Scholar’s Week Spring 2016

Last Spring the labrats put together a couple of posters and presented them at our local Scholar’s Week showcase. Both posters related to the DMAP1 project and give me a chance to see where we are at. I’m going to put up the poster slides here, mishing them together as they overlapped on some background stuff. True to form, I let the students play with powerpoint, illustrator and photoshop, to try and assemble a poster, but told them in advance I would be control freaky about requiring a very specific style, which I prefer (since the posters go up outside my lab and they MUST be clear).  I am always amazed by how my social media and computer savvy students don’t actually know how to put a poster together in powerpoint. It is always a brand new experience (and given the amount of time I spend using powerpoint, photoshop and the like, a useful one.)

When students come up with good design ideas (and some of them are really amazing) I do my best to make it so, together with them. So while the appearance of the poster and all the illustrations are mine (the fly pics typically have 50-100 component parts!) many of the layout and design ideas come from the students.

So, five students, two posters. Group 1: Genetic analysis of recovered lesions. Group 2: single embryo PCR analysis. What they had in common of course was the gene (DMAP1) and the model (Drosophila) and the male recombination scheme performed by students in Dr. Kathleen Fitzpatrick’s class at Simon Fraser University.  But I asked one group of students to focus on the genetic complementation tests that were used to define the limits of the deficiencies we obtained, and the other group to concentrate on the single embryo PCR (AKA getting DNA out of pieces of lint). Here are the abstracts for each poster. Poster 1: the genetics poster, has a blue dominant theme, while poster 2, the single embryo PCR poster, is green dominant (because, well, GFP, as you will see).

abstractsThe links to stem cell identity  and the immune system  are discussed here. Common to both posters was a description of the genomic landscape in which DMAP1 resides, and an outline of the male recombination scheme used to generate the deletions removing DMAP1 (and downstream flanking genes), more information about which can be found here.

So, a summary of the male recombination scheme carried out at SFU under Dr. Fitzpatrick’s supervision: this was included as part of the Single Embryo PCR poster (hence the green dominant theme).

male-rc-composite-slide

And a description of the genetic landscape where DMAP1 resides on the right arm of chromosome 2 (this slide appeared in both posters so there the blue version):

slide-4-poster-1-spring-2016

For the genetics poster, poster 1, we began by presenting evidence that DMAP1 might be essential, using transgenic RNAi.  A more detailed discussion about whether being essential matters or not can be found here.

transgenic-rnai-slidesHaving established that DMAP1 is likely essential using RNAi, we wanted to figure out how many genes were deleted by our male recombination-induced lesions. So we ordered a few overlapping deficiencies from the Drosophila stock center in Bloomington Indiana and performed complementation tests. Very cool, since some of the kids were in my genetics class at the time and so this was a practical application (“see? It does happen in the real world!”) The red lines in slide 5 below indicate two of those overlapping deficiencies (note that the red line indicates the DNA MISSING in the deficiency ) – one (BSC 402) takes out DMAP1 and MANY additional upstream and downstream genes, while the other (BSC 404) appears to break just upstream of a gene called “exu” (exuperantia). Slide 6 shows the results of the complementation test: all pairwise tests between K8-15 and 10-14 (the two male recombination deletions we made) and BSC 402 and 404 (the deficiencies of known limits). The bottom line is that deficiencies that remove DMAP1 and flanking genes fail to complement our male recombination mutants (10-14 and K8-15) whereas a deficiency that begins downstream of DMAP1 somewhere in the vicinity of exu complement our male recombination mutants. These data tell us that both lesions we made using male recombination are pretty large, and theoretically should end in the vicinity of exuperantia.

complementation-slidesBest laid plans, and all that. Sometimes, the molecular breakpoints of these deficiency fly stocks that one orders from a center like Bloomington have been mapped, sometimes not. Theoretically, Df(2R)404 has been molecularly mapped, and does in fact delete exuperantia. So based on the complementation data, our male recombination mutants should include exuperantia, right? So we’ve mapped the breakpoint, right?

