FACS analysis:

Approximately 1 × 10⁷ cells were collected from log-phase cultures and processed as described (Jorgensen et al. 2004). DNA was stained with Sytox Green (Molecular Probes, Eugene, OR) and profiles were analyzed using a Becton Dickinson (San Jose, CA) FACS Calibur machine and the CellQuest Pro and ModFit LT software (BD Biosciences).

DNA damage and genotoxic stress assays:

Saturated cultures were adjusted to OD₆₀₀ = 0.8 and serially diluted in 10-fold steps and 4-μl volumes spotted onto untreated medium or medium containing or exposed to 0.02% (v/v) methyl methanesulfonate (MMS), 200 mm HU, 5 μg/ml camptothecin, 0.1 μg/ml 4-nitroquinolone-1-oxide (4-NQO), 200 J/m² UV or 100 Gy X rays. Plates were incubated at 30° for 2 days. Conditions used to test for the intra-S checkpoint were as described (Paulovich and Hartwell 1995). Plasmids expressing full-length DIA2 (pMT2484) or DIA2^ΔF-box (pMT2742) from the endogenous DIA2 promoter were used in MMS sensitivity complementation tests.

Protein detection:

Co-immunoprecipitation experiments with FLAG- and MYC-tagged constructs were as described previously (Ho et al. 2002). For immunoblots, proteins were separated on a 10% or 12% SDS–PAGE, transferred to PVDF membranes, and probed with 9E10 monoclonal antibody (1:10,000), anti-FLAG M2 monoclonal antibody (1:2000, Sigma, St. Louis), or anti-Skp1 polyclonal (1:5000), as indicated. Detection was with HRP-conjugated secondary antibodies at 1:10,000 dilution (Amersham, Buckinghamshire, UK) followed by Supersignal West Pico chemiluminescent substrate (Pierce, Rockford, IL) and analysis on a Fluor-S Multiimager (Bio-Rad, Hercules, CA). To detect Rad53 isoforms, log-phase cultures were arrested in G₁ phase with α-factor, washed, and released into fresh media. Total protein extracts were resolved by 7.5% SDS–PAGE, transferred onto nitrocellulose, and probed with a primary rabbit anti-Rad53 antibody (1:750) and secondary donkey anti-rabbit HRP-conjugated antibody (1:10,000, Amersham). For in situ autophosphorylation (ISA) assays, log-phase cultures were either treated or not treated with 200 mm HU for 1 hr, and then processed as described (Pellicioli et al. 1999).

Synthetic genetic array screen:

A dia2ΔURA3 MATα strain (yMT1940) was mated to a collection of 4422 individual xxxkanR MATa haploid deletion mutants (Giaever et al. 2002) by manual replica pinning as described (Tong et al. 2001). Three independent screens were carried out in triplicate. All genetic interactions identified in at least two of the three screens were subsequently confirmed by tetrad analysis. Synthetic lethal interactions were defined as double mutants that failed to grow either in the original tetrad or upon restreaking onto fresh medium; synthetic sick interactions were defined as double mutants that grew notably more slowly than either single gene deletion strain. Two-dimensional hierarchical clustering of dia2Δ synthetic genetic interactions was performed against a data set of 284 synthetic genetic array (SGA) and diploid synthetic lethal analysis by microarray (dSLAM) screens (Tong et al. 2004; Pan et al. 2006; Reguly et al. 2006) using an average linkage clustering algorithm (Eisen et al. 1998) and MapleTree (http://mapletree.sourceforge.net/). Graphical representations of genetic interactions were rendered with the Osprey visualization suite (Breitkreutz et al. 2003). All data are available from the BioGRID interaction database at http://www.thebiogrid.org (Stark et al. 2006).

