Rad52-Independent Mechanisms

Molecular Cell, Vol. 12, 1499–1510, December, 2003, Copyright 2003 by Cell Press

Branch Migrating Sister Chromatid Junctions Form at Replication Origins through Rad51/Rad52-Independent Mechanisms Massimo Lopes,1,2,3 Cecilia Cotta-Ramusino,1,2 Giordano Liberi,1,2 and Marco Foiani1,2,* 1 Istituto F.I.R.C. di Oncologia Molecolare Via Adamello 16 20141 Milano Italy 2 Dipartimento di Scienze Biomolecolari e Biotecnologie Universita` degli Studi di Milano 20122 Milano Italy

Summary Cells overcome intra-S DNA damage and replication impediments by coupling chromosome replication to sister chromatid-mediated recombination and replication-bypass processes. Further, molecular junctions between replicated molecules have been suggested to assist sister chromatid cohesion until anaphase. Using two-dimensional gel electrophoresis, we have identified, in yeast cells, replication-dependent X-shaped molecules that appear during origin activation, branch migrate, and distribute along the replicon through a mechanism influenced by the rate of fork progression. Their formation is independent of Rad51- and Rad52mediated homologous recombination events and is not affected by DNA damage or replication blocks. Further, in hydroxyurea-treated rad53 mutants, altered in the replication checkpoint, the branched molecules progressively degenerate and likely contribute to generate pathological structures. We suggest that cells couple sister chromatid tethering with replication initiation by generating specialized joint molecules resembling hemicatenanes: this process might prime cohesion and assist sister chromatid-mediated recombination and replication events. Introduction Eukaryotic DNA replication is a complex process that has to be highly coordinated with cell cycle progression, checkpoints, transcription, recombination, repair, sister chromatid cohesion, and chromatin remodeling (Bell and Dutta, 2002; Aguilera, 2002; Gerbi and Bielinsky, 2002; Nasmyth, 2001; Osborn et al., 2002) to ensure the faithful transmission of genetic information to daughter cells. A failure in tuning together these cellular pathways may generate pathological DNA structures, chromosome lesions, mutations, genome instability, cell death, and cancer (Hickson, 2003; Kolodner et al., 2002). DNA replication, by itself, represents a dangerous event for the cell as chromosome lesions may arise due to the action of nick and closing enzymes (i.e., DNA topoisomerases and ligases), and nucleotides are often *Correspondence: [email protected] 3 Present address: Institute of Cell Biology, ETH Ho¨nggerberg, CH8093 Zu¨rich, Switzerland.

misincorporated during DNA synthesis (Kornberg and Baker, 1992). Further, the replisome often has to deal with repetitive sequences on the template that might lead to slippage of the newly synthesized chains, with highly transcribed chromosomal regions that slow down DNA synthesis or with genomic regions known as fragile sites that represent hot spots for genomic rearrangements (Deshpande and Newlon, 1996; Ivessa et al., 2000; Cimprich, 2003). The situation is even more dramatic when cells experience DNA damage while they are replicating the genome: a damaged template represents a barrier for the replisome leading to transient replication blocks (Page`s and Fuchs, 2003). Several cellular pathways have been implicated in the repair of DNA damage during replication; most of them employ sister chromatids to allow the exchange of genetic information through recombination, to restart replication following double-strand break formation or even as a potential template for replication bypass processes (Paques and Haber, 1999; Kadyk and Hartwell, 1992; Gonzales-Barrera et al., 2003; Malkova et al., 1996; Higgins et al., 1976). These repair processes could be highly influenced by the establishment of sister chromatid cohesion that contributes to keep sister chromatids in close proximity until chromosome segregation takes place through the formation of proteinaceous ring-like structures and, possibly, by generating yet unidentified topological interlocks between DNA sister molecules (Nasmyth, 2001; Murray and Szostak, 1985, and references therein; Koshland and Hartwell,1987). The two-dimensional neutral/neutral agarose gel electrophoresis (2D gel) method (Brewer and Fangman, 1987) has been extensively used to describe conformational changes of replication intermediates implicated in the initiation, elongation, and termination steps of DNA synthesis as well as of recombination-related structures. In recent years, replication-dependent fourways branched (X-shaped) molecules have been observed by 2D gels and consistently attributed to recombinative Holliday junctions. They have been described during meiotic S phase where recombination is known to accompany DNA replication (Collins and Newlon, 1994; Borde et al., 2000; Schwacha and Kleckner, 1994), or during mitotic chromosome replication at amplified loci, where homologous regions might easily prime recombination events (Delidakis and Kafatos, 1989; Heck and Spradling, 1990; Dijkwel et al., 1991; Liang et al., 1993; Zou and Rothstein, 1997). Recently, X-shaped molecules have been also associated with mitotic chromosome replication of single copy regions, in Physarum polycephalum (Be´nard et al., 2001) and in S. pombe (Segurado et al., 2002); in both cases, these structures have been connected with recombination events, although in Physarum no genetic approach was possible. Further, in S. pombe, it has been shown that accumulation of these X molecules at replication origins was prevented in mutants defective in homologous recombination (Segurado et al., 2002). These intra-S recombination structures have been suggested to play a role in assisting specialized replication-coupled repair pathways

