Strains, plasmids and growth conditions

All strains that were used in this study are listed in Table 1. To estimate the stability of mitochondrial microsatellite repeat sequences, we used a strain harbouring in its mtDNA the arg8^m::(GT)₁₆(+2) reporter in place of the COX3 gene [26]. The strain (YAK136) was obtained by cytoduction of mtDNA from the original strain with the mitochondrial reporter (strain CAB193-1; reference as above) into a rho⁰ MATa arg8Δ::URA3 strain derived from our wild-type strain FF18733 via a MATα kar1-1 strain (DFS160 [27]). The URA3 cassette for inactivation of the nuclear ARG8 gene was amplified by a PCR on the template of a genomic DNA extracted from the DFS160 strain (primers ARG8_A and OA54 in S1 Table). For assays of mitochondrial allelic homologous recombination, we used the method described in [28] with a parental pair of wild-type strains: YAK349 and YAK243. YAK243 was obtained in a way analogous to that described above for the construction of YAK136, i.e. through cytoduction of mtDNA from CAB183-1 (with the arg8^m::(AT)₁₆(+1) reporter replacing the COX3 gene) into a rho⁰ MATa arg8Δ::URA3 strain derived from FF18733 via a DFS160 carrier strain. YAK349, in turn, was isolated through cytoduction of mtDNA from YAK29/1 (via a MATa kar1-1 strain, JC8/55) into a rho⁰ MATα arg8Δ::URA3 strain derived from PJD1 [29]. The procedure used for those and other cytoduction experiments is described in Supplemental Information (S1 Text).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 1. S. cerevisiae strains used in this study.

https://doi.org/10.1371/journal.pone.0180153.t001

Erythromycin-resistance (E^R) mutagenesis was tested in the genetic context of the FF18733 strain [30]. The strain YAK225 was isolated by crossing FF18733 with the isogenic FF18734 strain [30]. Strains that are indicated in Table 1 as segregants were isolated after dissecting tetrads issued from sporulation of heterozygous strains that had been obtained by crossing appropriate haploid strains or transformation of diploid strains (congenic with FF18733 and FF18734) with deletion cassettes. All genetic manipulations were done essentially as described by Amberg et al. [36]. To introduce kanMX4-, hphMX4- or natMX4-marked deletions into our strains, we used mainly the PCR-mediated strategy described earlier [35] with antibiotic-resistance cassettes according to [37] and ORF-specific and marker-specific primers as listed in S1 Table. Deletion cassettes to inactivate studied genes with the kanMX4 maker gene were derived from strains in the Institute collection of yeast deletions from the Saccharomyces Genome Deletion Project [38], cited earlier in [39]. For inactivations of the RAD27 gene, we followed either the above-mentioned procedure, using cassettes with long flanks (primers RAD27_A and RAD27_D; S1 Table) or, as a cautionary measure to avoid introducing inadvertent modifications in the promoter of the adjacent APN1 gene, PCR-amplified kanMX6 or hphMX4 cassettes flanked with shorter flanks (40 bp long). For de novo amplification of the rad27Δ::kanMX6 cassette with short flanks, using the Phusion polymerase (Thermo Scientific), we utilized primers OA117 and OA118 (S1 Table) and plasmid pFA6a-kanMX [40] as a template. For amplification of the rad27Δ::hphMX4 cassette with short flanks, we used primers OA136 and OA137 (S1 Table) and, as a template, genomic DNA extracted from hygromycin B-resistant segregants obtained in tetrad dissection of the YAK385 strain (heterozygous rad27Δ::kanMX4; Table 1) in which the kanMX4 marker had been replaced for hphMX4 according to [37]. DUN1 deletion was performed by transformation of diploid strains with the dun1Δ::kanMX4 fragment (short flanks) amplified with primers OA152 and OA153 on the template of a genomic DNA extract from the BY4741 dun1Δ strain [38]. SML1 deletion strains (sml1Δ::natMX) were constructed according to [41], using primers listed in S1 Table. RRM3 was deleted using an hphMX4-marked cassette with long flanks amplified with primers RRM3_A and RRM3_D (S1 Table). All isolates were verified by PCR amplifications of modified loci, using primers listed in S1 Table, following the methodology described in Johnston et al. [42].

The wild-type copy of the MSH1⁺ gene was replaced by the msh1-R813W allele using the two-step gene replacement method in which EcoRI-linearized plasmid pCK14 [28] was integrated into the chromosomal MSH1 locus in the FF18733 strain (Table 1). The excision of the plasmid was selected in the resulting transformants by passaging twice on a medium with 5-fluoroorotic acid (5-FOA) [36]. Since the R813W substitution in the MSH1-encoded protein had been designed to create a PvuII site in the mutated allele [43], the presence of the mutant copy of the gene in isolated 5-FOA-resistant strains was verified by PvuII digestion of PCR products amplified with primers MSH1_C1 and MSH1_D (S1 Table) on genomic DNA isolated from the strains. The verified msh1-R813W isolates were crossed with FF18734 to obtain heterozygous diploid strains. The PvuII digestion of MSH1_C1-MSH1_D products was also used for tracking msh1-R813W allele segregations in meiotic progeny obtained by sporulation of msh1-R813W heterozygous strains.

To obtain centromeric plasmid pCK62 encoding RAD27, the BglII fragment containing the gene and the LEU2 marker was isolated from pEAI141 [13], kindly provided by Eric Alani, and inserted into pFL38 [44] to substitute for the URA3 containing–BglII fragment. The resulting plasmid was named pCK54. Subsequently, the BamHI site derived from the original multiple cloning site of pFL38 was removed by double cleavage with SmaI and XbaI, followed by Klenow treatment and re-ligation. A plasmid isolated after the BamHI site removal was named pCK62. To convert the wild-type allele of RAD27 into to the rad27-R325* allele, pCK62 was mutagenized, following BamHI cleavage and dephosphorylation with shrimp alkaline phosphatase (Thermo Scientific), by a gap repair recombination using a pair of overlapping mutagenic PCR-generated fragments. One mutagenic fragment was amplified with a pair of primers RAD27_C (S1 Table) and OA112 (ditto) on the template of pCK54 plasmid using the Phusion polymerase (Thermo Scientific). The other fragment was also generated in a Phusion-catalyzed PCR with the pair of primers OA111 (S1 Table) and RAD27_D (ditto) on the template of pCK54. The final construct with the rad27-R325* allele was verified by sequencing and named pCK64.

