phage forms a plaque, most infected cells lyse and produce infectious phage, but some form lysogens that grow and divide within the plaque.  Usually many such cells form causing the center of the plaque to appear cloudy ('turbid') rather than having visible colonies.

Questions about CRISPR-mediated phage immunity

Thursday's post described the hypothesis that bacteria might use gene transfer agent particles to inoculate other cells in the population with fragments of phage DNA, and outlined an experiment to test this.  Now I'm realizing that I need to know a lot more about the kind of immunity I should expect to see if this GTA-as-vaccine hypothesis is correct.

Simplistic outline of the experiment:
  1. Infect GTA-producer strain of R. capsulatus with phage under conditions where the infection is inefficient and few cells lyse.
  2. Remove cells and debris from the culture, to get a supernatant that will contain GTA particles and (unavoidably) some phage particles.
  3. Expose a new culture to the supernatant so cells obtain DNA from the GTA particles, again under conditions where successful phage infection will be minimized.
  4. Wash the surviving cells to remove phage (as much as possible).  Allow time for CRISPR formation if needed.
  5. Expose the cells to a titer of phage suitable for selecting resistant cells.  As a control, also expose cells not treated with GTA.  
  6. Plate to isolate colonies from surviving cells.
  7. Test the survivors for phage resistance.
  8. Compare the frequency of resistance in treated and control cultures.
  9. Test resistant colonies for CRISPR changes.

Things I should find out before I do the experiment:

1.  How efficiently do introduced DNA fragments give rise to CRISPR spacers?  If this efficiency is too low relative to the background rate of phage resistance, I won't be able to detect an effect.  This paper (Hynes et al. 2014, thanks to @AprilPawluk for pointing me to it) might let me estimate the  efficiency.  They exposed cells to a mixture of infectious and damaged phage (damaged by a restriction enzyme in the cell or by prior UV irradiation) at a multiplicity of infection (moi) of 0.1-0.2, and then examined the resulting confluently lysed lawns for phage-resistant colonies.  Unfortunately they only report relative changes in frequency of resistant cells (maxima 16-fold and 6 fold for restriction and irradiation respectively), but in their Methods they mention that the highest frequencies of resistance they observed were about 10^-6.  I don't know if this is for naive cells or pre-exposed cells, but even if it's for naive cells, the max frequency of CRISPR resistance I might expect is only about 10^-5.  This would not pose a detection problem, but it would limit the population-level benefits of the proposed vaccine system.

2.  What fraction of the survivors of a phage infection are genetically resistant, and what fraction of phage resistance arises by non-CRISPR mechanisms?  If most survivors are just lucky, then it might be a lot of work to identify the genetically resistant ones.  In the Hynes et al. experiments, all of the colonies were genetically resistant, and all had new CRISPR spacers.  However this might be quite different for different phages.  If most resistant cells have altered phage receptors rather than phage-specific CRISPR spacers, the effect of GTA-mediated CRISR resistance will be hard to detect.

3.  How quickly does CRISPR-mediated phage resistance arise after exposure to phage DNA?  I don't know.  Cells in the Hynes experiment might have had one or two lytic-cycle durations between being infected by the damaged phage and being infected by an infectious phage.

4.  What fraction of phage infections are abortive and thus could lead to CRISPR immunity to subsequent infection?  Inspired by the Hynes experiment, I can increase abortive infections by UV-irradiating the phage lysate.  (I know how to do this well from previous work.)

5.  How efficiently do phage spacers prevent phage infection?  April Pawluk (via Twitter) says they reduce infection by several orders of magnitude. In the Hynes work acquisition of a phage-derived CRISPR spacer enabled cells to form a colony in a sea of phage.

OK.  I have lysates of five sequenced R. capsulatus phages (from Dave Bollivar via Tom Beatty), and I have the R. capsulatus strain these phages were isolated on, as well as GTA-producing and recipient strains. Time to get to work!

Why GTA genes can't be maintained by 'selfish' transmission

Below is the line of reasoning showing that the genes responsible for producing GTA particles cannot maintain themselves or spread into new populations by GTA-mediated transfer of themselves into new cells.  I initially worked this out with a rigorous set of mathematical equations, but then realized that the problem was so glaringly obvious that math isn't needed.

The main GTA gene cluster is too big to fit inside a single GTA particle, so GTA particles can't transmit DNA that converts a GTA- cell into a GTA+ cell.  Some genes outside the main cluster are also required for GTA production.

But GTA particles can (and do) contain one or more individual GTA genes.  If a fragment containing a particular GTA gene is injected into a formerly-GTA+ cell that is now GTA- because it has a mutated version of this gene, the resulting recombination can restore the cell's original GTA+ genotype.

But these transfer events would not allow GTA+ cells to invade a GTA- population, or to maintain themselves in the face of loss of GTA function by mutation.  That's true for all known GTA systems, even in the simplest (imaginary) case where production of GTA particles requires only a single gene that could easily fit into a GTA particle, as illustrated below.  