Molecular Cell 1500

or in promoting replication restart processes (Zou and Rothstein, 1997; Be´nard et al., 2001; Segurado et al., 2002). Recombination-dependent X molecules have been described also at replication forks in RecQ-helicase-defective cells experiencing intra-S DNA damage (G.L. and M.F., unpublished data), and Holliday junctionlike structures (reversed forks) have been shown in checkpoint mutants exposed to replication blocks (Sogo et al., 2002). In the latter cases, though, these replication-related recombination or pseudo-recombination events are thought to contribute to genome instability. In this study, we describe a species of intra-S fourways branched molecules that form during an early step of DNA synthesis, at origins of replication, through mechanisms independent of Rad51 and Rad52-mediated homologous recombination. These chromosomal structures are able to branch migrate but do not resemble any of the intermediates described so far. Their dispersion along newly replicated DNA is strictly related to the rate of fork progression, and their stability when forks stall depends upon a functional checkpoint. Our results suggest that newly replicated chromatids are transiently connected by physical junctions via a recombination-independent mechanism. These joint structures resemble hemicatenanes that might tether sister chromatids, modulate their metabolism under physiological conditions, and contribute to the pathological formation of abnormal structures in checkpoint-defective backgrounds. Results We have analyzed the quality and progression of replication forks during unperturbed chromosomal replication in S. cerevisiae. DNA replication intermediates (RIs) were separated according to their mass and shape by neutralneutral two-dimensional gel electrophoresis (2D) (Brewer and Fangman, 1987); we have focused our study on Chromosome III on those forks that arise from the early origin of replication ARS305 and invade adjacent regions that also contain the dormant origins ARS301-304 (Newlon et al., 1993; Figure 1). In wild-type (wt) cells released from a G1 block under unperturbed conditions, bubbles and large Ys accumulate at ARS305 within 25–30 min (Figure 1). The bubble arc represents origins that have been fired bidirectionally, while large Y molecules result from asymmetric progression of replication forks out of the restriction fragment containing ARS305. Forks progressively invade adjacent regions as shown by the appearance of the Y arc on restriction fragments A–D. Region A (positioned at 5 kilobases [kb] to the left of ARS305) is replicated at 30 min, while regions B and C (16 kb to the left and 17 kb to the right of ARS305, respectively) at 45 min, and region D (26 kb to the left) at about 60 min. Region C is equidistant from the early origins ARS305 and ARS306 and, consequently, also exhibits double Y-shaped intermediates, resulting from forks converging from the two replicons. X-shaped structures can also be detected on ARS305 fragment: they begin to accumulate at 30 min when the bubble arc is clearly visible, they reach the maximal accumulation at 45 min when bubbles start to decrease in intensity, and they decrease at 60 min; at later time