Wrong. Lethality is one phenotype that results from a failure to complement. Lethality means there is an overlap between the known deficiency and our male recombination mutants. By sheer chance (and curiosity) one of Kathleen’s students tested the progeny from a cross between Df(2R)BSC404 and 10-14 or K8-15 for fertility. SURPRISE: progeny are both male AND female STERILE. So in fact, 10-14 and K8-15 do NOT complement Df(2R)BSC404. Take a look at the region deleted by this deficiency (the red bar highlighted in yellow on the map below):

df2rbsc404-map

Exuperantia is the tiny little blue triangle near the top of this figure on the left hand side. Our male recombination mutants in fact fail to complement Df(2R)BSC404, because progeny heterozygous for this deficiency and either K8-15 or 10-14 are sterile. Sterility, from an evolutionary point of view, is just as bad as lethality, and therefore is just as much a failure to complement. So what is exu? What does it do? Is sterility a predictable phenotype?

Exuperantia has an illustrious pedigree. It was isolated as a maternal effect gene necessary for the normal anterior-posterior patterning in the Drosophila embryo. Exu was one of an elite suite of genes isolated and studied in the mid 1980’s by Trudi Schüpbach and Eric Wieschaus in a screen to study pattern formation in development. This work revealed a highly conserved genetic determination of body patterning and opened up the fascinating field of developmental genetics. Oh yes, and resulted in a Nobel Prize in 1995.  Bottom line: if the mothers are homozygous for loss of function mutations in exu, the embryos fail to develop, and they show anterior patterning defects. How does the exu gene product do this? Presumably by controlling some of the other mRNA’s that mum packs into the cytoplasm of her unfertilized egg, providing the egg with patterning coordinates (head end, tail end, dorsal side ventral side etc.)

exu-fcn-flybase

A number of mutations in exu have been isolated since then. Most are female sterile, one is male sterile. A couple are lethal (but we have determined that at least one of those lethals is due to a second side hit on the same chromosome). So, yes, theoretically, male and female sterility is a reasonable phenotype for our male recombination lesions, and tells us that they both extend at least out to exuperantia.

Buggah.

Can we confirm these data using PCR? Certainly we know our lesions take out DMAP1 and do not disrupt the region upstream (see here), but how far downstream do these lesions extend? More primer design for the students,  targeting genes in the vicinity of exu.

slide-7-poster-1-spring-2016And our results? Using DNA from adults means using balanced flies, with the wild type genetic region on the balancer. (Remember what a balancer is? No? Explanation to follow). Thus, the data below show the results using a primer to the P element which we know is still present upstream of DMAP1 (at least half of it is) and gene-specific primers. P-out is the P element primer, in combination with the upstream gene CG33785 we have a product, but nadda in combination with anything down stream, including exuperantia.slide-8-poster-1-spring-2016Now, these data make a bold assumption, that so much DNA has been deleted in  10-14 or K8-15, that the region between the P element and the exu gene-specific primer is now small enough to be spanned by garden-variety PCR (usually a limit of about 2 to 3 kilobases of DNA). What if that’s wrong? Also, what if the priming site in the P element isn’t there anymore, for down stream amplification? Really the only way to find that out is to try PCR on single K8-15 or 10-14 homozygous individuals. And as those are dead, we may be stuck.

Or are we? We put both K8-15 and 10-14 over a balancer chromosome that also carried a transgene expressing the Green Fluorescent Protein (GFP). What’s a balancer? Recall that if you want to maintain a recessive lethal mutation in a population of individuals, meiosis will eventually purge the population of the mutation – lethality rather interferes with fitness, after all. Drosophila geneticists have developed a useful tool called a balancer chromosome, which has essentially all the right DNA in it, but not necessarily in the right order, having been scrambled somewhat by radiation (which reminds me of the famous Morecambe and Wise sketch with Andre Previn: “I am playing all the right notes, just not necessarily in the right order” – see around 11:00 in the sketch). The balancer is usually marked by a mutation that has a dominant phenotype (curly wings, stubbly bristles, messed up eyes etc) that is recessive lethal. That way you can instantly see if your stock is balanced. What this means is that during meiosis, when a chromosome with a lethal mutation on it attempts to pair with a balancer homolog, well, it can’t, and all recombinant products are lost. Thus, the population consists entirely of individuals that are heterozygous for the balancer and the chromosome with the desired mutation. A marvellous tool, so far unique to Drosophila.