Microarray analysis:

Genomewide expression profiles were determined for dia2Δ (yMT2078) and isogenic wild-type (yMT1448) strains grown to early log phase (OD 0.2–0.4) at 30°. Polyadenylated RNA was fluorescently labeled with Cy3- or Cy5-conjugated dCTP (Amersham) and dia2Δ and wild-type samples were competitively hybridized against full genome 6200 feature ORF arrays (UHN Microarray Centre, Toronto). All hybridizations were replicated with fluor reversal. Arrays were scanned with a Scanarray 4000 (GSI Lumonics) and images were processed by eliminating corrupted spots, normalizing spot intensity to total Cy5 and Cy3 signal, eliminating spots with low intensity, and averaging duplicate spots (Jorgensen et al. 2004).

Genome instability assays:

Chromosome loss rate of a CFIII-SUP11-HIS reporter fragment was measured as described (Hieter et al. 1985). After growth to saturation overnight in SD–trp–his 2% glucose medium to maintain the artificial chromosome, cells were diluted into SD–trp 2% glucose media, grown for 6 hr at 30° to early log phase (2–4 × 10⁶ cells/ml), sonicated briefly, and plated onto SD–trp–ade 2% glucose agar media supplemented with a limited amount of adenine (10 mg/liter). Chromosome loss rate per generation was calculated as the number of half-sectored colonies divided by the number of colonies. GCR rates were determined with a strain in which HXT13 (~7.5 kb telomeric to CAN1 on Chr V) is replaced by URA3 (Chen and Kolodner 1999). Control (yMT3420) and dia2Δ (yMT3812) revertants that arose from loss of the region encompassing CAN1 and URA3 were scored on canavanine and 5-fluoroorotic acid (5-FOA)-containing medium. Rates were calculated by fluctuation analysis using the method of the median as described (Kanellis et al. 2003) and represent the average of two independent experiments using sets of five independent cultures. Mitotic recombination rates were determined by loss of an ADE2 marker integrated into the rDNA locus on Chr XII as described (Christman et al. 1988). Recombination rate per generation was calculated by dividing the number of half-sectored colonies by the total number of colonies.

Pulsed-field gel electrophoresis and hybridization:

Standard pulsed-field gel electrophoresis (PFGE) procedures were used according to the manufacturer's instructions (Bio-Rad). Agarose plugs containing 3 × 10⁸ cells/ml were loaded onto a 1% agarose gel in 0.5 × Tris–borate EDTA (TBE) buffer and electrophoresed at 6 V at an angle of 120° for 24 hr at 14° with an initial switch time of 60 sec and a final switch time of 120 sec. Gels were stained and photographed and then transferred to nylon membranes and probed with γ-³²P-ATP-labeled 35S rDNA or MCM3 fragments. To assess formation of ERCs, plugs were prepared as above and DNA was resolved on a conventional 1% agarose gel. Gels were transferred to nylon membranes and probed with a γ-³²P-ATP-labeled 35S rDNA fragment.

Genetic interactions of DIA2 with DNA replication, repair, and checkpoint pathways:

To delineate Dia2 function, we employed SGA technology to construct a collection of double mutants between dia2Δ and an ordered array of 4422 viable single gene deletion strains (Tong et al. 2001). Double-mutant combinations that result in inviability (synthetic lethality) or in reduced fitness (synthetic sickness) identify genes that either converge on an essential biological process or exhibit a damage-response relationship (Hartman et al. 2001). The SGA screen identified 55 deletion mutants that exhibited overt synthetic genetic interactions with dia2Δ, all of which were confirmed by tetrad analysis. Of the 55 double-mutant interactions, 19 were synthetic lethal (Figure 2). Many of the interactions occurred with genes implicated in DNA replication (e.g., RRM3, PIF1, RAD27, RNR4), recombination (e.g., RAD52, MRE11, XRS2), chromosome segregation (e.g., BFA1, MAD1, CIK1, KAR3), and the DNA damage checkpoint (PPH3). Significantly, DIA2 also exhibited strong interactions with a spectrum of genes that promote replication fork restart, namely SGS1 and TOP3, SLX1 and SLX4, SLX5 and SLX8, and MUS81 and MMS4; each of these gene pairs encode protein complexes that play different roles in fork restart (Fabre et al. 2002; Fricke and Brill 2003; Zhang et al. 2006). In addition, DIA2 displayed synthetic interactions with genes required for oxidative stress metabolism (e.g., SOD1, LYS7), transcription (e.g., CTK1, SWI4), and chromatin structure (e.g., SWR1, HST4). Recently, a cohort of dSLAM screens focused on DNA replication and chromosome segregation pathways recovered 112 synthetic lethal/synthetic sick interactions with a dia2Δ query strain, and an additional 9 dia2Δ interactions with other query strains (Pan et al. 2006). Of the 55 confirmed hits in our dia2Δ screen, only 23 overlapped with the Pan et al. (2006) data set, indicating that neither experimental approach exhaustively identified dia2Δ genetic interactions. The myriad interactions recovered by each systematic dia2Δ screen suggested that Dia2 function is required to cope with a host of defects in normal replication, repair, and recombination processes, as caused either directly or indirectly by various mutations.