points, they completely disappear from ARS305 fragment (data not shown). The relative intensity of the X spike on ARS305 is similar to those of bubble and Y molecules. X molecules can be also visualized on region A, although they appear 15 min later than the Y arc and their relative intensity is lower compared to that of the Y arc on region A and that of the X spike on ARS305 throughout the time course. No X molecules were detected on regions B, C, and D. We found that the visualization of the X structures is particularly enhanced when replication intermediates are prepared using conditions that restrain branch migration of joint molecules during in vitro manipulation (Allers and Lichten, 2000; see also Experimental Procedures) (Figure 2A, Method 1). Conversely, purification procedures that employ columns to enrich DNA intermediates (Figure 2A, Methods 2 and 3) and glass beads to break the cells and isolate the nuclei (Figure 2A, Method 3) minimize the detection of the X structures. This finding suggests that the X molecules are particularly labile and/or undergo in vitro transitions that do not depend upon DNA synthesis but, rather, resemble branch migration events that might shift the junctions outside the restriction fragment of interest, preventing their visualization by southern. We therefore tested whether the X molecules can indeed experience branch migration. Following the first-dimension gel electrophoresis, agarose slices were treated under conditions that allow branch migration (see Experimental Procedures) and, subsequently, processed for electrophoresis in the second dimension (Figure 2B). Incubation of the intermediates under branch migration conditions results in the progressive decrease of X molecules and in the concomitant accumulation of linear intermediates within a section of the linear line that, horizontally, spans the region occupied by the population of X molecules. We therefore conclude that the X molecule may represent cruciform DNA structures able to branch migrate in vitro. The result shown in Figure 2B suggests the obvious hypothesis that the X molecules represent recombination structures. Recombination-dependent X-shaped chromosomal structures have been described during mitotic DNA replication in S. cerevisiae at the rDNA locus (Zou and Rothstein, 1997), at damaged replication forks in sgs1 cells (G.L. and M.F., unpublished data), and in S. pombe at replication origins (Segurado et al., 2002). The four-way structure of the Holliday junctions, in the presence of Mg2⫹ ions folds by pairwise coaxial stacking of helices into the stacked X structure that is unable to efficiently branch migrate (Duckett et al., 1988, 1990; Lilley, 1997). We therefore tested whether the branch migration of the X molecules was prevented by the addition of Mg2⫹ ions using the method described in Figure 2B. We found that the kinetics of branch migration of the X molecules was not affected by the presence of Mg2⫹ ions: after 1 hr incubation in branch migration buffer, almost 50% of molecules are converted into linear structures regardless of the presence of Mg2⫹ ions (Figure 2C) and, by 4 hr, nearly 90% of the intermediates are in the linear conformation (data not shown). This result suggests that the joint molecules are not Holliday junctions in the stacked conformation. To further confirm this observation, we tested whether the formation of X molecules was dependent upon Rad51- and/or

Sister Chromatid Junctions at Replication Origins 1501

Figure 1. Joint DNA Molecules Form at ARS305 Replication Origin ARS305 and A–D represent the restriction fragments analyzed; restriction sites are indicated as follows: EcoRV (E), HindIII (H), NcoI (N). W3031A cells were presynchronized by ␣ factor (␣F) treatment in G1 and released into fresh YPD; DNA was prepared from cells collected at the indicated times using “CTAB extraction” (see Experimental Procedures), cut with EcoRV and HindIII, and analyzed by 2D gel, by sequential hybridization of the same membrane with the different probes.

Rad52-dependent recombination pathways. We found that in rad51⌬ and rad52⌬ cells, released from a G1 block into S phase under unperturbed conditions, replication intermediates and X molecules accumulate on ARS305 and region A with kinetics similar to those of wt cells (Figure 3; also compare with Figure 1). We conclude that, differently from S. pombe (Segurado et al., 2002), in S. cerevisiae homologous recombination is not required for origin firing. Further, our results strongly suggest that the X structures are not Holliday junctions and do not derive from Rad51- or Rad52-mediated recombination processes. We have therefore identified chromosomal branched intermediates that, on 2D gels, separate like Rad51/ Rad52-dependent recombination structures (Zou and Rothstein, 1997; G.L. and M.F., unpublished data) although their formation does not require homologous

recombination but depends upon origin-dependent initiation of DNA synthesis. These X-shaped structures seem to specifically form at the origins of replication and then resolve as forks move far from the origin region, possibly as a result of differences in topological constraints behind replicating forks on newly replicated regions positioned far from the origin fragment (Olavarrieta et al., 2002a). Alternatively, they could branch migrate independently of the movement of the forks and asynchronously scatter throughout the newly replicated regions, between the forks and the origins of replication. In both cases, we expected that the branched molecules would be present between two diverging forks within an elongating bubble (Figure 6A); thus, we reasoned that their turnover and/or migration extent would be affected by changes in the rate of fork elongation.

