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Meiotic recombination is triggered by programmed DNA double-strand breaks (DSBs), which are catalyzed by Spo11 protein in a type II topoisomerase-like manner. Meiotic DSBs can be detected directly using physical assays (gel electrophoresis, Southern blotting, and indirect end-labeling) applied to samples of genomic DNA from sporulating cultures of budding and fission yeast. Such assays are extremely useful for quantifying and characterizing many aspects of the initiation of meiotic recombination, including the timing of DSB formation relative to other events, the distribution of DSBs across the genome, and the influence on DSB formation of mutations in recombination factors and other gene products. By varying the type of gel electrophoresis and other parameters, the spatial resolution of DSB analysis can range from single nucleotides up to whole yeast chromosomes.
To detect meiotic DSBs in S. cerevisiae, genomic DNA is extracted from synchronously sporulating cultures, then digested with restriction enzyme(s) as necessary to yield an appropriate sized fragment. Subsequently, these DNA fragments are separated by gel electrophoresis, and both the parental length (unbroken) DNA and the DSB fragments are detected by Southern blotting and indirect end-labeling by hybridization to an appropriate probe (Fig. 1).
Meiotic DSBs do not occur randomly throughout the genome. Instead, DSBs (and the resulting recombination products) form preferentially within small (≤2 kbp) regions called hotspots (1). DSBs show specificfeaturesof distribution according to the resolution of the methods used to detect them. When observed at the resolution of a whole chromosome (100’s to 1000’s of kbp in S. cerevisiae), alternating hot and cold domains are observed that are typically on the order of 50–100 kbp wide (see Fig. 2 for an example). When examined at the resolution of individual genes (i.e., examining ~5–20 kbp at a time), individual hotspots can be observed that are separated by several kbp of DNA in which few, if any, DSBs are formed (Fig. 3A). Most hotspots in S. cerevisiae are located within transcriptional promoter regions (2). Mapping at yet higher resolution reveals that each hotspot consists of multiple DSB sites clustered within regions of ~75–250 bp (Fig. 3B). Finally, mapping at the resolution of individual nucleotides reveals that Spo11 cleaves the DNA to yield a two-nucleotide 5′ overhang and that some positions within hotspots are cut more frequently than others, although no obvious DNA sequence preference has yet emerged (3–6).
Because there is no single method for separating and detecting DNA fragments across all of the size ranges outlined above, it is necessary to choose DSB mapping strategies appropriate to the purpose of the experimental study. Here we describe procedures for DSB analysis at four levels of spatial resolution, which we define as chromosome level, medium resolution, high resolution, and nucleotide level mapping. These differ from one another primarily with respect to DNA sample preparation and the method of gel electrophoresis (pulsed-field, conventional agarose, polyacrylamide, or sequencing gel, respectively). These protocols are modified from established methods (5, 7, 8).
In many studies, the position and the frequency of meiotic DSBs are measured using rad50S or sae2Δ mutants, which are deficient in a process of endonucleolytic release of covalently attached Spo11 from DSB ends (5, 6, 9, 10). These mutants accumulate unrepaired DSBs, making observation of the normally transient meiotic DSB much easier. The frequency of DSBs varies from one hotspot to another and, in general, DSB frequencies measured in rad50S or sae2 strains correlate well with overall recombination frequencies in corresponding RAD50+ or SAE2+. However, it is important to note that DSB frequencies are substantially reduced in rad50S or sae2 mutants in regions where replication is experimentally delayed (8), and recent genome-wide studies demonstrate that DSBs in certain regions are specifically underreported in rad50S-like mutants (11, 12).
To obtain well-synchronized meiotic samples, cells are precultured in presporulation medium (SPS), and then transferred to sporulation medium (SPM). In the SK1 strain background, the DSB frequency reaches maximum levels about 5–6 h after transfer to SPM. Premeiotic samples are harvested at 0 h (i.e., immediately upon transfer to SPM), and meiotic samples are harvested at appropriate time points thereafter (seeNote 1).
The frequencies of meiotic DSBs are low, so it is important that genomic DNA is extracted gently to avoid mechanical shearing. Excessive shearing generates a high background of hybridizing signal in lanes from the gel electrophoresis, and can obscure weaker DSB signals and/or make DSBs difficult to quantify. DNA samples in solution should be mixed gently, and vigorous pipetting and vortexing should be avoided.