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Figure 2.—

Systematic analysis of dia2Δ synthetic genetic interactions. A total of 55 synthetic lethal (red edges) and synthetic sick genetic interactions (black edges) detected in triplicate SGA screens were confirmed by tetrad analysis. Interactions are grouped according to indicated cellular functions and individual nodes are colored by a reduced hierarchy of gene ontology (GO) biological processes ranked in the order shown in the color key.

Genes that perform related functions tend to cluster together in genomewide synthetic lethal interaction profiles (Tong et al. 2004). We therefore performed two-dimensional hierarchical clustering of the dia2Δ genetic interaction profile in the context of a large combined data set of 10,334 synthetic lethal interactions (Figure 3A), as derived from 284 systematic SGA and dSLAM screens reported in the primary literature (Tong et al. 2004; Pan et al. 2006; Reguly et al. 2006). As revealed by the dominant central cluster, many of the systematic screens carried out to date have focused on DNA replication, DNA damage, and chromosome segregation pathways (Tong et al. 2004; Pan et al. 2006). Within this large data set, the dia2Δ profile clustered most closely with query mutations that disrupt aspects of DNA replication and repair, namely homologous recombination (rad51Δ, rad52Δ, rad54Δ, rad57Δ, hpr5Δ/srs2Δ), the replication checkpoint (csm3Δ, tof1Δ, mrc1Δ), an alternative replication factor C complex implicated in establishment of sister-chromatid cohesion (dcc1Δ, ctf8Δ, ctf18Δ), the Mre11–Rad50–Xrs2 complex that mediates the DNA damage response and facilitates DSB repair (mre11Δ, rad50Δ, xrs2Δ), and postreplicative repair (rad5Δ, rad18Δ). Overall, the correlated genetic profiles revealed by hierarchical clustering suggested a role for Dia2 in some aspect of DNA replication.

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Figure 3.—

Two-dimensional hierarchical clustering of synthetic genetic interaction profiles. (A) A combined unique set of 144 genetic interactions from dia2Δ query screens reported in this study and in Pan et al. (2006) were clustered against 284 systematic genetic screens curated from the primary literature (Reguly et al. 2006). A locally dense region of interactions that contains the dia2Δ profile is expanded and immediate dia2Δ neighbors are indicated by the red bar. The source of each genetic interaction is indicated by the color key. (B) Shared interactions among dia2Δ, rtt101Δ, rtt107Δ,, and mms1Δ. Network shows all interactions retrieved from the full 284-screen data set for each of the four nodes. Edges are colored by interaction source; nodes are colored by the same GO biological processes as in Figure 2.