Molecular Cell 1502

Figure 2. The Joint DNA Molecules Are Sensitive to the DNA Extraction Procedure and Able to Branch Migrate Samples were collected 45 min after the ␣F release using the conditions described in Figure 1. In (A), three samples were processed following different DNA extraction procedures (see Experimental Procedures); 2D gels were hybridized with the ARS305 probe. In (B), three DNA samples were extracted using the conditions described in Figure 1; DNA were digested with EcoRV and HindIII and subjected to electrophoresis in the first dimension; one of the agarose lanes was then processed for the second dimension under standard conditions (Control), while the other two were incubated in branch migration buffer at 65⬚C for 2 or 4 hr before running the second dimension. In (C), agarose slices from the first dimension gel were incubated prior to the second dimension in branch migration buffer, either in the presence or in the absence of 10 mM MgCl2 (see Experimental Procedures). Quantification of the signals is presented.

To test this hypothesis, we investigated the replication timing and the dynamics of formation/resolution/migration of X-shaped molecules of cells treated with sublethal concentrations of methyl methan sulphonate (MMS), a DNA damaging agent that does not affect the timing of early origin activation, but slows down the rate of fork elongation due to the presence of a damaged template (Paulovich and Hartwell, 1995; Tercero and Diffley, 2001). In wt cells, released from G1 in the presence of MMS, ARS305 fires at 20–40 min and the forks clear the origin fragment very slowly, not before 120 min (Figure 4A). Accordingly, the kinetics of invasion of regions B, C, and D were more delayed in time compared to untreated conditions, with the first forks reaching these regions on average 15–20 min later and replication being completed only at about 180 min (Figures 1 and 4A and data not shown). X-shaped molecules appear on the origin fragment as soon as ARS305 activation is detectable (40

min), and their relative amount is similar to untreated conditions (Figure 4A). Also in this case the formation of X-shaped molecules is RAD51- and RAD52 independent (Supplemental Figure S1 [http://www.molecule.org/ cgi/content/full/12/6/1499/DC1]). The disappearance of X-shaped intermediates from the origin region is significantly delayed compared to unperturbed replication, as a large fraction of this signal is still present on ARS305 90 min after the release, when, usually, untreated cells have completed S phase and approach cell division. Forks invading region A persist until 90–120 min, together with a faint signal corresponding to X-shaped molecules. Still 120 min after the release, some replicating forks are traversing regions B, C, and D, and low levels of X-shaped molecules are now detectable even 15 kb far from the origin on both sides, on regions B and C (Figure 4A). Thus, the presence of a damaged template not only

Sister Chromatid Junctions at Replication Origins 1503

Figure 3. Formation of Joint Molecules at ARS305 is RAD51- and RAD52 Independent rad51⌬ (CY2269) and rad52⌬ (CY2272) strains were released from G1 into fresh medium, under unperturbed conditions. DNA samples were processed as in Figure 1 and 2D gels hybridized to detect ARS305 and region A. Note that in these strains the activation of ARS305 is detectable 5–10 min earlier than in wt cells likely due to the presence of very large cells that need less time to reach the critical mass to enter S phase and therefore activate early origins of replication slightly in advance. The kinetics of formation and resolution of joint molecules is anticipated accordingly.

slows down fork progression but also delays the disappearance of the X molecules within a specific restriction fragment; further, under these conditions, the branched molecules can be detected on a broader region of the replicon. To further investigate the relative dependency between the movement of the branched molecules and fork progression, we have analyzed cells exposed to a replication block induced by hydroxyurea (HU) treatment. G1 cells released into S phase in the presence of HU arrest replication and activate the checkpoint. Forks under these conditions stall and accumulate stable replisome-fork complexes and short regions of ss-DNA coated with RPA (Sogo et al., 2002; Lucca et al., 2003; Zou and Elledge, 2003). We found that in HU-treated wild-type cells, X molecules can still be visualized and remain stable throughout the treatment (Figure 4B). This result indicates that fork arrest in wt cells prevents branch migration of the X molecules outside the origin restriction fragment. Rad53 has been implicated in preventing replication fork collapse in response to replication blocks and intra-S DNA damage and accumulation of abnormal intermediates that resemble Holliday junctions (Lopes et al., 2001; Tercero and Diffley, 2001; Sogo et al., 2002). We therefore tested whether Rad53 could influence the X molecule dynamics. Untreated rad53 cells exhibit a replication profile and kinetics of formation/disappearance of X-shaped molecules similar to wt cells, with the exception of a slight anticipation (5–10 min) of ARS305 activation (Supplemental Figure S2 [http:// www.molecule.org/cgi/content/full/12/6/1499/DC1]).