So we balanced our male recombination mutations over a balancer with GFP on it, which allowed us to do two things. First, we could identify the lethal phase, or when homozygotes for K8-15 and 10-14 died. Second, we could isolate individuals, dying, to be sure, that were homozygous for the lesions, which would be GFP minus, and extract their DNA for PCR analysis.

slide-4-poster-2-spring-2016In fact the graphic above is not entirely accurate: the GFP -/- individuals never made it to adult hood, as implied. They died as first instar larvae – fresh out of the egg.

single-emb-pcr

Panel C in slide 5 above shows a first instar larva that is homozygous for 10-14 or K8-15 – both lesions behaved the same way. Slide 6 shows how we used GFP fluorescence to select against all the genotypes we did not want, and select only those that did not fluoresce. Those larvae were placed individually into eppendorf tubes for DNA extraction and subsequent single embryo PCR. We tested a number of genes in the vicinity of exu – here are some data for CG13437: clearly it is absent from the male recombination lesion 101-14.

slide-7-poster-2-spring-2016

I can tell you we also tested using primers to exu, and to galla-1, which is even further away.  The Galla-1 results are puzzling because Galla-1, which may play a role in chromosome segregation through establishment of sister chromatid cohesion (PMID 25065591), is supposed to be lethal (same paper). But our “complementing” progeny from a cross between Df(2R)BSC404 (removes galla-1 completely) and our male recombination lesions (also remove Galla-1 – enough of it to be considered gone based on where the primers target) are alive. Sterile, but alive. What could this mean? Who knows.

Buggah, buggah, these are large deficiencies we made, when what we really wanted were small lesions confined to DMAP1.

Do we have ANY idea where the break point is? Both K8-15 and 10-14 are viable but sterile over Df(2R)BSC404.  Are there any essential genes, in terms of viability, included within the genomic territory defined by this deficiency? Aside from Galla-1? Yes, but perhaps we have to stick to the brut force single embryo PCR approach at this point. The next victims in my lab will design more primers getting further and further away from exu, Galla-1…until we get a PCR result that tells us a gene is actually present in our male recombination induced lesions.

We will have to map these breakpoints. We want to know how big these lesions are. The next step is to use them as known deficiencies over which to carry out a “local hop” – try to get the P element just upstream of DMAP1 to move out of its current location and into DMAP1, hopefully knocking the gene out. K8-15 and 10-14 may well be huge, but they are much smaller than the stock deficiencies available, like Df(2R)402 or 404. They may yet be useful. Fingers crossed.

Here are the concluding slides for the two posters my students presented last spring, and a couple of shots of students by their posters.

conclusion-slides

 

 

DMAP1 mutations: PCR analysis

So all our putative DMAP1 mutants are stably balanced (if lethal) and floating balancers (if viable – the balancer chromo is still in the stock but homozygotes abound). At this point, it becomes possible to purify genomic DNA from them and see how much we can find out about the molecular nature of any lesions in the region using PCR (I make the bold assumption all readers know how PCR works. If not, try this).

First of all, we need to generate a map of the region, and design primer pairs that are likely to be diagnostic of any changes in the DNA. One very important primer will anneal to the inverted repeat of the engineered transposable element that we used as a starting point for our male recombination scheme. There are two primers of use here: P-OUT which anneals to the 31 base pair inverted repeat sequence in such a way as to point OUT of the P element (3′ end of primer directed away from the internal P element sequence) and P-IN, simply the complement of P-OUT and of course, pointing INTO the P element.  By “pointing” into or out of, I mean, of course, that the Taq polymerase will extend from the 3′ end of the primer into or out of the P element.

So here is a map of the primers I’ve designed so far.

DMAP1 primer map

Recall that the original engineered P element is inserted into the 5′ UTR of the upstream (on the left) gene CG33785/6 (UTR = untranslated region – grey in the Flybase gene diagram above). The male recombination scheme is designed to select for events in which the P element partially excises from this location, taking with it material in the direction of DMAP1 – we want to leave the upstream region intact. If all goes according to plan, a successful event will retain the upstream (5′) portion of the P element, but should lose the downstream (3′) end, plus hopefully a nice chunk of the DMAP1 gene and nothing else. How MUCH DMAP1 DNA downstream of the P element is retained should be possible to determine by seeing if the downstream priming sites are retained.