In addition to the above well-characterized replication and repair functions, dia2Δ clustered immediately adjacent to rtt101Δ, rtt107Δ/esc4Δ, and mms1Δ/rtt108Δ (Figure 3A, red bar). The RTT genes were isolated in a screen for regulators of Ty1 transposition, which uncovered many pathways that dictate genome stability (Scholes et al. 2001). Rtt101 is the cullin subunit of an SCF-like ubiquitin ligase that has recently been implicated in replication fork progression through natural pause sites and damaged DNA (Luke et al. 2006); Rtt107/Esc4 is a BRCT repeat-containing protein that is required for replication restart after DNA damage (Rouse 2004); Mms1 is required for resistance to MMS and other genotoxins (Hryciw et al. 2002) and appears to operate in the same pathway as Mms22 (Araki et al. 2003). Direct inspection of all genetic interactions in this cohort revealed that 21 of 29 interactions with rtt101Δ, 34 of 55 hits with rtt107Δ and 38 of 49 hits with mms1Δ also interact with dia2Δ; in addition, many other cross-interactions were evident, including all possible pairwise synthetic lethal interactions (Figure 3B). These similar genetic profiles suggest that Dia2, Rtt101, Rtt107, and Mms1 functions converge at the replication fork, particularly under adverse circumstances. Because the dia2Δ mutant exhibited a larger and more diverse set of interactions than other members of the cohort (Figure 3B), Dia2 likely performs additional functions as well.

Dia2 is required for resistance to some but not all DNA-damaging agents:

Mutations that compromise S-phase progression often cause sensitivity to DNA-damaging agents or replication stress. Prompted by the synthetic genetic interactions identified in the SGA screen, we tested the sensitivity of a dia2Δ strain to various genotoxic agents, using appropriate checkpoint (mec1Δ sml1Δ), nucleotide excision repair (rad14Δ), and homologous recombination (rad52Δ) mutants as controls (Figure 4A). Plating efficiency of the dia2Δ strain was severely inhibited by the potent alkylating agent MMS. This sensitivity to MMS was most likely due to defective SCF^Dia2 activity because a dia2Δ strain bearing a version of Dia2 that lacked the F-box domain was as susceptible as the dia2Δ strain itself (Figure 4B). Intermediate growth inhibition was caused by 4-NQO, which forms bulky adducts that are substrates for nucleotide excision repair, and by camptothecin, a poison that traps covalent topoisomerase I/DNA intermediates. A high concentration of the replication inhibitor HU modestly reduced the plating efficiency of a dia2Δ strain. The sensitivity of the dia2Δ mutant to MMS, HU, and camptothecin has recently been reported in other studies (Ohya et al. 2005; Koepp et al. 2006; Pan et al. 2006). To determine if these defects reflected a general sensitivity to DNA damage, we also tested sensitivity of the dia2Δ strain to UV light, which induces thymidine dimers and other photo products, and to X rays, which produce DSBs. Unlike many other replication and DNA damage response mutants, the dia2Δ strain was insensitive to these agents (Figure 4A). Dia2 thus does not appear to have a role in nucleotide excision repair or homologous recombination per se, but rather seems to aid in replication fork progression in the presence of certain DNA adducts.

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Figure 4.—

Dia2 is required for resistance to agents that generate DNA adducts. The indicated strains were serially diluted in 10-fold steps and plated onto rich medium (A) or synthetic medium lacking tryptophan (B) in either the presence or the absence of 0.02% (v/v) MMS, 200 mm HU, 5 μg/ml camptothecin, 0.1 μg/ml 4-NQO, 200 J/m² of UV, or 100 Gy of X rays. Photographs were taken after 2 days at 30°.

The DNA damage checkpoint is essential in the absence of DIA2:

As cells lacking Dia2 exhibited a pronounced S and G₂/M cell cycle delay, reliance on nonessential checkpoint proteins recovered in the SGA screen, and hypersensitivity to certain DNA-damaging agents, we determined in further detail if the DNA damage checkpoint was necessary for viability of a dia2Δ strain (Figure 5). Both Mec1 and Rad53 are essential proteins that transduce signals in the DNA damage checkpoint. Viability of rad53Δ and mec1Δ mutant strains, but not their checkpoint deficiencies, is rescued by deleting the ribonucleotide reductase inhibitor SML1 (Zhao et al. 1998). Triple mutants of the genotype dia2Δ mec1Δ sml1Δ and dia2Δ rad53Δ sml1-1 were inviable, indicating that Dia2 normally prevents some form of endogenous DNA damage that invokes a necessary checkpoint response.