Why?  Three factors together require that production of GTA particles reduces the total number of GTA+ cells in the population:

Problem 1:  GTA particles can only be released to the environment if the GTA+ producer cell lyses.  So each production event removes one GTA+ cell from the population.

Problem 2:  The GTA genes in the producer cell are not over-replicated as a phage genome would be, so each production event can produce at most one G+ particle (containing the GTA gene or cluster).  

If all steps occurred with 100% efficiency, problems 1 and 2 would allow, at best, replacement of the lost GTA+ cell with a new one created by GTA-mediated recombination.  But this would not maintain the numbers of GTA+ cells in the face of occasional loss of GTA genes by mutation or deletion.  Nor would it allow GTA+ cells to invade a GTA- population.

Problem 3:   Production of GTA particle production, transmission of their DNA to recipient cells, and recombination with the recipient genome are all likely to be at least moderately inefficient.  Here's a partial list of expected inefficiencies:
  1. Burst size:  Actual burst sizes are unknown, but packaging all the DNA in a R.capsulatus. genome would need 841 particles, which is much larger than typical burst sizes for DNA phages.  Capsid proteins may be limiting, since they would be produced from single-copy GTA genes rather than replicated phage genomes.
  2. Dispersion:  The GTA particles will disperse in the environment, and many will probably not find cells to attach to.
  3. Stability:  Lab preps of GTA particles are unstable in non-optimal storage conditions, so many particles will likely fall apart.
  4. Recombination efficiency:  Only one DNA strand enters the cytoplasm, and some DNA degradation is likely.  The highest observed transduction frequency is only ~4^-4, (theor. max: 1.2^-3) so recombination efficiency is probably only ~0.3.  Recombining in a novel gene will be less efficient than simple strand replacement
  5. Self-conversion:  Some G+ particles may attach to cells that are already GTA+.

Might GTA be a vaccination system for infecting phages?

My work at Dartmouth (to be described in upcoming posts) showed conclusively that genes encoding Gene Transfer Agents (such as the GTA system of Rhodobacter capsulatus) cannot be maintained by 'selfish' transfer of either whole GTA gene clusters or single GTA genes into GA- recipients.  Neither can the GTA genes be maintained by general recombination benefits that can arise when fragments of chromosomal DNA are transferred into new cells.  So, although 'gene transfer agent' does accurately describe one activity of these genes, it cannot be the activity for which they are selected.

The main obstacle to the maintenance of GTA genes, which applies to all the benefits is that any GTA+ cell that actively produces GTA particles cells must die, since cell lysis is needed to release their particles into the environment.  Another obstacle, applying to selfish transfer, is that GTA genes are not over-replicated during GTA production (and are not preferentially packaged), so each cell death can produce only one GTA+ particle. 

I presented these results at the Analytical Genetics conference last week, and asked the other participants if they could think of alternative benefits of producing GTA particles.  Sanna Koskiniemi from Uppsala University made the very interesting suggestion that GTA particles could serve as a syringe, packaging DNA fragments from a phage that's infecting the producer cell and transferring these fragments into other as-yet-uninfected cells, where they could trigger development of CRISPR immunity.

I love this idea and want to test it.  It doesn't overcome the cell-death obstacle, but it does overcome the selfish-transfer obstacle since a single producer cell could produce many particles of phage DNA from a single phage genome, and more if the phage genome is replicated.

One way to see if this could provide sufficient benefits to maintain the GTA genes is by simulation modeling like that I used to examine the recombination benefits.  This could clairfy the important factors that would need to be examined.

Here I want to start considering experimental tests of this hypothesis.

The ideal test would be to infect the GTA-producing strain with a phage, preferably under low-growth conditions where phage infections are often abortive.  (Luckily R. capsulatus produces most of its GTA under such conditions.)  Then some recipient cultures would be exposed to the GTA-containing culture medium (and some not, as controls), and then all exposed to a lysate of the phage.

"But wait!", you say.  "Won't the GTA-containing culture medium also contain some phage?"  Yes, probably.  I don't think there's any way to inactivate the phage particles without also inactivating the GTA particles, or vice versa.  We might be able to come up with either perfectly-abortive infection conditions (where infected cells don't produce any phage), or a cellular mutation that prevents phage production.  If not, we might have to combine the GTA-exposure and phage-infection steps.

"And won't any phage lysate also contain some GTA particles?"  Yes, probably.  But we could use a GTA- mutant as the host for lysate production.  Not the mutant that can't lyse, but the one with the main GTA gene cluster completely deleted.

What resources are available for this project?  First I checked with my GTA colleagues, who confirm that R. capsulatus does have a CRISPR-Cas9 system.  Then I asked if there were any well-characterized phage systems able to infect R. capsulatus.  Until quite recently the answer would have been 'No', but a recent paper reported the isolation and sequences of 4 R. capsulatus phages.  A Mu-like phage of R. capsulatus has also been characterized, but it did not form plaques on SB1003.

The report about the 4 new phages used a different host strain (YW1-derived, not SB1003), so the first thing I'll need to do is check whether they form plaques on SB1003.  Then I'll need to play around with infection and plating conditions...  My idea of fun!