In MMS-treated rad53 cells, ARS305 is fired earlier then in wt cells and is already detectable at 20 min; as a result, the invasion of the closest region (region A) is slightly anticipated compared to wt (Figure 5A). Nevertheless, forks appear to invade regions B and C with a kinetic similar to wt cells: this is likely the result of collapse of a significant number of forks leading to a delay in the invasion of downstream regions by the bulk of DNA replication. This is consistent with the finding that, while in MMS-treated wt cells most of the forks migrate out of the origin fragment within circa 60 min since origin activation, in rad53 cells forks are clearly detectable on ARS305 fragment even 100 min after origin activation, exhibiting a “complete Y arc” pattern which resembles the fork collapse-phenotype already described in HUtreated rad53 cells (Lopes et al., 2001). Further, as a fraction of forks remains stacked on ARS305 and region A, the same seems to be true for the X molecules, strengthening the idea that X molecule migration requires efficient fork progression. In MMS-treated rad53 cells, dormant origins of replication exhibit an unscheduled firing (Shirahige et al., 1998); a weak bubble ark is indeed detectable at 60 min on region D (containing the ARS301 dormant origin) in MMS-treated rad53 but not wt cells (Figure 5A, asterisk; Figure 4A). Unprogrammed firing of ARS301 in rad53 cells results also in accumulation of X-shaped molecules, clearly detectable 30 min after origin firing (Figure 5A arrows); conversely, no X molecules were detected in wt cells during passive replication of the same region (Figure 4A). Thus, this result reinforces the link between origin activation and forma-

Molecular Cell 1504

Figure 4. Kinetics of DNA Replication and Joint Molecules Distribution in MMS- and HU-Treated Wild-Type Cells (A) W303-1A cells were presynchronized by ␣F treatment in G1 and released into fresh YPD containing 0.033% MMS. Samples were collected at the indicated times and processed as in Figure 1. (B) Presynchronized W303-1A cells were released in YPD containing 0.2 M HU. DNA was extracted as in Figure 1 from samples collected at the indicated times, digested with NcoI, and run in 2D gels. Membranes were hybridized with the ARS305 probe.

tion/accumulation of X-shaped molecules during S. cerevisiae chromosomal replication. Further, we can also conclude that X molecule formation does not require Rad53. Rather, the presence of a functional Rad53 ki-

nase in MMS-treated cells influences the number of forks actively engaged in replication elongation and, consequently, also the migration of a fraction of X molecules.

Sister Chromatid Junctions at Replication Origins 1505

Figure 5. Kinetics of DNA Replication and Joint Molecules Distribution in MMS- and HU-Treated rad53 Cells (A) rad53-K227A (CY2034) cells were presynchronized by ␣F treatment in G1 and released into fresh YPD containing 0.033% MMS. Samples were collected at the indicated times and processed as in Figure 1. (B) Presynchronized rad53-K227A cells were released in YPD containing 0.2 M HU. Samples were collected at the indicated times and processed as in Figure 4B.

Finally, we analyzed the fate of X molecules in HUtreated rad53 cells (Figure 5B). rad53 cells were released from G1 into S phase in the presence of HU. At 30 min from G1 release, rad53 forks exhibit a 2D profile similar to wt cells. However, with time, forks progressively de-

generate and accumulate small Ys and a cone signal as a result of fork collapse. The population of X molecules seems to follow the fate of the rest of the intermediates: at 30–60 min they form a well-defined spike, but later on (90–120 min) the spike becomes more diffuse and,