To obtain a plasmid encoding the fusion of α lacZ fragment with the C-terminus of Exo1, we based our construction on the plasmid p680 (a generous gift from Ophry Pines). The plasmid contains a gene encoding a fusion protein Aco1-α [45] cloned in pRS425-GAL1 [46]. For the construction, p680 was linearized with PstI and dephosphorylated. Subsequently, the cut plasmid was gap-repaired by recombination with a PCR Expand High-Fidelity Taq polymerase (Roche)-amplified fragment, containing EXO1 open-reading frame (ORF), obtained with primers OA85 and OA86 (S1 Table) on the template of a genomic DNA extract from the wild-type strain FF18733. The fragment for the gap repair of the linearized p680 contained the sequence of EXO1 ORF, without its STOP codon, embedded in appropriate sequences homologous to p680 that were directing the recombination. 5’-terminus of EXO1 ORF sequence was linked to the GAL1 promoter. The 3’-terminal end of the gene (omitting the EXO1 STOP codon) was joined to the sequence coding for the α lacZ fragment (with a linker peptide of 4 amino acids, Pro-Gly-Gln-Thr, as described in [47]). Thus, in this gap repair construction, EXO1 ORF (without its STOP codon) replaces the ACO1 ORF in p680. The final construct pRS425-GAL1-EXO1-α was named pCK48. In an analogous way, we constructed pRS425-GAL1-RAD51-α plasmid (pCK40), using primers OA68 and OA69 (S1 Table).

To construct pYES2-CTA1, which enables the expression of the CTA1 gene under its native promoter from a URA3-containing 2μ plasmid, the 2.8 kb-EcoRI fragment, containing the gene, was recloned from YEp352-ura3Δ::kanMX-CTA1 [35] into pYES2 linearized with the same restriction enzyme. The final plasmid pYES2-CTA1 was named pCK78.

Plasmids: pYES-DEST52-EXO1-V5-6xHis and pYES-DEST52-RAD27-V5-6xHis were kindly provided by Miguel Blanco and Stephen West [48]. Plasmid pRDK480 (EXO1 on 2μ) was a generous gift from Richard Kolodner [49].

Media for growth (YPD, YPG as well as synthetic complete media for plasmid selection) and sporulation are essentially as described [36]. For synthetic complete media with appropriate requirement omissions, supplement drop-out mixes were used from BioShop (distributed by Lab Empire). For experiments with strains harbouring the ARG8^m gene, we used a drop-out supplement mix according to [50]. For preparing synthetic media without uracil, a formula based on yeast nitrogen base with ammonium sulfate (Difco) and supplemented with casamino acids at 0.5% and tryptophan (as below) was also used. For nuclear mutagenesis assays to measure frequencies of forward mutations in the CAN1 gene eliciting resistance to canavanine, cultures of tested strains were plated on a minimal glucose medium with all the auxotrophic requirements, unless a particular requirement (uracil or leucine) was omitted in case of plasmid selection, and supplemented with canavanine at 60 mg/l. Auxotrophic requirements were added to synthetic media at concentrations recommended in [51]: tryptophan at 80 mg/l, leucine at 217 mg/l, uracil at 22.4 mg/l, histidine at 47 mg/l, lysine at 162 mg/l. Non-respiring petite colonies were scored on YPG medium supplemented with 0.1% glucose (YPGd). Erythromycin-resistant (E^R) mutants were scored on YPG medium (buffered at pH 6.2) containing 4 g/l erythromycin. For plasmid amplification in bacteria, Escherichia coli strain XL1Blue (Stratagene) was used.

Isolation and genetic analysis of erythromycin-resistant mutants for sequencing analysis

For this experiment, cells were cultured by a method proposed in [52] with some modifications. Cells were spread on YPG plates without sub-cloning directly from frozen stocks at -80°C and incubated for 4 days at 28°C. Subsequently, cells were re-plated on YPD for single colonies. After 3-day incubation of the plates at 28°C, individual colonies were suspended in 20 μl sterile water. A 5-μl-aliquot of the suspension was diluted appropriately in water and plated on YPGd to assess, upon incubation for several days at 28°C, the number of rho⁺ and petite cells in tested suspensions (using the tetrazolium overlay method [53]). The rest of a cell suspension was patched on YPG+erythromycin plates. Cell patches were arrayed in a 12-node grid per plate, i.e. one-colony suspension was patched on a 2-cm² square in the grid and 12 independent gridded YPD-grown colonies per strain were tested in parallel for resistance to erythromycin on one 8.5 cm-diameter plate. Altogether, 12 such plates for the wild-type strain (FF18733) with 12 independent 1-colony-patches were processed and 10 plates for rad27Δ (YAK254; Table 1). Plates were incubated at 28°C and scored for erythromycin-resistant colonies after 7 days and 14 days of incubation. Mean values of E^R mutant frequencies are reported for this experiment in Table 2. 95% confidence intervals of the mean (CI 95%) were calculated, using the GraphPad Prism 6 software. To determine the localization of E^R mutations and ensure the independent origin of analyzed clones, for all the patches, only one E^R colony per patch was picked up and purified by an additional passage on fresh YPG+erythromycin plates for further analyses. Each isolated mutant was subjected to a cytoduction test (S1 Text). In addition, to isolate heterozygous strains for tracking the E^R trait in meiotic progeny, E^R isolates lacking the RAD27 gene were also crossed in parallel with a rho⁰ derivative of the wild-type strain FF18733 carrying a prototrophic allele of the HIS7 gene (YAK1070; Table 1). Diploid cells (His⁺ G418^R) generated in the crosses were selected as described in S1 Text. Tetrads issued from sporulation of isolated diploid clones were dissected on YPD plates and spore clones originating from complete tetrads were analyzed for segregating markers.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 2. The effect of Rad27 deficiency on the frequencies of total and mitochondrial erythromycin-resistant (E^R) mutants.

https://doi.org/10.1371/journal.pone.0180153.t002

Sequencing of mutations in the 21S rRNA locus in mtDNA of mitochondrial erythromycin resistant mutants

Total genomic DNA extracts were prepared from strains with E^R mutations localized in mitochondria by the cytoduction test. For extractions (from 3 ml of YPD culture of strain per extraction), a scaled-down modification of the method with a zymolyase digestion (a yeast DNA miniprep) described in [36] proved to give DNA extracts of a sufficient yield and quality for subsequent PCR amplifications preceding the final sequencing step. The primer pair ERY-1 and ERY-2, according to [54], was used for PCR amplification of the mitochondrial 21S rRNA and sequencing. The primers amplify a mtDNA fragment from the nucleotide at position 1731 to the nucleotide at position 2293 (the coordinates correspond to the 21S rRNA sequence from Saccharomyces Genome Database [55] and not to the GeneBank sequence cited in [54]). Before sequencing, PCR-amplified fragments were purified on Clean Up (A&A Biotechnology) columns. Reported sequence changes in the 21S rRNA were confirmed by two sequencing reactions with both primers.