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Figure 5.—

The DNA checkpoint is required for viability of a dia2Δ strain. (A) Representative tetrads were dissected for dia2Δ mec1Δ sml1Δ, dia2Δ rad53Δ chk1Δ sml1-1, dia2Δ rad9Δ rad24Δ, and dia2Δ mrc1Δ rad9Δ heterozygous diploids. Wild-type spores are unmarked, as are inviable spores whose genotype could not be inferred. The dia2Δ rad53Δ chk1Δ cross carries an unmarked sml1-1 allele to allow viability of the rad53Δ mutation. Note that, in the last tetrad shown, rad53Δ alone is inviable because sml1-1 did not segregate to this spore clone; however, specific inviability of dia2Δ rad53Δ sml1-1 was inferred from the viability of rad53Δ chk1Δ sml1-1 in the adjacent cross, as well as from other tetrads not shown. The mrc1Δ rad9Δ cross carries a <RNR1 LEU2 2μm> plasmid to maintain viability of the mrc1Δ rad9Δ double mutant. (B) Legend for determined genotypes. (C) Summary of genetic interactions. Minus sign indicates full inviability; plus sign indicates full viability. (D) Failure of various DNA damage checkpoint mutants to bypass the G₂/M cell cycle delay of dia2Δ strains. DNA content of asynchronous cultures of the indicated strains was determined by FACS analysis.

To delineate the checkpoint pathway(s) that maintain dia2Δ viability, we assessed genetic interactions between dia2Δ and mutations in the S-phase (mrc1Δ) and DNA damage (rad9Δ, rad24Δ) branches of the checkpoint. Absence of the fork-associated checkpoint protein Mrc1 moderately reduced the growth rate of the dia2Δ mutant, whereas elimination of the checkpoint adaptor Rad9 and/or the clamp loading factor Rad24 only slightly impaired growth rate (Figure 5, A–C). Loss of both the Mrc1 and the Rad9 branches results in synthetic lethality, but this lethality can be suppressed by overexpression of RNR1 (Alcasabas et al. 2001). As expected, a mrc1Δ rad9Δ dia2Δ triple mutant was inviable despite overexpression of RNR1. These results suggested that while either of the S-phase and DNA damage branches of the DNA checkpoint suffice to allow viability of the dia2Δ mutant, the S-phase branch is the more important of the two arms. Given that checkpoint activation is usually accompanied by a cell cycle delay, we anticipated that the accumulation of cells in G₂/M in the dia2Δ mutant might be dependent on Mrc1 and/or Rad9/Rad24. However, abrogation of either of the main checkpoint branches failed to alter the G₂/M cell cycle delay in the dia2Δ strain, as shown by FACS analysis (Figure 5D). This delay thus either requires both arms of the checkpoint or is mediated by an as yet uncharacterized checkpoint effector.

Interestingly, as recovered in the SGA screen, the dia2Δ strain also requires the phosphatase Pph3 for viability (Figure 2). Because Pph3 is required for dephosphorylation of γ-H2Ax and recovery from the checkpoint response (Keogh et al. 2006), persistent DNA replication defects in dia2Δ cells may cause permanent checkpoint arrest in the absence of desensitization.

Dia2 prevents endogenous DNA damage in S-phase:

The requirement for Mec1, Rad53, and Mrc1 for viability of the dia2Δ mutant and the accumulated S-phase fraction in asynchronous dia2Δ cultures (Figure 1D) suggested a possible S-phase progression defect in dia2Δ strains. We therefore assessed the kinetics of S-phase progression in synchronous cultures released from a mating pheromone-induced G₁ phase arrest. While most of the dia2Δ population appeared to progress from G₁ phase to G₂/M phase with wild-type kinetics, a significant subpopulation accumulated in S-phase (Figure 6A). Signals that trigger the replication checkpoint result in Mec1-dependent activation of Rad53 (Sanchez et al. 1996; Sun et al. 1996), which can be monitored by an ISA (Pellicioli et al. 1999). Given that dia2Δ cells have a cell cycle delay, we tested whether this defect correlates with activation of Rad53 kinase activity. As expected, upon treatment of wild-type cells with 200 mm HU, Rad53 was activated, as revealed by its autophosphorylation in vitro (Figure 6B). However, in dia2Δ strains, Rad53 exhibited marked activation in untreated asynchronous cultures. Exposure to 200 mm HU further induced Rad53 autophosphorylation in dia2Δ cells, possibly because fewer cycling cells are in S-phase as compared to cultures arrested in HU. Because MMS and 4-NQO also further activated Rad53 in the dia2Δ strain (data not shown), Dia2 is not required for maximal activation of the checkpoint. To determine whether checkpoint activation in dia2Δ cells required S-phase progression, we monitored Rad53 phosphorylation in synchronous cultures by virtue of its phosphorylation-dependent mobility shift (Figure 6C). In the absence of genotoxic stress, wild-type cells passed through S into G₂ without Rad53 phosphorylation. In contrast, while Rad53 was unphosphorylated in G₁ phase in the dia2Δ strain, as cells progressed through S-phase and into G₂ phase, slower migrating Rad53 isoforms became apparent. This result suggested that Dia2 normally prevents endogenous DNA damage from arising in S-phase.

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Figure 6.—

Activation of the DNA damage response in the dia2Δ strain. (A) Cell cycle progression of unperturbed wild-type and dia2Δ strains. Cells were arrested in G₁ phase with α-factor and released into rich medium. DNA content was assessed by FACS at the indicated time points in minutes. (B) Rad53 kinase activity was detected using an ISA assay on lysates of the indicated strains grown in either the presence or the absence of 200 mm HU for 1 hr. Arrow indicates in situ ³²P incorporation into Rad53. (C) Cell cycle dependence of Rad53 activation. The indicated strains were arrested with α-factor and released into rich medium and lysates were prepared at the indicated time points. Immunoblots were probed with a polyclonal Rad53 antibody. (D) Cell cycle progression of MMS-treated wild-type and dia2Δ strains. Cells were arrested in G₁ phase with α-factor, released into rich medium containing 0.033% MMS, and assessed for DNA content at the indicated time points. (E) DNA damage repair foci in a dia2Δ strain. Rad52^YFP was detected by fluorescence microscopy in wild-type and dia2Δ cells. A composite of 21 collapsed Z-sections was taken for a minimum of 300 cells/strain, which were classified as unbudded (G₁) and budded (S/G₂/M) cells. Bars indicate standard error.

Despite the ability of a dia2Δ strain to activate Rad53 in response to HU, MMS, and 4-NQO, the strain is nevertheless sensitive to MMS. To examine whether this sensitivity might result from a partially defective S-phase checkpoint, G₁ phase cultures were released into medium containing a low dose of MMS, which activates the intra-S-phase checkpoint but not the G₁ DNA damage checkpoint (Sidorova and Breeden 1997). Wild-type cells required at least 210 min to complete replication in the presence of MMS, ~175 min longer than without DNA damage. In contrast, MMS-treated dia2Δ cells were unable to progress past mid-S-phase for the duration of the 270-min experiment (Figure 6D). As checkpoint defective cells normally progress through S-phase faster than wild-type cells (Paulovich and Hartwell 1995), sensitivity of the dia2Δ strain to MMS was not a consequence of a defective intra-S checkpoint, but rather due to an inability to recover from MMS-induced DNA damage.