Molecular Cell 1506

concomitantly, a population of intermediates migrating as a cone appears (Figure 5B, arrows); the cone signal likely represents reversed forks (Lopes et al., 2001; Sogo et al., 2002) and, differently from the X spike, is particularly stable as it can be detected regardless of the method used to extract the DNA intermediates (data not shown), likely because the homology of the regressed arm is able to stabilize reversed forks. A possible explanation for this observation is that in HU-treated rad53 cells X molecules migrate very fast outside the restriction fragment. However, we can exclude such a possibility, as we could not observe a progressive accumulation of X molecules on adjacent regions (data not shown). Hence, we conclude that in rad53 cells either the turnover of X molecules is higher compared to wt cells or that they are progressively converted into other structures. Discussion Chromosomal X-shaped DNA structures on 2D gels have been consistently related to Holliday junctions and therefore connected with recombination events. In S. cerevisiae, X molecules can be visualized during the meiotic cell cycle (Collins and Newlon, 1994; Schwacha and Kleckner, 1994) and their appearance has been temporally and spatially related to recombination hot spots (Schwacha and Kleckner, 1994). X molecules have also been detected at mitochondrial DNA (Brewer et al., 1988) and at the rDNA locus (Zou and Rothstein, 1997), and, in both cases, their accumulation was dependent upon recombination pathways. Further, X structures can be visualized in RecQ helicase-defective cells at damaged replication forks and their accumulation can be rescued in rad51⌬ and rad52⌬ strains (G.L. and M.F., unpublished data). DNA replication-dependent X molecules have been described in Physarum during S phase and interpreted as Holliday junctions (Be´nard et al., 2001); in S. pombe, X-shaped recombination structures have been identified at replication origins (Segurado et al., 2002). Using 2D gels, we have identified a species of X molecules that, based on physical and genetic evidence, cannot be classified as recombination structures. The main characteristics of these molecules are the following: (1) their formation occurs under physiological conditions during mitotic S phase and depends upon firing of replication origins; (2) although they migrate on 2D gels like recombination structures their formation does not require Rad51- or Rad52-mediated recombination pathways and, therefore, differ from the structures described in S. cerevisiae and S. pombe; (3) they are able to branch migrate and their movement depends on fork progression; (4) they can be distinguished by Holliday junctions based on their ability to branch migrate in the presence of metal ions, the genetic requirements, the relative stability, and the visualization procedure; (5) their formation is neither prevented nor enhanced by the presence of MMS or HU; conversely, in Physarum X-molecule formation was prevented by HU treatment (Be´nard et al., 2001); (6) they rapidly disappear when exposed to replication blocks in the absence of a functional checkpoint. Hence, despite some similarities with Holliday junctions, these joint molecules likely represent recombination independent X structures.

A tantalizing possibility is that these branched molecules represent hemicatenanes structures (Figure 6A). Hemicatenanes are X-shaped jointed molecules in which one strand of a duplex is coiled around one strand of the other duplex (Figure 6A); on 2D gels, they have been shown to migrate like Holliday junctions (Lucas and Hyrien, 2000). They are not recombination molecules and their structure does not require base pairing, although, in theory, they could be converted into structures resembling pseudo double Holliday junctions by generating pairing between the two newly synthesized strands (Figure 6A, Schwacha and Kleckner, 1995). Further, not only hemicatenanes would be able to branch migrate (Bianchi et al., 1983; Figure 6A) but the two strands engaged in the coiled junction could even slide relative to each other and, as a result, two sequences that share no homology could be coiled together (Lucas and Hyrien, 2000; Figure 6A); hemicatenane formation would not require Rad51- or Rad52-dependent processes, although it is formally possible that under certain conditions or in certain genetic backgrounds hemicatenanes could be converted into pseudo-recombination structures and therefore engaged by recombination proteins. Further, differently from Holliday junctions and consistently with our data, it is expected that the presence of metal ions will not prevent branch migration of the hemicatenane structure as it cannot be converted in the stacked conformation. Hemicatenanes are expected to be more labile than Holliday junctions: a single nick might resolve hemicatenanes but not Holliday junctions that would be still stabilized by base pairing; this consideration might explain the observation that certain harsh procedures used to prepare DNA intermediates minimize their visualization (Figure 2A). We observed that, once formed, the migration of the X structures is not coordinated with the movement of the fork but still depends upon fork progression. Indeed, it is expected that branch migration of hemicatenanes, formed by newly synthesized chains coiled together, would be uncoupled from fork movement but still dependent on the progression of the fork, as forks might represent physical barriers that would prevent branch migration of the junctions; this consideration could explain the observation that in HU-treated wt cells both forks and X molecules stall (Figures 4B and 6B). During unperturbed conditions, we failed to detect X molecules on regions positioned far from the replication origins (Figure 1). At least two possible explanations could account for this result: if junctions are formed continuously behind replicating forks, their formation might be counteracted (or resolution increased) as forks move far from the origin. Alternatively, if junctions are exclusively formed at the origins and chase the forks, their migration out of the origin region might be somehow limited or not coordinated with the movement of the elongating forks; in the latter case, X molecules might scatter asynchronously throughout the replicon while forks diverge (Supplemental Figure S3 [http:// www.molecule.org/cgi/content/full/12/6/1499/DC1]), and, consequently, their visualization on the regions located far from the origin might be progressively prevented. The finding that, during replication of a damaged template, the branched molecules can be detected on