Determination of erythromycin-resistant (E^R) mutant frequencies

The frequency of E^R mutants was estimated as described in the subsection “Isolation and genetic analysis of erythromycin-resistant mutants for sequencing analysis” (results in Table 2) or in liquid cultures. To start cultures for mutagenesis assays, 3–4 day old colonies growing on YPG plates were inoculated independently into 3 ml of an indicated medium. In experiments with strains harbouring plasmids, colonies with plasmid-transformed cells were picked up directly from transformation plates to inoculate cultures in appropriate selective media. For experiments with plasmids coding for genes under the GAL1 promoter, transformants were isolated on plates with a complete synthetic selective medium with 2% sucrose as the carbon source. Cells from late logarithmic cultures (grown on glucose-containing media for 16 hours, or on media with galactose as the carbon source for 24 hours) were harvested by centrifugation, washed with sterile water and plated on YPG+erythromycin medium. Appropriate dilutions of the cultures were also spread on YPGd plates to calculate the number of petite and grande (rho⁺) cells. All plates were incubated at 28°C. Colonies on YPGd plates were counted after 3–5 days of incubation and scored as rho⁺ or rho¯/rho⁰ (i.e. petite) by the tetrazolium overlay method [53]. E^R colonies were scored after indicated incubation times. Median values derived from several combined experiments with at least 10 independent cultures of each strain were used to determine the frequency of mutants resistant to erythromycin calculated per number of rho⁺ cells. P values for statistical significance of any differences in mutant frequencies between pairs of strains were determined by calculating the nonparametric Mann–Whitney statistics, using the GraphPad Prism 6 program. The same software was used for drawing the graphs and calculating 95% confidence intervals of median values.

We also used the same statistical analysis for processing results of other mutagenesis assays in the current study, with the exception of mitochondrial allelic recombination (details in the section Mitochondrial allelic recombination tests) and the erythromycin-resistance mutagenesis control test (results in Table 2) accompanying the mtDNA sequencing analysis, as described in the preceding section Isolation and genetic analysis of erythromycin-resistant mutants for sequencing analysis.

Determination of canavanine-resistant (Can^R) mutant frequencies

To measure levels of nuclear mutagenesis in tested strains, mutations in the CAN1 gene conferring resistance to canavanine were selected on a medium with canavanine, as described in the subsection Strains, plasmids and growth conditions. Cells were grown in rich or selective media (as indicated), harvested, rinsed with water and plated (around 2.5 x 10⁷ cells/5.5 cm diameter-plate). Can^R mutants were scored after 5 days of incubation at 28°C. Median frequency values were analyzed as described above.

Determination of phenotypes in strains harbouring variants of the mitochondrial reporter gene ARG8^m

Colony size estimation

Frozen cells (at –80°C in 15% glicerol) of E^R strains were thawed by plating at low density on plates with YPG (pH 6.2) medium with erythromycin and grown for 4 days at 28°C. The plates were imaged using a 24-megapixel (6000 X 4000 pixels) dSLR camera (Sony) with an APS-C sensor (23.5 X 15.6 mm) and a macro lens with 1:1 reproduction ratio (Tamron), resulting in a scale of ~256 pixels/mm. Diameters of colonies (n = 25 for each strain) were measured in pixels using the line tool in the NIH ImageJ2 software, Fiji distribution [56–58]. Colony diameter mean values were compared using the unpaired t-test.

α-complementation assay

The wild-type haploid strain FF18733 was co-transformed, on the one hand, with plasmids encoding proteins to be tested in a C-terminal fusion with the α fragment of lacZ protein (either pRS425-GAL1-EXO1-α, alias pCK48, or pRS425-GAL1-RAD51-α, alias pCK40) and, on the other hand, with a plasmid encoding either the lacZ ω fragment localized in the cytoplasm (p877, alias pYES/M15 or pWc [59]) or a plasmid coding for the ω fragment directed to mitochondria (p999, alias pWm [59]). For the α-complementation test, transformed strains were cultured in a non-repressing selective synthetic complete medium with 3% glycerol and 0.1% glucose for 2 days. 5 μl-aliquots of undiluted cultures were spotted on a solid selective medium with 2% galactose as a sole carbon source (buffered at pH 7.0) and with X-gal (5-bromo-4-chloro-3-indolyl β-D-galactopyranoside) indicator at 40 mg/l (after the protocol in [36]). The color of yeast colonies, arising due to β-galactosidase activity on the detector plates, was observed after 5 days of incubation at 28°C.

Fractionation of cells expressing Exo1-6xHIS and analysis of mitochondrial fractions

Cells of a rad27Δ::hphMX strain (YAK1441) transformed with pYES-DEST52-EXO1-V5-6xHis were grown in selective medium with 2% galactose and 0.1% glucose up to OD₆₀₀ 1.0 and harvested. The mitochondrial fraction was isolated by differential centrifugation according to a described procedure [60]. Samples of the mitochondrial fraction (15 μg/sample) were analyzed by treatment with proteinase K (at concentration 50 μg/ml) in a reaction set-up according to [61]. Digestions were terminated by adding phenylmethane sulfonyl fluoride (PMSF) at 2mM and centrifuged at 12 500 x g for 5 min at 4°C. The pellets were analyzed by 12% SDS-PAGE and a Western blot. His-tagged proteins were detected with the GenScript anti-His monoclonal antibody (at the dilution of 1:5000) and the mitochondrial control Cox2 protein with the monoclonal anti-MTCO2 antibody [4B12A5] from Abcam at the concentration recommended by the manufacturer. As a secondary antibody, a polyclonal goat anti-mouse immunoglobulin with horseradish peroxidase from DakoCytomation (at the dilution of 1:2500) was used, followed by a chemiluminescence detection procedure, as in [28].