Of course the best way to do this is to carry out a Southern analysis.  Cut up the DNA with a well-chosen restriction enzyme (well-chosen based on the sequence we know), fractionate the digest in a gel by electrophoresis, blot the pattern of fragments onto a membrane, incubate the membrane with a DMAP1 cDNA probe, labelled in some way to make it possible to visualize (usually radioactivity) and analyse the pattern of bands. Southerns are still important experiments – you see them commonly described in four point font in the materials and methods section of knock-out mouse studies. But they are not simple experiments, and they are not cheap, so PCR it is for us at SmallScienceWorks.

So without further ado, here are the results. Note that we did these PCRs at the same time as the complementation tests, so some of these data make more sense in hindsight, which I’ll discuss a bit later. For the homozygotes, I selected males only (mated females might have eggs from balanced sibs that will yield false positives – remember the region is wild type on the balancer)

In Figure 1, top row of lanes, we tested all the viable isolates and w[1118] (which has no P element and is wild type for the region) using gene specific primers in DMAP1. The bottom row, on the other hand, tests for the presence of the DOWNSTREAM (or 3′) end of the P element in all the isolates, by using P-OUT together with a series of DMAP1 gene specific primers.

Gel 1a

From these data, top row, which tests for gene structure in the mutants that are viable (as homozygotes) you can see that there appears to be no disruption in the region covered by all these primers (which is pretty much the whole gene). The band sizes for the mutants are indistinguishable from w[1118], which is wild type (the gel is frowning a bit – the bands are distorted on the edges – not a big deal). Were we disappointed? Yes. NO! Of course not! We are dispassionate scientists, who want what we get, not get what we want. Buggah.

From the data on the bottom row though, you can see that the P element is absent in all mutants EXCEPT K14, and of course in still present in GS10389, the original P stock used in the male recombination scheme (and therefore serving as a positive control). You may note some laddering in the bottom row, right hand side. That’s OK, that is non specific product, resulting from some weak false priming of the primers that shows up after 30 cycles.

The data shown in Figure 2 are even more disturbing, at least the top row is. The NGFPF primer is a forward primer that anneals to the very beginning of DMAP1, starting at the ATG. The subsequent lanes are testing for amplification across the P element insertion site. CG33785 is upstream of DMAP1 – remember the P element is inserted into the 5′ UTR of this gene.

The bottom row is another test for the downstream or 3′ portion of the P element.

Gel 2a

So the data in this gel tell us that the gene structure is probably completely unaltered in the homozygous “mutants” – I use inverted commas because they are scarcely looking like mutants at all. Any lesions, if they exist, must be so small there is no difference in band size when the “mutants” are compared with w[1118], which remember, is wild type for this region. Hmm.

The bottom row tells us what we already know. The downstream portion of the P element is absent in all mutants except K14. Perhaps, in the homozygotes, we had a perfect excision of the P element, plus a recombination event further down the chromosome, to put the brown eye color mutant onto the same chromosome. Who knows.

So what about the P element upstream of DMAP1? According to the scheme, the 5′ portion of the P element should still be there, though this is hard to believe with the homozygotes (top row Figure 2) – PCR across the insertion site shows no size difference with w[1118]. But of course, we did the PCR to prove it to ourselves. Here it is, Figure 3.

Gel 3a

So all peculiarities solved, right? Everything makes perfect sense, the end.

Alas no. According to the top row of data, the upstream portion of the P element is present, (as it should be in theory) EXCEPT for 20-13 homozygotes, and 22-13/SM6a heterozygotes. Great. There is a little bit of false priming in the negative lanes but the upshot is, for three out of four homozygous viable “mutants”, the upstream or 5′ portion of the P element is there, and yet, not there, when PCR across the site is performed. The P element is there and not there. How very Heisenberg.

Huh? I didn’t get all that. And I wrote it.

For the mutants that are homozygous viable, the upstream portion of the P element is present (except in 20-13). The smallest amount of P element that could be left is 31 bp – the annealing site of the P-OUT primer. Unlikely – if the priming site is there, a chunk of the P element must be there. So when you do PCR using primers on either side of that chunk, then the band should be BIGGER than it is in w[1118] which doesn’t have the P element. BUT all the bands representing PCRs across the site are identical in size to the w[1118] stock. BUT the upstream P element is still there. According to the PCR with P-OUT and the upstream gene CG33785/6. So for these homozygous viable mutants, the P element is there, but not there. See? It is a headache inducing contradiction. But there must be a solution, because this is real matter, this is Nature, this is real DNA. We just have to figure it out.