Sister Chromatid Junctions at Replication Origins 1507

Figure 6. A Model Representing Hypothetical Hemicatenane Transitions under Physiological or Pathological Situations (A) A single-stranded junction between replicated fragments (hemicatenane) might form early after origin activation, and undergo local isomerization to form a pseudo double Holliday junction (by pairing of newly synthesized strands), or branch migration, eventually coiling together nonhomologous sequences. (B) Joint molecules degenerate in HU-treated rad53 cells. When junctions between newly replicated strands migrate and reach stalled forks deprived of DNA polymerases, a fraction of them might be resolved into single-stranded gaps, while others could form reversed forks (see Discussion for further details).

a broader region of the replicon (Figure 4A), supports this last hypothesis: branched molecules might indeed migrate from the origin and invade adjacent regions, although delayed compared to the movement of the forks, and the damaged template could simply counteract the dispersions of the joint molecules that would be therefore clustered and detectable also in regions B and C (Figure 4A and Supplemental Figure S3). We found that in HU-treated rad53 cells the X molecules are particularly unstable. HU-treated rad53 cells fail to maintain DNA polymerases stably associated with stalled forks (Lucca et al., 2003) that progressively degenerate leading to the formation of reversed forks and partially replicated intermediates (gapped molecules) (Sogo et al., 2002). Again, the observation that X molecules are able to form in HU-treated rad53 cells but then rapidly disappear could be explained by the hemicatenane model based on the following considerations: (1) hemicatenanes reaching a stalled fork deprived of polymerases could engage the ends of newly synthesized strands in pairing (Figure 6B), leading to the formation of reversed forks that would be further stabilized by nucleosome formation (Sogo et al., 2002); this is also consistent with the observation that the disappearance of the X spike is concomitant with the appearance of the cone signal and that the formation of both type of structures does not require Rad51 and Rad52 recombination proteins (Lopes et al., 2001). In this view, hemicatenane formation would directly influence the accumulation of reversed forks. Conversely, the presence of stable stalled replisome-fork complexes, in HU-treated wt cells, would prevent the transition from hemicatenanes into reversed forks (Figure 6B), thus explaining the observation that, in HU-treated wt cells, X molecules remain stable (Figure 4B) and no reversed forks can be detected (Sogo et al., 2002). Hence, the hemicatenanes would be a potential source of dangerous recombination events in the absence of a functional replisome or in

checkpoint, defective cells experiencing replication stress. (2) A fraction of hemicatenanes could also be resolved into linear molecules by migration across the gapped forks likely resulting from lagging strand defects and unscheduled polymerase dissociation (Lopes et al., 2001; Sogo et al., 2002; Lucca et al., 2003). Again, this would not happen in wt cells that exhibit normal stalled forks (Sogo et al., 2002) and stable replisome-fork complexes (Lucca et al., 2003). Altogether, our data suggest that the newly synthesized chains would have to be coiled together in forming the hemicatenane structure (Figure 6), although more work will be required to firmly establish the nature of these molecules. The Rad53 contribution in the stabilization of X molecules seems to be specific for HU-induced replication blocks. However, it should be pointed out that, in the presence of MMS, the majority of replication forks do not experience a real block rather they are simply delayed and only a fraction of them collapses (Tercero and Diffley, 2001). Consequently, very few hemicatenanes would resolve into linear molecules and reversed forks. The X molecules form sometimes during origin firing or early step of DNA synthesis and their relative amount, compared to the one of the other initiation molecules, strongly suggests that they represent a novel initiation intermediate rather than a rare event; this is supported by the finding that X structures arise even when unscheduled firing of dormant origins occurs (Figure 5A). The mechanism leading to formation of these molecules at the origin still remains elusive, but we can exclude that, at least in S. cerevisiae, this process is mediated by homologous recombination. Although other possibilities could be envisaged, we speculate that formation of these joint structures at the origin could occur by coiling around each other the newly synthesized chains during initiation of DNA synthesis, perhaps by promoting a transient annealing of newly synthesized strands at the fork,