Verification of mitochondrial phenotypes caused by Rad27 inactivation

RAD27 inactivation has previously been shown to cause mitochondrial phenotypes, such as: an increase in the frequency of point mutations in mtDNA and a decrease both in mitochondrial recombination and microsatellite instability [20]. To elucidate these phenotypes, we repeated and extended the analysis of the effect of Rad27 deficiency on point mutations in mtDNA leading to the resistance to erythromycin by verification of mitochondrial location of erythromycin-resistant (E^R) mutations. We also employed systems alternative to those described earlier to measure mitochondrial recombination and microsatellite stability in rad27Δ strains.

Mutations conferring resistance to erythromycin are acquired through specific changes in the mitochondrial 21S rRNA gene [62–64] and the acquisition of resistance to this drug is a standard method of measuring point mutagenesis in mtDNA [52, 65]. Table 2 shows that the rad27Δ mutant displayed an about 2-fold increase in the frequency of E^R mutants as compared to that of the wild-type strain. However, when scoring E^R colonies, we observed that incubation of cells on the antibiotic-containing medium for periods longer than 7 days, the minimal time routinely used in our laboratory for scoring E^R, resulted in the appearance of additional erythromycin-resistant colonies. In the wild-type strain, we observed after 14 days less than a 2-fold increase in the frequency of E^R mutations as compared to that estimated after 7 days (Table 2). In the rad27Δ mutant, the frequency of E^R mutations increased by about 12 fold after the longer incubation time, as compared to that estimated after the 7-day incubation (P < 0.0001, Mann-Whitney test). The observation that E^R mutants accumulate progressively on selective medium during prolonged incubation times has previously been reported for rad27Δ mutants [20]. We designated those E^R mutants which arise up to the 7th day of incubation as early mutants, and those which arise after additional incubation as late mutants.

To confirm that the E^R phenotype is really due to a mitochondrial mutation, we determined whether the E^R trait could be transferred to a rho⁰ kar1-1 strain by cytoduction. In parallel, we also crossed each mutant with a wild-type strain to obtain a heterozygous strain for meiotic segregation analysis, so that we could corroborate the nuclear location of an E^R mutation if it had given a negative result of the mitochondrial location by the cytoduction test. We found that all of the tested early E^R mutants in the wild-type and rad27Δ strains (41 and 35 isolates, respectively) were able to transfer their resistance during the abortive mating with the rho⁰ recipient strain, indicating the mitochondrial location of these mutations (Table 2). In contrast, only about 50% of the tested 43 late E^R mutants in the rad27Δ strain were able to transmit their resistance phenotype by cytoduction, suggesting that a large fraction of late E^R mutants in rad27Δ was due not to mitochondrial, but to nuclear mutations (Table 2). The nuclear nature of the E^R mutations which had failed to be transmitted with mitochondria in the cytoduction test was confirmed by the segregation analysis of spores from dissected tetrads issued from sporulation of heterozygous strains obtained in crosses between late rad27Δ E^R mutants and a wild-type strain. For those late E^R mutations, heterozygous strains produced tetrads in which the resistance trait segregated 2:2 (tetrads were analyzed from 11 independent heterozygous diploid strains), indicating that non-mitochondrial E^R mutations were nuclear and a single mutation of this type was sufficient for a cell to acquire resistance to erythromycin. Although, contrary to long-standing views in literature [20, 52], our analysis shows that at least for rad27Δ strains a mutation causing resistance to erythromycin cannot be automatically equated with a mutation in mtDNA, our results confirm the earlier results by Kalifa et al. [20] that the level of mitochondrial point mutations increases due to Rad27 deficiency.

Previous studies on the role of Rad27 in the control of stability of microsatellite sequences indicated that, in contrast to the nucleus in which Rad27 promotes microsatellite stability, inactivation of RAD27 resulted in inhibition of rearrangements in those sequences in mitochondria [20]. However, whereas the notion that the Rad27 activity limits the stability of microsatellite sequences in the nucleus is based on several independent studies [24, 66–69], to date only one study has reported the opposite effect of Rad27 in mitochondria [20]. Therefore, we decided to verify the observation in another genetic background than the one used in the cited report. We introduced the arg8^m::(GT)₁₆(+2) reporter [26] into a rho⁰ derivative of our wild-type strain devoid of the nuclear ARG8. Subsequently, we inactivated the RAD27 gene in the resulting strain and tested the stability of GT dinucleotide repeats in the mitochondrial ARG8 reporter. Our results showed that deletion of the RAD27 gene decreased over 10 fold the frequency of frameshifts within the repeated tract leading to an Arg⁺ phenotype (Fig 1A). Introduction of a low-copy-number plasmid encoding RAD27 to rad27Δ cells not only compensated for the decrease, but even slightly increased the frequency of Arg⁺, confirming that Rad27 stimulates mitochondrial microsatellite instability. The plasmid containing a defective truncated allele rad27-R325*, which confers nuclear phenotypes closely resembling those exhibited by rad27Δ strains [13, 70], failed to suppress the increased stability of the di-nucleotide tract in the mitochondrial reporter, further corroborating the requirement of the fully functional Rad27 nuclease in regulation of mtDNA sequence stability.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Fig 1. Effects of Rad27 deficiency on other mtDNA-related phenotypes.