To observe DSBs at different levels of spatial resolution, the lengths of the parental and DSB fragments need to be considered, and an appropriate probe needs to be chosen for indirect end-labeling of Southern blots. Parental and DSB fragments should be well separated by gel electrophoresis in order to accurately estimate DSB frequencies. It is recommended that probes are designed within open reading frames so that the region to which the probe hybridizes is unlikely to itself be a site of DSB formation. Additional specifications are discussed in the individual methods sections below.
Restriction enzymes; electrophoresis power supply; agarose gel electrophoresis apparatus suitable for running 30 cm-long gels; agarose; TBE buffer (see Subheading 2.2.2), 6× loading buffer (0.25% bromophenol blue, 0.25% xylene cyanol FF, 15% Ficoll PM400); 10 mg/mL ethidium bromide solution.
Whatman 3MM filter paper; transfer apparatus (BioRad Trans-Blot® SD Semi-Dry Electrophoretic Transfer Cell, or equivalent); Nylon membrane (uncharged, GeneScreen™ Hybridization Transfer Membrane (PerkinElmer) or equivalent); UV crosslinker; 20× SSC (see Subheading 2.2.3).
High voltage electrophoresis power supply; sequencing gel electrophoresis apparatus; sequencing gel plates (30 × 40 cm); plate coating reagent (Sigmacote); shark tooth combs and spacers (0.4 mm thick). See Subheading 2.4.1 for acrylamide gel solutions and loading buffer.
UV crosslinker; 20× SSC (see Subheading 2.2.3); 35 × 45 cm blotting paper (BioRad); Whatman 3MM filter paper; electro transfer apparatus (TE 90 - GeneSweep™ Sequencing Gel Transfer Unit (Hoefer Scientific Instruments)).
Taq polymerase, 333 μM dNTP mix without dCTP, [α-32P] dCTP (6000 Ci/mmol), Taq amplification buffer, 50 mM MgCl2, probe template DNA (see Subheading 2.2.4), 20 μM PCR primer b (Fig. 4).
This section is based on (13) with minor modifications.
This method is suitable for observing whole chromosomes and DSB fragments of 0.2–2.0 Mbp after separation by pulsed-field gel electrophoresis (PFGE). It is necessary to prepare high molecular weight DNA under conditions of gentle cell lysis to prevent any mechanical DNA breakage. This is achieved by embedding cells in low-melting point agarose as soon as they are collected and performing all enzymatic reactions (preparation of spheroplasts, lysis and DNA purification) in the resulting DNA “plugs”. Enzymatic degradation of chromosomal DNA is prevented by maintaining high concentrations of EDTA, which inhibits nucleases, at all steps of DNA plug preparation. For indirect end-labeling on Southern blots, a good choice for probe is often the open reading frame closest to one extremity of the chromosome of interest. A unique, unrepeated gene should be chosen.
This method has been described in (14).
DSB mapping using separation by conventional agarose gel electrophoresis is suitable for observing DSBs across ~2–15 kb regions, with a resolution of ~50–100 bp. For accurate quantification, parental fragments of less than 10 kb are recommended so that transfer during Southern blotting is efficient. Restriction enzymes can be chosen as appropriate to give good spatial separation between the DSB fragments and the parental fragment (see Fig. 3A). We generally use 0.5–1 kb DNA fragments as probes, which can be efficiently labeled by random priming. Ideally, the probe should hybridize to sequence close to one end of the parental size restriction fragment (Fig. 3A).
Parental and DSB bands can be visualized using a Phosphoimager and quantified using ImageQuant (Molecular Dynamics) as described in (7). The frequency of DSBs is calculated as the percent of radioactivity in DSB fragments relative to the total radioactivity in the lane (i.e., parental and DSB fragments). Appropriate exposure times can be selected to allow accurate determination of the intensity of the parental band (i.e., within the linear response range of the phosphor screen). Background signals are often present in the lane because of random shearing of genomic DNA during preparation of samples, etc; these background signals should be subtracted.
DSBs can be detected at higher resolution (±10~20 bp) using polyacrylamide instead of agarose gel electrophoresis (see Fig. 3B). We generally obtain better resolution and signal strength using denaturing (6% polyacrylamide containing 8 M urea) rather than nondenaturing gels. Good resolution is achieved when DSB fragments are 150–300 bp long. To provide sufficient spatial separation from DSB fragments of this size, parental fragments should be 500–1000 bp. Restriction sites should be chosen with these size ranges in mind. The size of the DNA probe fragment should be 100–200 bp, which is often difficult to label to sufficient specific activity by random priming. Therefore, we use PCR to amplify the probe fragment in the presence of [α-32P] dCTP.