The S-phase-dependent activation of Rad53 suggested that dia2Δ cells accumulate endogenous DNA damage. This idea is also supported by the genetic result that RAD52-dependent DSB repair is required for viability of dia2Δ strains (Figure 2), whereas in direct tetrad analysis, the Ku70/Ku80 nonhomologous end joining factors required to repair breaks in G₁ phase are not required (data not shown). To directly assess the occurrence of DNA strand breaks in dia2Δ cells, we measured the occurrence of DNA repair foci in live cells using YFP fusions to DNA-damage-responsive proteins (Lisby et al. 2001; Melo et al. 2001). As expected, Rad52^YFP foci formed at a low frequency in log-phase cultures of wild-type cells. In contrast, the dia2Δ mutant showed a significant increase in foci formation of 2- and 4.5-fold in unbudded and budded cells, respectively (Figure 6E). A similar increase was seen by both a TUNEL assay for DNA strand breaks and a Ddc1^YFP damage foci reporter (data not shown). This increase in repair foci implies that DNA lesions, most likely DSBs, accumulate in dia2Δ cells and activate the DNA checkpoint response. The slight increase of Rad52 foci in G₁ phase dia2Δ cells suggested an additional defect, perhaps due to incomplete repair of lesions before desensitization of the G₂/M checkpoint arrest. However, we note that the extent of G₁ phase damage foci in dia2Δ cells is modest compared to other DNA repair/checkpoint mutants, such as rmi1Δ (Chang et al. 2005).

Dia2 ensures high-fidelity chromosome segregation:

We examined the dia2Δ phenotype by genomewide expression profiles, which can often be used to deduce gene function (Hughes et al. 2000a). The overall expression profile of the dia2Δ strain was highly similar to that of wild-type cells, with the notable exception of a 1.5- to 3.5-fold increase in ribonucleotide reductase (RNR) transcripts in the dia2Δ mutant (Figure 7A). RNR induction is a hallmark checkpoint response to replication stress (Elledge and Davis 1990). In addition, several general stress responsive genes, including XBP1 and HSP26, were elevated in the dia2Δ strain. Aside from the obvious transcriptional difference in RNR transcript levels, comparison of chromosome-wide mean expression levels revealed that two independent dia2Δ isolates, one haploid and one diploid, exhibited significantly higher levels of gene expression from Chr I and Chr X, respectively (Figure 7B). This result indicates that these dia2Δ isolates were aneuploid, consistent with previous detection of aneuploidy in other dia2Δ isolates (Hughes et al. 2000b).

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Figure 7.—

Dia2 is required to ensure faithful chromosome transmission. (A) Induction of RNR transcripts in a dia2Δ strain. Two replicate dia2Δ haploid isolates were competitively hybridized against wild-type control mRNA on genomewide microarrays (5989 ORFs detected) and plotted against one another. The top 10 induced genes in the dia2Δ strain (fold increase indicated in parentheses) were MET17 (6.5×), GPH1 (4.2×), HSP26 (3.4×), RNR4 (2.9×), YGP1(3.0×), RNR2 (3.4×), HSP12 (3.0×), YMR250w (3.1×), XBP1 (2.3×), and LAP4 (2.1×). Signals for RNR genes in replicate dia2Δ hybridizations are shown separately. (B) Aneuploidy in dia2Δ mutants detected by microarray analysis. Median gene induction per chromosome was calculated for independent haploid and homozygous diploid dia2Δ isolates. (C) Chromosome loss rates. Wild-type and dia2Δ strains bearing an artificial test chromosome (CFIII-SUP11-HIS3) were plated onto nonselective media for 2 days and scored for half red/white colonies (i.e., first division missegregation events). At least 2400 colonies were scored per strain.

To directly test whether dia2Δ plays a role in chromosome segregation, the rate of chromosome loss was measured by a colony-sectoring assay (Hieter et al. 1985). In this assay, cells carry a reporter chromosome fragment with the SUP11 (ochre-suppressing tRNA) gene, which is used to complement an ade2-101 (ochre) mutation. Colonies that retain the test chromosome are white, whereas cells that undergo a loss event accumulate a red adenine biosynthesis intermediate. The dia2Δ mutation caused an ~200-fold increase in chromosome loss rate relative to wild type (Figure 7C). In comparison, the spindle checkpoint mutants bub1Δ and bub3Δ exhibit only a 50-fold increase in chromosome loss rate under identical conditions (Warren et al. 2002). Dia2 thus plays a significant role in ensuring the fidelity of chromosome transmission, likely through suppression of replication-induced DSBs, which can lead directly to aneuploidy through missegregation events (Kaye et al. 2004).