Panel A. Rad27 stimulates GT dinucleotide repeat instability. Frequencies of Arg⁺ prototrophs were determined in cultures of the wild-type strain (YAK136), harbouring mtDNA with the arg8^m::(GT)₁₆(+1) gene, and its rad27Δ derivative (YAK1629) transformed with either control vector (pRS415), pRAD27⁺ (pCK62) or prad27-K325* (pCK64). Values represent median frequencies in three combined independent experiments with 10–15 cultures of each transformant strain. P-values computed with the Mann-Whitney statistics–***: 0.0008; ****: < 0.0001. Panel B. Allelic recombination in the mitochondrial arg8^m reporter gene is decreased due to Rad27 deficiency. Frequencies of mitotic Arg⁺ segregants were determined in diploid cultures after crosses of wild-type strains (YAK349 xYAK243) and their rad27Δ derivatives (YAK1094 x YAK1097) transformed with the indicated plasmids (as in Fig 1). Values represent average frequencies in cultures obtained in 16 crosses for each pair of transformed parental strains. Error bars are SEM (standard error of the mean). P-values computed with the two-tailed unpaired t-test–**: 0.0024; ****: < 0.0001; ns: not significant.

https://doi.org/10.1371/journal.pone.0180153.g001

Frequency of homologous recombination has also been reported to be affected by Rad27 deficiency in opposite ways in the nucleus and mitochondria. Cells lacking Rad27 exhibit a mitotic hyper-recombination phenotype [15, 71]. In contrast, the mitochondrial intra-chromosomal recombination, measured as the frequency of deletions of the intervening sequence between a 96 bp-long direct repeats, is limited by Rad27 deficiency [20]. To get broader insight into the role of Rad27 in the mitochondrial recombination, we used an ARG8^m recombination assay previously developed in our laboratory to measure the frequency of mitochondrial allelic recombination [28]. Wild-type or rad27Δ parental strains carrying mitochondrial heteroalleles of the ARG8^m reporter gene were crossed and frequencies of arginine prototrophs were scored in final diploid cultures. The results showed that Rad27 deficiency caused a significant decrease (p = 0.0024) in the frequency of Arg⁺ recombinants (Fig 1B), indicating that mitochondrial inter-chromosomal recombination is also dependent on the active Rad27 nuclease. The conclusion was confirmed by complementation of rad27Δ-dependent recombination frequency decrease within ARG8^m with the plasmid containing the wild-type RAD27 gene and lack of complementation with the plasmid carrying the truncated rad27-R325* allele. The similar effects of RAD27 inactivation on both intra- (between direct repeats) and inter-chromosomal recombination of mtDNA (heteroallelic) further support the general conclusion that, in contrast to its role in recombination in the nucleus, Rad27 activity stimulates mitochondrial homologous recombination.

Inactivation of Rad27 alters the spectrum of mitochondrial mutagenesis

The mechanism underlying the enhanced mitochondrial mutagenesis conferred by Rad27 deficiency is not understood [20]. To get more insight into the mitochondrial mutator phenotype caused by the lack of the Rad27 nuclease, mutations arising in mtDNA of rad27Δ deletion mutants were subjected to a detailed analysis. Because individual mutagenic mechanisms often demonstrate their signatures in DNA sequence [72–74][75], the region from the position 1731 to 2293 (the coordinates of the 21S rRNA locus sequence in Saccharomyces Genome Database [55]) of the mitochondrial 21S rRNA gene was sequenced from independently isolated early and late E^R colonies. This region contains a stretch of three nucleotides (GAA, positions: 1957–1959) whose mutations commonly confer resistance to erythromycin [54, 62–64]. The sequence analysis of E^R mutations indicated that the majority of early mutations found in the wild-type strain and in the rad27Δ single mutant were AT → GC transitions and AT → TA transversions at position 1958 (Table 3). Those two mutations were frequent also in late mitochondrial E^R in the mutant strain, but, mainly, a remarkable GC → AT increase at position 1957 was found (the frequency of 19.1/ 10⁷ rho⁺ in rad27Δ vs. 0.41/ 10⁷ rho⁺ in the wild-type; Table 3).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 3. Nucleotide changes in the sequenced region of the mitochondrial 21S rRNA gene (rib3 locus).

https://doi.org/10.1371/journal.pone.0180153.t003

Since the GC → AT transition at position 1957 appeared to be a hallmark of late E^R mutations arising in the rad27Δ strain, we asked about the origin of these mutations. Theoretically, GC → AT mutations could have occurred in pre-cultures (in this case, in pre-cultured colonies) before plating cells on the selective medium and then appeared with a delay on erythromycin plates because of a slower growth rate. In point of fact, we did notice some heterogeneity of colony sizes among strains growing on the erythromycin-containing medium. Alternatively, the characteristic GC → AT mutations could arise as post-plating mutations in cells exposed to the erythromycin-mediated toxic stress. In an attempt to distinguish between these two possibilities, we measured the growth rates of a GC → AT mutant and compared them to those of AT → GC and AT → TA mutants, both in the wild-type and rad27Δ background. The growth test was performed by measuring average colony sizes of the mutant strains after a 4-day incubation at 28°C on the erythromycin medium. Table 4 shows that GC → AT mutants, both in the wild-type and rad27Δ background, exhibited a significant decrease in the average colony size as compared to those of AT → GC (P-values for comparisons of colony diameters for both the wild-type and rad27Δ were < 0.0001 by the two-tailed unpaired t-test) or AT → TA mutants (P-values, as above). On the other hand, a similar analysis of mutant colonies growing on YPG plates without the addition of erythromycin showed that in general there were no significant differences between colony sizes of all three mutants in both the wild-type and rad27Δ. Thus, the GC → AT mutation at position 1957 in the 21S rRNA gene confers a growth defect only on YPG medium in the presence of erythromycin, suggesting that the substitution at this gene position is suboptimal for the expression of the erythromycin resistance phenotype. The slower growth rate of the GC→AT mutants goes some way towards explaining the heterogeneity of observed colony sizes on erythromycin-containing media and may also be a contributing factor in delaying the growth of some pre-plating mutants.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 4. Average colony sizes of different mitochondrial E^R mutants.

https://doi.org/10.1371/journal.pone.0180153.t004

GC→AT transitions are frequently caused by either spontaneous deamination of cytosine and generation of pro-mutagenic G:U mispairs [76], or by oxidative deamination of cytosine to 5-hydroxyuracil and uracil glycol [77]. Consequently, GC→AT transitions are considered to be common mutations resulting from oxidative damage to DNA [78]. We considered the possibility that the frequent generation of GC→ AT transitions in rad27Δ may be related to oxidative stress-induced lesions in mtDNA. To verify this hypothesis, we examined whether the generation of E^R mutants in rad27Δ is altered in cells harbouring a multicopy plasmid bearing the CTA1 gene. The CTA1 gene encodes catalase A, a well-known scavenger of H₂O₂. Catalase A localizes both to peroxisomes and mitochondria and a significant import of catalase A into mitochondria occurs when cells are grown under respiratory conditions [79]. We have previously shown that introduction of a multicopy plasmid encoding CTA1 suppresses antimycin A-induced rearrangements of mtDNA [35]. As shown in Fig 2A, overproduction of catalase A did not decrease the level of E^R mutations occurring in the rad27Δ strain, suggesting that the mitochondrial mutations in Rad27 deficient yeast did not originate from stress-induced oxidative lesions in mtDNA that could be counteracted by overproduction of catalase A. Consistently, we did not detect significantly increased levels of reactive oxygen species (ROS) in rad27Δ strains, using 2′,7′-dichlorofluorescein as a probe for ROS detection (S1 Fig).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Fig 2. Neither oxidative stress, nor Mec1/Rad53-dependent induction of DIN7 expression determines the increase in E^R mutations in Rad27 deficient yeast.