|333 μM dNTP mix, without dCTP||0.5 μL|
|[α-32P] dCTP (400 Ci/mmol)||5 μL|
|20 μM Forward Primer*||0.625 μL|
|20 μM Backward Primer*||0.625 μL|
|10× Buffer||1.25 μL|
|50 mM MgCl2||0.625 μL|
|Taq polymerase (5 U/μL)||0.25 μL|
|Template DNA fragment||5 ng|
|Water||to 12.5 μL|
In order to detect DSB sites at single nucleotide resolution, DSB fragments must be separated on a sequencing gel. Before carrying out this type of analysis, it is recommended to first determine the position and frequency of DSBs at the locus of interest by performing high resolution mapping (Subheading 3.4). Based on the result of this mapping, a restriction enzyme that cuts 150–200 bp from DSB site can be chosen, along with primer sets to amplify probe and template for sequence standards (see Fig. 4).
Meiotic DSBs made by Spo11 have 2-nucleotide 5′ overhangs (seeNote 18). In rad50S or sae2Δ mutants, Spo11 remains covalently attached to the 5′ ends. Even with extensive proteinase K treatment, oligo-peptides of Spo11 still remain covalently attached to 5′ DSB ends, which causes the 5′-terminal DNA strand to migrate slower on sequencing gels and thus makes it difficult to map the 5′ ends accurately (3–6). To circumvent this problem, 5′ ends are mapped indirectly by filling in the 3′ ends with a DNA polymerase that does not add extra nucleotides. After the fill-in reaction, the ends of the DSB fragments will be blunt and the filled-in 3′ ends will match the original 5′ ends (see Fig. 5). The filled-in 3′ ends and untreated (original) 3′ ends can then be detected side by side on the same gel by probing a Southern blot with a strand-specific probe (Fig. 5). Sequence standards are prepared by linear amplification in the presence of dideoxy nucleotide triphosphates (ddNTPs) with a primer that corresponds to the end of the sequence cleaved by the restriction enzyme (Fig. 4, primer a). This method is based on (5, 15).
After defining DSB sites at single-nucleotide resolution by this method, it is advisable to confirm the accuracy by mapping the DSB ends on the other side of the hotspot. This is accomplished by repeating the procedure using an appropriate restriction digestion and probe, as diagrammed in Fig. 4 (probe B, primer c, primer d, and RE2).
|5× buffer||10 μL|
|25 mM dNTP mix||0.4 μL|
|Total volume||50 μL|
This step assumes the use of TE 90 GeneSweep™ Sequencing Gel Transfer Unit (Hoefer Scientific Instruments).
|333 μM dNTP mix without dCTP||0.5 μL|
|[α-32P] dCTP (6000 Ci/mmol.)||5 μL (50 μCi)|
|20 μM Primer*||0.625 μL|
|10× Buffer||1.25 μL|
|50 mM MgCl2||0.625 μL|
|Taq polymerase (5 U/μL)||0.25 μL|
|Template DNA fragment**||5 ng|
|Water||to 12.5 μL|
1These instructions assume the use of diploids of the SK1 strain background, with either rad50S or sae2Δ mutation. If other strain backgrounds are used, adjustments to the culture media and/or times for sample collection may be necessary.
2Be careful that the agarose solution is not too hot when you pour it because this may partially melt the DNA agarose plugs and detach them from the comb.
3The volume of SPS should be less than 20% of the flask volume to ensure good aeration. Use a flask of ≥250 mL for 50 mL of SPS culture.
4The volume of SPM should be less than 10% of the flask volume to ensure good aeration, e.g., use a 1 L flask for 100 mL of SPM culture.
5Appropriate time points to collect cells are dependent on the purpose of the experiment. Usually, the DSB frequency reaches a maximum by 5–6 h after transfer to SPM in the SK1 background when using rad50S or sae2Δ mutants.
6The mixture of Solution 1 and the LMP agarose mix should be kept at 40°C as briefly as possible (maximum 3–4 min) in order not to inactivate the zymolyase enzymatic activity. Therefore, it is better not to make an agarose mix for more than 6 samples at a time.
7If you have more than 15 samples to run on the same gel, you may run a 14 cm long by 21 cm wide gel using the same gel casting stand, but with a 21 cm wide comb. The running conditions must be adjusted such that total run time is 30.5 h but all the other conditions are unchanged. An example of such a run is shown in Fig. 2B.