Here is a summary of the data in Table 1:

Table 1

Note K8-15* – this is a new mutant, generated by my colleague Kathleen Fitzpatrick this year up at Simon Fraser University in BC Canada. It looks good – fails to complement 10-14, which fails to complement a deficiency for the region, and it behaves just as 10-14 in the PCR assays as well.

Now we know from the complementation tests that of the lethals, 22-13 and K-14 complement deficiencies for DMAP1 obtained from the Bloomington stock center. So these are lethal hits elsewhere in the genome, and therefore should be chucked (which is hard because Drosophila geneticists tend to be hoarders of genotypes). Of the viables, 20-11 shows a weak failure to complement with 10-14 so I am inclined to keep it, in case it turns out to be a hypomorph (partial loss of function). Though, like the other viables, it shows this bizarre P element there-and-not-there molecular phenotype. BUT: there is an outside chance that these viables might in fact be duplications for the region, which, if you examine the original paper (PMID: 8978050) indicates that duplications can occur with approximately equal frequency to the deletions. We might be able to detect these by PCR – a forward primer pointing downstream in DMAP1 and a reverse primer upstream of the forward primer might amplify a product, if a local duplication as occured. Worth a try.

We have no idea about the gene structure for DMAP1 in the lethals. These stocks are balanced, and so primer pairs NOT involving P-OUT will amplify from the wild type sequence present on the balancer. I need homozygotes for whatever the lesion is in 10-14 (or K8-15) in order to test for gene structure using solely gene specific primers. I tried rebalancing the lethal 10-14 over a CyO-GFP balancer and looking for GFP-minus larvae but saw none, suggesting this genoytpe has a very early lethal phase (dying as an embryo perhaps, or a first larval instar, where the GFP doesn’t show up).

So plan B: note from the complementation tests that 10-14 complements the original P strain GS10389. If I can purify DNA from flies transheterozygous for the P element-bearing chromosome, and the 10-14 deletion chromosome (or K8-15 for that matter), I might have a chance at mapping the gene structure, using primers that span the insertion site. For the GS10389 chromosome, the region is too large for conventional PCR (the engineered P element insertion is about 7000+ base pairs of DNA – conventional PCR is good to about 3000 bp), and so if I find a primer pair that anneal to the remaining DNA, I should be able to amplify a product. If that happens, I will certainly cut out the amplified band from the gel, extract the DNA and clone the fragment, and stick it into my freezer until I can find some $$$$ to sequence it. At that point, we will know the precise molecular nature of the lesion. Hurrah!

 

DMAP1 Fertility Tests

So my colleague Kathleen and her inexhaustable supply of undergrads at Simon Fraser University in B.C., Canada, have generated seven potential DMAP1 lesions, using the male recombination screen described in the last post. Recall that in this screen, a P transposable element very close to the upstream gene CG33785 was induced to transpose in the male germline, hopefully generating a deletion in the direction of DMAP1, and thereby recombining a recessive brown eye colour mutation onto the same chromosome. Note here that four out of seven potential lesions produce viable brown-eyed (and therefore homozygous) flies. They look normal but are they really? In Figure 1, each potential mutant that produces homozygous progeny is tested for fertility. For each strain, four crosses, with virgin male and female parents, 3-4 of each sex (i.e, as close to identical conditions as possible). First, balanced males and females are crossed to each other (grey and black representing Cy males and females respectively – remember Cy is the dominant curly wing mutation that marks the SM6 balancer). Then, balanced males are crossed to homozygous females (which are dark blue in my colour scheme) – this tests the homozygous female fertility. Then the other way around: homozygous males (light blue) crossed to balanced females – this tests the homozygous males’ fertility. Finally, homozygous males and females to each other.

Fertility tests

The total number of flies for each cross exceeded 100, but I expressed the proportions of each genotype as a percentage of 100 in this graph. For balanced parents crossed to each other, the ratio of balanced to non balanced progeny (or blue shades to grey shades) should be 0.5.  (Think A/a X A/a, where A is Cy, and Cy/Cy is lethal. Half as many a/a to A/a, or 1/3 : 2/3). Note that doesn’t really happen except for 12-14 (and that ratio turns out to be 0.48).  For the two middle crosses, heterozygous males to homozygous females and the other way around (think A/a X a/a) half the progeny should be homozygous, and half should be balanced, or 1:1.  So you should see as much grey/black and light/dark blue for those crosses. Do you? Of course, homozygotes crossed to each other are depicted solely in the blue shades.