Dia2 suppresses GCRs:

DNA double strand breaks can lead to whole or partial chromosome loss (Kaye et al. 2004). We therefore tested whether dia2Δ cells are prone to genomic alterations by using a mutator assay that measures the incidence of GCRs (Chen and Kolodner 1999). Rearrangements are detected in haploid cells by the simultaneous loss of CAN1 and URA3 from the left arm of Chr V, such that cells lacking CAN1 and URA3 are able to form colonies on medium containing 5-FOA and canavanine (can). Strikingly, deletion of dia2Δ in this reporter strain caused a 117-fold increase in GCR rate (Figure 8A). This rate is comparable to that reported for S-phase checkpoint mutants (Myung et al. 2001).

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Figure 8.—

Increased GCR rate is correlated with hyperrecombination at the rDNA locus in dia2Δ cells. (A) Assay used to detect gross chromosomal rearrangements. Sensitivity to canavanine (can) and 5-FOA selects for colonies that have undergone simultaneous loss of both genes via a GCR event. GCR rates were from two independent experiments; fold induction was calculated as the mean of dia2Δ GCR rate divided by the mean GCR rate of the parental control strain. (B) Chromosomal DNA from a control and seven independent can^r 5-FOA^r dia2Δ isolates was resolved by PFGE, stained with ethidium bromide, and probed with a Chr V-specific MCM3 sequence. (C) Chromsomal DNA from five of the same isolates was rerun and probed with a 35S rDNA sequence. (D) Recombination at the rDNA locus. Wild-type and dia2Δ strains bearing an ADE2 marker at the rDNA locus were plated onto rich medium and grown for 3 days. Recombination rates were calculated from counting first division missegregation events for at least 20,000 colonies. Bars indicate standard error. (E) Accumulation of ERCs. Genomic DNA isolated from each of the indicated strains was resolved by electrophoresis and probed with rDNA sequences. Asterisk indicates chromosomal rDNA and arrows point to mono- and multimeric ERCs (red and black arrows, respectively). (F) Subcellular localization of Dia2^GFP. A wild-type strain expressing Dia2^GFP was grown to log phase and visualized by fluorescence microscopy. Dia2^GFP signal that segregates late in anaphase as a string of fluorescence stretched across the bud neck is indicated by the arrow. Numbers in parentheses indicate the percentage of cells that show nucleolar Dia2 localization at each cell cycle stage.

Three classes of rearrangements are detected by the GCR assay: interstitial deletions, nonreciprocal translocations, and partial deletions of Chr V with de novo telomere addition (Kolodner et al. 2002). To characterize the spectrum of GCRs that occur in dia2Δ strains, we analyzed Chr V structure by PFGE. Chromosomal DNA from the control parental strain and seven independent 5-FOA^r can^r dia2Δ isolates was resolved and probed with a Chr V-specific MCM3 sequence (Figure 8B). In six of the seven dia2Δ GCR isolates, MCM3 sequences migrated slightly faster than in the control lane, presumably due to a partial chromosome deletion. In addition to these discrete chromosomal rearrangements, each dia2Δ strain tested retained a significant amount of chromosomal DNA in the well, suggestive of unresolved replication and recombination structures (Desany et al. 1998).

We thank Jef Boeke, Xuewen Pan, Grant Brown, and Patrick Paddison for helpful discussions and communication of unpublished results; Rodney Rothstein, Charlie Boone, Hannah Klein, and Howard Bussey for providing reagents; and Nizar Batada and Jan Wildenhain for advice on data analysis. D.B. was supported in part by a Canadian Institutes of Health Research (CIHR) Doctoral Award. P.K. is a recipient of a studentship from the National Cancer Institute of Canada. D.D. and M.T. are supported by grants from the CIHR and are holders of Canada Research Chairs. M.P. was supported by grants from the Swiss Federal Institute of Technology and the Swiss National Science Foundation.