Panel A. Overexpression of CTA1 does not decrease the E^R mutation frequency in a rad27Δ strain. Frequency of E^R mutations in control strain (FF18733) and its rad27Δ derivative (YAK1405) transformed with indicated plasmids. The strains (10–30 independent cultures/strain) were grown overnight in a synthetic medium with casamino acids (supplemented with tryptophan as described in Materials and Methods) with 3% glycerol and 0.1% glucose (non-repressing conditions) and plated on the medium containing erythromycin. Median frequencies of E^R mutants scored on the 10^th day of incubation are shown. ns—not significant according to the Mann-Whitney statistics (P = 0.08); ****: P-value < 0.0001. Vector: pYES2; pYES2-CTA1: pCK78. Panels B and C. Enhanced mutagenesis to erythromycin resistance in a rad27Δ strain does not depend on the presence of Din7. The following strains: WT (FF18733), din7Δ (YAK1446), rad27Δ (YAK1441), and rad27Δ din7Δ (YAK1444) were grown overnight in YPD medium and tested on medium containing erythromycin as described in Materials and Methods. E^R mutants were scored on the 7^th (Panel B) and on the 12^th (Panel C) day of incubation at 28°C. Median frequencies are shown from combined three independent experiments with 10–12 cultures of each strain. ****: P-value < 0.0001; *: P-value: 0.0343; ns: not significant.

https://doi.org/10.1371/journal.pone.0180153.g002

Mutagenesis leading to erythromycin resistance in strains lacking Rad27 does not depend on Din7

It has been shown that activation of the conserved Mec1/Rad53 cell-cycle checkpoint (called the DNA damage response; DDR in short) pathway due to instability of the nuclear DNA leads to the increased copy number of mitochondrial DNA [80]. Since Rad27 deficiency results in replication stress [15, 17, 81] and activation of checkpoint kinases [16, 17], we considered the possibility that the mitochondrial mutator phenotype of rad27Δ may result from activation of the cell-cycle checkpoint pathway. It is known from our previous studies that activation of the Mec1/Rad53 pathway results in increased cellular abundance of the uniquely mitochondrial nuclease Din7 [82, 83]. Furthermore, we have also shown that enhanced production of Din7 results in increased mitochondrial mutagenesis [43, 83, 84]. Thus, the mutator phenotype of rad27Δ strains could reflect the elevation of the Din7 nuclease level due to Mec1/Rad53 activation in Rad27 deficient yeast. To test this possibility, we compared the frequency of E^R mutants generated in the rad27Δ din7∆ double mutant strain to that of the rad27∆ single mutant strain. As shown in Fig 2B, early E^R frequencies were similar in both mutants, indicating that enhanced mitochondrial mutagenesis in cells lacking Rad27 is not dependent on the elevated level of Din7. Furthermore, after a prolonged incubation time the median frequency of E^R mutants in the rad27∆ din7∆ double mutant strain increased slightly over that in the rad27∆ single mutant strain (Fig 2C), though with a marginal statistical significance, pointing to a weak synergistic effect between deletions of RAD27 and DIN7. Altogether, the results clearly show that Din7 activity is not required for the E^R mutagenesis in strains lacking the Rad27 nuclease.

Dun1-dependence of mitochondrial mutagenesis in strains lacking Rad27

The major target of activated Mec1/Rad53 pathway is the pool of DNA synthesis precursors. Since yeast cells carrying the rad27∆ mutation require both Mec1 and Rad53 to survive, we analyzed the role of Dun1, the effector kinase of the checkpoint phosphorylation cascade, in the mitochondrial mutagenesis caused by Rad27 deficiency. We constructed a rad27∆ dun1∆ double mutant strain and established the frequency of Can^R and E^R mutations. Intriguingly, the results indicated that deletion of DUN1 in rad27∆ cells suppressed both nuclear and mitochondrial mutator phenotypes, resulting in an over 80% decrease of Can^R and an over 95% decrease of both early and late E^R mutations (Table 5).

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 5. The effect of Dun1 and/or Sml1 deficiencies on the nuclear and mitochondrial mutagenesis in rad27Δ cells.