Clearly two of the potential mutants show sterility defects: 20-13 is male AND female sterile (no progeny at all for the second, third and fourth crosses), and 12-14 looks male sterile. Incidentally, in the very first post about this project, I yanked the ovaries out of 20-13/20-13 females, (look here), and asserted the testes didn’t look quite right either, so that’s consistent so far. (Could still be something in the background, but relish it for now!)

As for 20-11 and K2-13, the second and third crosses don’t produce quite the right ratios.  In addition, 20-11/20-11 parents produced approximately 70% fewer progeny than 20-11/SM6 parents, and for K2-13 that reduction was around 50%. So there might be viability and fertility issues for both lines.

In summary, 20-13 is homozygous male and female sterile, 12-14 is male sterile, 20-11 and K2-13 homozygotes may be semi fertile and perhaps even semi viable (particularly 20-11). But are these all mutations in the same gene? Stay tuned.

Screening for DMAP1 deletions

Here is a description of the male recombination screen used to target DMAP1  which my colleague Dr. Kathleen Fitzpatrick (Simon Fraser University (SFU) in Burnaby BC Canada) and I wrote together. Dr. Fitzpatrick used this screen as a part of her undergraduate genetics lab teaching program, and so this work is discussed largely by her (I made the pretty pictures). An application of the Awesome Power of Undergrads in the service of a very tricky screen!

In our class (Bisc 302W) at SFU, we spent several semesters screening for a directed deletion of the gene DMAP1 in Drosophila melanogaster. We used a P transposable element located between DMAP1 and its near neighbour CG33785, on chromosome 2. The P element is called GS10389,  but I called it P* in the class, so I’ll refer to it that way here as well. Note here that P* is closer to the gene CG33785 than it is to DMAP1.

There are two main requirements for P elements to move in the genome: first: an enzyme called transposase and second, special DNA sequences on either end of the P element called inverted terminal repeats. The transposase enzyme recognizes those repeats, and can cut the P element out of the original position and insert it into a new position somewhere else in the genome. P elements tend to have insertion preferences: places where they are more likely to insert and places where they won’t insert. But there is no way to predict exactly where a P element will go. In this sort of work, you mobilize a P element – allow it to move – and then you have to select for the result you want. You can’t just make it do exactly what you want.

The majority of transpositions are precise – we call them clean (or perfect) excisions. Sometimes transposition is not neat and tidy; sometimes it’s a bit messy – the element is not cut out and moved discretely, but some DNA to one side or another of the element is cut out too. This is referred to as imprecise excision. For our purposes, we are very interested in those specific imprecise excision events that also include a crossover event with a homologous sequence, because in this way, we might be able to select the direction in which the deletion takes place  (and remember, P* is closer to a gene we do NOT want to disrupt). In 1996, Preston et al (PMID: 8978050) published the results of their analysis of crossovers (recombination events) induced by P elements. They found a crossover rate of about 1% of the chromosomes tested (though it varied with temperature) and – here’s the important thing – deletions were generated from the insertion’s original site to one side or the other of the P element in some of the crossover chromosomes. The direction of the crossover reflected the direction of the deletion. Thus, if you induce a P element to move, and the element starts out near a gene you are interested in, careful selection any recombinants that result may net you some deletions of your gene. The mechanism is kind of complicated – the P element sequences located on sister chromatids try to pair with each other, forming a complex structure, the resolution of which may generate the deletion we need. Take a look at Movie 1 (and many thanks to Dr. Bill Engel for letting us use it).

 

So, here is a strange thing about fruit flies that turns out to be very useful in this context. Everybody knows that during meiosis, homologous chromosomes pair and undergo crossing over during Prophase I. This is one of those very important generators of variability upon which our evolutionary history depends. But, in male fruit flies, for reasons I think are still not well understood, there is NO meiotic recombination. Very strange. In fact, reduced rates of recombination associated with heterogametic organisms – those with different sex chromosomes like X and Y – have been observed since the early part of last century – here’s a recent paper using Zebrafish that describes it well (PMID: 11861568). Male fruit flies represent an extreme example with no meiotic recombination at all, but that turns out to be very helpful to us, because if we do our directed deletion experiment with male flies, any crossovers that result are almost certainly due to the P element “attempting” to move and making a useful mistake (for us), as opposed to a general recombination event we can’t control.