8Try to avoid leaks to protect the phosphor screen and to prevent the blot from drying. If the blot is kept moist, the probe can be stripped by washing twice with 1% SDS for 15 min. However, once the blot is dried with the hybridized probe, it becomes difficult to strip the probe.
9If the region to be analyzed is larger than 10 kb, it is recommended to extract and digest genomic DNA in low melting point agarose as described in Subheading 3.2.1.
10If a residual pellet exists after overnight incubation at 4°C, mix the DNA by gentle tapping. Do not vortex. Often, meiotic samples are cloudy, presumably because of polysaccharide or other components from the ascus or spore walls. However, this does not affect later steps (restriction enzyme digestion, etc.).
11Usually 10 μl of sample corresponds to approximately 1 μg of DNA. For more precision to ensure that the quantity of DNA is similar from sample to sample, the DNA concentration can be quantified by Hoechst dye fluorescence using a fluorometer.
12For accurate DSB mapping, it is important to include appropriate DNA size standards on the same gel, and it is also important that they appear in the autoradiograph of the Southern blot after probing. There are two convenient ways to achieve this goal. The first is to use a commercially available molecular weight marker, which is run in an adjacent lane in the gel (e.g., lambda DNA digested with BstE II; see Fig. 3A, lane M). To visualize the marker on the autoradiograph, marker DNA is also added to the random primed labeling reaction at a probe to marker ratio of 1000:1. Alternatively, molecular weight markers can be prepared from yeast genomic DNA. To do so, approximately 1 μg of 0 h sample (i.e., DNA from a premeiotic culture) is digested with the same restriction enzyme used to digest the meiotic samples. Then, an aliquot of the digested DNA is digested with an appropriate second enzyme which cuts within the region of interest. A set of such double digests is then pooled with the undigested DNA after heat inactivation of the restriction enzymes (see Fig. 3B, lane M). The amount of DNA that is subjected to double digestion should be adjusted dependent on the expected DSB frequency, such that the marker fragments are not stronger than the DSB signal. A typical starting point would be to perform the secondary digestions on aliquots of 1% of the first digest (i.e., ~0.01 μg of DNA). Either method works well for providing size standards, but the second method provides somewhat more accurate size information because it allows one to control for DNA sequence composition and for the amount of DNA loaded in the lane.
131 × TAE may be used instead of TBE. If so, it is essential to circulate the buffer during electrophoresis.
14If the restriction digest worked well, each sample will show the same pattern on ethidium stained gel.
15We obtain good results using a vacuum blotter to transfer DNA from the gel to a hybridization membrane. Alternative methods using capillary transfer under neutral or denaturing conditions may also be satisfactory.
16Depurination is not necessary for DNA fragments less than 5 kb, but we observe that transfer is partial for DNA fragments larger than this. Thus, depurination (and subsequent nicking of apurinic sites) is critical to obtain accurate estimation of DSB frequencies. The depurination step can be replaced by UV treatment at 120 mJ/cm2, which produces alkali-labile photoproducts in the DNA.
17This step is to check the quality of the gel. If there is a problem (e.g., leaks or air bubbles that cannot be removed, etc.), discard the gel and prepare it again.
18We have mapped breaks at single nucleotide resolution at several DSB hotspots (Murakami et al., manuscript in preparation). When the 5′ ends were mapped independently from both sides of DSB hotspots using separate restriction digests and probes, we confirmed that DSB ends always have 2-nucleotide 5′ overhangs, as reported previously (4, 5). However, it is important to note that some 3′ DSB ends were not identical to what was expected based on the mapping of 5′ ends from the other side of the DSB site. Specifically, 3′ ends were often 1–2 nucleotides longer than expected. Therefore, it appears that 3′ ends may sometimes be filled in by DNA polymerase, either in vivo or during the preparation of genomic DNA (B. de Massy, personal communication).
19If the DSB signals are too weak, more genomic DNA can be used. We have confirmed that up to 5 μg of DNA can be loaded in a single lane.
20Mixing the sequence standard with genomic DNA from the 0 hr sample allows for equal amounts of total DNA to be loaded in each lane. This controls for the effects of genomic DNA on the migration pattern on the sequencing gel, and is essential for accurate DSB mapping.
21The electrophoresis time depends on the length of the DSB fragment to be resolved. The following are optimal total times using the conditions described in this protocol:
|150 nucleotides||190 min|
|180 nucleotides||250 min|
|220 nucleotides||290 min|
|240 nucleotides||320 min|
22If the DSB hybridization signals are too weak, increase the total volume with the same concentrations of all reagents. We have used up to 50 μL of total reaction volume successfully.