https://doi.org/10.1371/journal.pone.0180153.t005

Activation of Dun1 kinase by the Mec1/Rad53 pathway causes transcriptional induction of the ribonucleotide reductase (RNR) genes [85] as well as phosphorylation and concomitant degradation of Sml1, the inhibitor of the large subunit of ribonucleotide reductase, Rnr1 [86]. Consistently, in a recent report by Hendry et al. [16], the authors propose that Rnr3 abundance can be used as a specific read-out of the DDR pathway activation and genome instability (similar results were also reported earlier by Tang et al. [87]). A strain lacking Rad27 displays a highly increased abundance of Rnr3 both in spontaneous conditions (2^nd rank among over 5000 mutants) and when tested in the presence of a DNA-methylating agent, methylmethane sulfonate (1^st rank) [16]. RNR3 transcription is repressed in unstressed cells by Crt1/Rfx1 repressor which is phosphorylated by the Dun1 kinase upon its activation by the DDR Rad53 kinase [88]. Hyperphosphorylation of Crt1/Rfx1 releases the repressor from DNA which allows the induction of target gene transcription. Thus, the increased abundance of Rnr3 in rad27Δ cells strongly suggests that the Mec1/Rad53 pathway is activated due to nuclear genome instability in those cells. Consequently, the activation of the signaling pathway should lead to an increase in dNTP synthesis. Elevation of dNTP pools, in turn, could result in increased mutagenesis of both nuclear and mitochondrial genomes. To check this possibility, we inactivated the SML1 gene in the rad27Δ dun1Δ double mutant background. We expected that deletion of SML1 should at least partially restore dNTP pools and mutator phenotypes in the rad27Δ dun1Δ strain. The prediction turned out to be correct because elimination of the ribonucleotide reductase inhibitor from rad27Δ dun1Δ cells brought back elevated levels of nuclear (Can^R) and mitochondrial (E^R) mutations (Table 5). This result confirms the hypothesis that both nuclear and mitochondrial mutator phenotypes in Rad27 deficient cells result from the increased dNTP pools that are a consequence of Mec1/Rad53/Dun1 pathway activation. Interestingly, however, whereas the moderate early E^R mutagenesis in the triple deletion rad27Δ dun1Δ sml1Δ mutant reached exactly the level of that measured in the rad27Δ mutant, the late E^R mutations are significantly less frequent (by 2.7 fold) in cultures of the triple deletion rad27Δ dun1Δ sml1Δ mutant than those detected in cultures of the rad27Δ single mutant. On the contrary, the level of Can^R mutagenesis in the rad27Δ dun1Δ sml1Δ strain is slightly, though significantly, higher than that in the rad27Δ strain. Thus, although our results point to an increase of dNTP pools as an actual source of mutagenesis in rad27Δ, this increase does not provide an explanation for all of the Dun1-dependent mutations in mtDNA. There is yet another Dun1-dependent mutagenic mechanism affecting mitochondrial DNA, which remains to be established.

To find out if mutations in other repair genes have a similar to rad27Δ indirect effect on mitochondrial DNA, we combined a DUN1 deletion allele with a deletion of the RRM3 gene coding for the Rrm3 DNA helicase involved in rDNA replication [89]. Rrm3 deficiency induces RNR3 expression, albeit at a lower level than that induced by the lack of Rad27 activity [16]. On the other hand, it has previously been reported that inactivation of RRM3 results in increased levels of mitochondrial point mutations [90]. Surprisingly, in our strains lack of Rrm3 resulted in only a slight (2-fold), though statistically significant, increase of E^R mutation levels (Fig 3), suggesting that the previously described effect of rrm3Δ on mitochondrial DNA could be dependent on the specific background of the strain under study and/or on the capacity of mitochondrial DNA repair machinery. Therefore, we analyzed the rrm3Δ-induced mitochondrial mutagenesis in yeast strains harbouring a deficient allele of the MSH1 gene encoding the mitochondrial DNA repair protein [91][28]. The R813W mutation in the ATPase domain of the Msh1 protein increased the frequency of E^R mutations by 20 fold in the RRM3⁺ context, but in the rrm3 null background the msh1 allele caused a synergistic effect increasing the frequency of E^R by 85 fold (Fig 3). This result confirms that the detection of rrm3Δ effect on mtDNA stability depends on the mtDNA repair potential and suggests that Msh1 is involved in the repair of rrm3Δ-provoked DNA lesions in this compartment. Dun1 deficiency entirely abolished the synergistic effect of rrm3Δ msh1-R813W double mutation on E^R mutation frequency (Fig 3). The results indicate that rrm3Δ-induced mitochondrial point mutations are Dun1-dependent, which suggests in turn that the indirect effect of Dun1 activation on mtDNA stability concerns not only cells lacking Rad27, but generally also other cells in which the DNA damage checkpoint pathway is activated. This suggestion, however, requires further investigations.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Fig 3. Inactivation of the DUN1 gene suppresses the increased mitochondrial mutagenesis in an msh1-R813W rrm3Δ strain.

The following strains: WT (FF18733), rrm3Δ (YAK1339), msh1-R813W (YAK1445), msh1-R813W rrm3Δ (YAK1337), and msh1-R813W rrm3Δ dun1Δ (YAK1884) were grown overnight in YPD medium and tested on the medium containing erythromycin as described in Materials and Methods. E^R mutants were scored on the 14^th day of incubation at 28°C. Median frequencies are shown from combined two to five independent experiments with 10–12 cultures of each strain. ****: P-value < 0.0001.

https://doi.org/10.1371/journal.pone.0180153.g003

The effect of Exo1 overproduction on mitochondrial phenotypes resulting from Rad27 deficiency

The results showing that Dun1 activation-dependent activities are responsible for mitochondrial mutagenesis in rad27Δ mutant points to the perturbation of nuclear DNA metabolism in this mutant as the main cause of the mitochondrial mutator effect. To verify this idea, we suppressed the nuclear effects of rad27Δ by overproducing Exo1. Exo1 is a 5’-3’exonuclease and a flap endonuclease engaged in DNA recombination and several other DNA repair pathways [92–94]. It was previously reported that overproduction of Exo1 leads to suppression of nuclear phenotypes connected with Rad27 deficiency [13, 17, 49, 95–97]. We analyzed the effect of EXO1 overexpression under control of the GAL1 promoter on the frequency of E^R in the rad27Δ background. As a control, we used an isogenic construct overexpressing RAD27 and, in parallel, we monitored the effect of overproduction of both nucleases on the level of nuclear mutagenesis (Can^R). As shown in Table 6, the nuclear and mitochondrial mutator phenotypes are suppressed to the same extent due to Exo1 overproduction.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Table 6. Overexpression of EXO1 suppresses both the mitochondrial and nuclear mutator phenotypes in rad27Δ cells.