But how do we set this up so that we can select for those aberrant male recombination events that move in the direction of DMAP1? We really want to make a deletion in DMAP1 and only DMAP1. How can we distinguish between deletions in the left or right direction?

The trick is to use other genetic markers – mutations in other genes, to the left and to the right of DMAP1 that can be recombined onto the chromosome at the same time, during the same recombination event in the male germline that generates the deletion when the P element is mobilized. So this is what we did in our class (Figure 1): we crossed flies with the P element beside DMAP1 (P*) to flies with eye colour mutations on chromosome 2, and a source of transposase enzyme on chromosome 3. We collected male offspring that combined P* with the transposase source. Then we crossed those males to females with the recessive eye colour mutations cinnabar (cn) and brown (bw) on both homologues of chromosome 2.  Here’s the important thing: these eye colour genes are located on either side of DMAP1 and P*, so ultimately, we can select for deletions in one direction or the other by selecting for one eye colour phenotype or the other (this is also depicted in Movie 1). The males were heterozygous for the recessive eye colour mutations, and therefore had wild type (red) eye colour, and the females were homozygous for the mutant forms (which combine to yield a white eye colour – a subject for a later post; just accept this curiosity for now). Therefore any crossover in the males could be detected in the offspring as either mutant for cinnabar but normal for brown (bright orange eyes) or mutant for brown and normal for cinnabar (brown eyes).

DMAP1 male RC 04

Most of the progeny will have either normal red eyes (one wild type allele of each gene) or white eyes (mutant alleles of both genes). Hopefully Figure 1 will help you work through the chromosome shuffling that’s going on. Figure 2 is what it looks like when the crosses are written out in fly language:

Cross scheme

Since we wanted deletions in the direction of DMAP1 (and not CG33785) we kept any brown-eyed males and crossed them individually to a stock of flies containing the SM6a balancer chromosome (having the dominant Curly (Cy) mutation and the recessive cn mutation, along with a bunch of rearrangements that prevent further recombination, thus allowing us to maintain potentially useful deletions). Brown eyed males have a recombinant chromosome that is likely to have a deletion towards, hopefully into, possibly all the way through, the DMAP1 gene. We had to cross each brown-eyed male individually because theoretically, each male arose from the union of a paternal gamete (sperm) in which the desired recombination event took place, and an egg from the white eyed, cn,bw mother. Every F2 male, in other words, represents a gamete from the F1 father, in which the P element was induced to recombine.

Preston et al. found that about 1/3 of the recombinant chromosomes they tested had deletions. The deletions ranged in size from just a few base pairs to more than 100,000 base pairs, but the majority fell within about 2000 base pairs. This is good because the DMAP1 gene is about 1800 base pairs long, and we hope to delete most or all of the DMAP1 gene without deleting additional genes. I set up the cross to make the F1 males, and gave those males to groups of students in several successive Genetics Lab semesters. Students crossed those males en masse to cn, bw females, and then sorted through the F2 progeny for brown eyed flies. Those males, when found, were crossed INDIVIDUALLY to females with the multiply rearranged balancer for the second chromosome that was also marked with cn, and in the next generation, the F3, the recombinant chromosome bearing the brown allele and P* was recovered and balanced as a genetically uniform population of flies, or a stock. If the induced recombination event caused a lethal mutation, then the stock should remain balanced, all flies showing curly wings, with no brown-eyed homozygotes present. If the event produced a mutation that was not lethal, then brown-eyed homozygotes should show up in the stock.

There’s a lot to keep track of in this screen. Lots of different markers and mutations, understanding what balancers are and what they are good for, why some matings are en masse (lots of males and females together in a bottle or vial) and why others have to involve single males. After several semesters, we generated seven stocks carrying independently induced recombinant events that hopefully have deleted into DMAP1. Table 1At first glance (Table 1), we see that some stocks are lethal, and some are not, which could mean that DMAP1 encodes an essential function, and we have possibly generated an allelic series, ranging from complete lethality due to a large deletion that removes most or all of the coding region (a null), to perhaps a small change that simply reduces the expression of the gene (a hypomorph). But lots of other, less appealing results are also possible – the generation of an off-site hit in some other gene (that happens to be essential), or a complex rearrangement that will be awful to try an analyze. We have lots of work to do…so we had better get started!