https://doi.org/10.1371/journal.pone.0180153.t006

Exo1 is considered a cytoplasmic/nuclear protein and as far as we know no report has indicated its entrance into mitochondria. However, since in our experiments Exo1 was overproduced, we wanted to make sure that the Exo1-mediated suppression was not due to the possible moonlighting activity of this nuclease in mitochondria. We employed a previously described lacZ α-complementation assay for dual protein localization in yeast cells [47, 59] and constructed a plasmid encoding a fusion protein in which the α fragment of β-galactosidase was attached to the C-terminus of Exo1. The Exo1-α fusion protein was produced under the control of the GAL1 promoter. This plasmid was co-expressed in yeast cells with an ω fragment of β-galactosidase localized either to the cytoplasm (ω_cyto/ ω_c) or mitochondria (ω_mito/ ω_m). As shown in Fig 4A, the β-galactosidase activity was detected in a plate assay in yeast strains which beside the Exo1-α protein fusion produced ω_c, but not ω_m. In contrast, we obtained positive results of the α-complementation assay for both ω_c and ω_m when we tested the reference Rad51-α fusion protein, which could be expected to reside both in the cytoplasm and mitochondria according to a recent report by Stein et al. [98]. The absence of Exo1 from mitochondria was additionally confirmed by fractionation of rad27Δ cells harbouring pGAL1-EXO1 and analysis of resulting mitochondrial fractions with proteinase K followed by Western blotting (Fig 4B). Although we detected Exo1 in the crude fraction containing mitochondria and other cellular membranes, the Exo1 band disappeared after treatment of the fraction with proteinase K. Altogether, the results confirm that the full length Exo1 does not enter mitochondria and that the overproduction of Exo1 in the nucleus is sufficient to suppress mitochondrial E^R mutagenesis resulting from Rad27 deficiency.

Download:

• PPT
  PowerPoint slide
• PNG
  larger image
• TIFF
  original image

Fig 4. Overproduced Exo1 indirectly suppresses most of the mitochondrial phenotypes of Rad27 deficient strains.

Panels A and B. Exo1 is not localized to mitochondria in yeast cells. Panel A. Cultures of the wild-type haploid strain (FF18733) co-expressing indicated α fusions with a cytosolic ω fragment (ω_cyto) or ω attached to a mitochondrial targeting sequence (ω_mito) were spotted on galactose medium plates containing X-gal and incubated for 5 days at 28°C. Panel B. Mitochondrial localization of Exo1-6xHis was verified by fractionation of cells from a galactose grown culture of a rad27Δ strain harbouring the pGAL1-EXO1 plasmid (the same strain was tested in experiments presented in Table 6). Equal portions of the resulting mitochondrial fraction were or were not treated with proteinase K and analyzed by Western blotting using an anti-His antibody for the detection of the Exo1 protein and an anti-Cox2 antibody for the detection of the control Cox2 protein localized in the inner mitochondrial membrane. Panels C-E The effects of EXO1 overexpression on mitochondrial phenotypes caused by Rad27 deficiency. Panel C. The frequency of arginine prototrophs arising by rearrangements of mitochondrial microsatellite sequences in strains transformed with control vector (YEp13) or pRDK480 overexpressing EXO1 (pEXO1) and bearing in their mtDNA the ARG8^m gene interrupted out-of-frame by a tract of GT dinucleotide repeats. The values represent medians of Arg⁺ frequencies in cultures grown in a glucose selective medium from three combined separate experiments with 10–15 independent cultures/transformant strain. P-values were computed using the Mann-Whitney statistics. **: 0.003; ****: < 0.0001. WT: YAK136; rad27Δ: YAK1629. Panel D. Mitochondrial allelic recombination within the ARG8^m reporter gene in crosses of wild-type strains (YAK349 xYAK243) and their rad27Δ derivatives (YAK1094 x YAK1097) transformed with indicated plasmids (the same as in panel A). Frequencies of Arg⁺ prototrophs were determined in diploid cultures grown under conditions selective for the presence of the plasmids. The columns correspond to average values of Arg⁺ frequencies in cultures obtained in 16 crosses for each pair of parental strains. Error bars are SEM. P-values were computed using the two-tailed unpaired t-test. *: 0,0462; ****: < 0.0001. Panel E. Levels of direct-repeat mediated deletions in mtDNA of a reference strain (EAS748) and its rad27Δ derivative (YAK1659) transformed with a vector plasmid (as above) and pRDK480. The columns represent median frequencies, from two combined separate experiments with 10–20 colonies/transformant strain, of respiring cells arising after the ARG8^m deletion in mtDNA in 4 day-old colonies of indicated transformant strains, grown on plates with a synthetic complete medium selective for the presence of the plasmid (without leucine), but supplemented with arginine. P-values by Mann-Whitney statistics: **: 0.003; ns: not significant.

https://doi.org/10.1371/journal.pone.0180153.g004

This result raised the question of how the Exo1 overproduction affects other mitochondrial phenotypes caused by Rad27 deficiency. We introduced a multicopy plasmid pRDK480 [49] containing an insert with the EXO1 gene into cells carrying the mitochondrial ARG8^m gene interrupted by the GT₁₆ (+2) tract. As a control, to verify the ability of Exo1 overproduction to suppress the nuclear mutagenesis in Rad27 deficient cells in the background of the Arg^¯ non-respiring strain, instead of using the standard canavanine-resistance assays, we tested the cycloheximide-resistance mutagenesis as an indicator of accumulating point mutations in the nuclear genome (as used previously in e.g. [99]). Introduction of this plasmid into rad27Δ cells caused an over 90% decrease in the level of nuclear mutations conferring resistance to cycloheximide (S2 Table), confirming the expected suppression effect. The overexpression of EXO1 also significantly increased the level of mitochondrial microsatellite instability (Fig 4C), indicating that suppression of DNA instability in the nucleus affects microsatellite stability in mitochondria of rad27Δ cells. However, although we noticed a significant increase in Arg⁺ frequency due to the presence of pRDK480, this increase did not reach the Arg⁺ level measured in cells producing the wild-type Rad27 protein. This partial effect suggests that the fraction of Rad27 residing in mitochondria can contribute to the stimulation of mitochondrial microsatellite instability as well. In the allelic recombination assay with Exo1-overproducing rad27Δ cells, over 70% of this recombination was restored by introduction of pRDK480 into rad27Δ cells (Fig 4D). On the basis of the results, we cannot exclude that the presence of Rad27 in mitochondria also directly influences allelic recombination of mtDNA. However, this effect seems to be marginal, since the full suppression effect could be expected at the level of 90%. Altogether, the result indicates that nuclear processes activated in rad27Δ cells are responsible for limiting a great part of mitochondrial allelic recombination. Curiously, the overproduction of Exo1 in yeast cells, carrying a previously described system of direct repeats in mtDNA [20, 34], did not affect the level of direct repeat-mediated deletions (DRMD) in the rad27Δ mutant (Fig 4E). This result suggests that, in contrast to the above-mentioned allelic recombination, mitochondrial Rad27 functions directly in the homologous recombination pathway leading to DRMD in mtDNA.