Molecular Cell 1508

leading to the formation of short double-stranded reversed forks coiled in the double helix conformation: while reverting to re-anneal to the parental strands, the newly synthesized filaments might remain partially coiled together, thus generating hemicatenanes. In this view, reversed forks and hemicatenanes would represent two conformational states of the same molecular transition, as in HU-treated rad53 cells, where reversed forks would result from hemicatenane run off (Figure 6B). If that is the case, it is very unlikely that these transitions occur spontaneously; rather, they should be mediated by a network of origin proteins and, in theory, it should be possible to uncouple origin firing from hemicatenane formation in certain genetic backgrounds. It will be relevant to address which are the mechanisms and the factors promoting the branch migration of the X molecules in a chromatin context and whether this process is regulated in response to DNA damage. It is expected that mutant backgrounds defective in such a process would accumulate X molecules (or their derivatives) and possibly exhibit DNA damage sensitivity. A remarkable peculiarity of these structures is their relative stability during chromosome replication and following S-phase arrest (at least in wt cells). This suggests that their resolution has to occur outside of S phase, likely prior to anaphase, to allow disentangling of sister chromatids before chromosome segregation. Hemicatenanes are potential substrates for type I topoisomerases; we therefore tested whether the X molecules’ formation and/or accumulation was affected in top1 and top3 mutant backgrounds. We found that the relative amount of X molecules in top1 and top3 mutants was comparable to the one of wt cells (data not shown). We conclude that the formation and resolution of these structures during normal S phase does not require functional Top1 or Top3. However, considering the functional redundancy of topoisomerases we cannot exclude the possibility that other topoisomerases could mediate the function of Top1 or Top3. It is possible that these structures are not a substrate for topoisomerases, at least during unperturbed S phase, perhaps because they are actively masked or because topoisomerases are prevented to act on the X structures during replication. Hemicatenane structures have been described in fully replicated SV40 minichromosomes (Sogo et al., 1986; Laurie et al., 1998) and postulated on yeast circular minichromosomes (Wellinger et al., 2003), although, in both cases, their formation has been connected with the molecular events related to replication termination. Further, hemicatenanes have been suggested to form within replication bubbles on partially replicated plasmids in E. coli (Olavarrieta et al., 2002b). Origin-dependent formation of recombination-independent joint intermediates, able to branch migrate behind the forks, might have important implications that could reflect their physiological functions under normal growing conditions. By generating DNA-mediated joint structures during initiation of DNA synthesis, cells would couple sister chromatid tethering with replicon firing. Such a mechanism could very well contribute to sister chromatid cohesion and provide an explanation on how sister chromatids are tethered in order to be efficiently and precisely packed together into the cohesin-dependent proteinaceous loop (Haering et al., 2002). This

would also support previous analysis suggesting that sister chromatids are topologically interlocked until anaphase (Murray and Szostak, 1985, and references therein). A corollary of this hypothesis is that the efficiency of origin activation, and therefore the number of initiation events, should influence the establishment of physical links between sister chromatids; thus any genetic situation altering the replicon/chromosome ratio might, in turn, affect sister chromatid cohesion. Following genotoxic insults, this mechanism could allow the cell to search for homology between sister chromatids and even prevent loss of chromosomal regions as a result of double-strand break formation. Recent observations indicate that when replication forks encounter a damaged template, the strand that hits the lesion transiently stalls while the other one proceeds further (Page`s and Fuchs, 2003). Hemicatenanes could play an important role within this context: by reaching the stalled strand, hemicatenanes could actively promote template switching by displacing the stalled strand from the template and allowing its pairing with the other newly synthesized chain; in this way, the stalled strand could copy a correct template. Hence, hemicatenane formation at the origin might allow the cell to program an error-free replication bypass process (i.e., template switching, Higgins et al., 1976) when the template is damaged. It will be a challenge for the future to identify the genetic pathways required for the formation, migration, and resolution of these origin-dependent branched molecules. Experimental Procedures S. cerevisiae Strains and Growing Conditions The strains used in this study are isogenic derivatives of W303-1A (Thomas and Rothstein, 1989). rad51 (CY2269), rad52 (CY2272), top1 (CY2278), and top3 (CY2284) KanMX4-deletion strains were constructed using the PCR-based strategy already described (Wach et al., 1994); rad53-K227A (CY2034) mutant has been described in Pellicioli et al., 1999. Strains were grown in YPD, presynchronized in G1 by adding 2 ␮g/ml ␣ factor, and released from the G1 arrest by centrifugation and resuspension in fresh YPD medium. DNA Extraction Procedures Total genomic DNA was isolated mainly according to Allers and Lichten (2000), with modifications (CTAB extraction). 200 ml cultures (2–4 ⫻ 109 cells) were arrested by addition of 0.1% Sodium Azide (final concentration) and cooled down in ice. Cells were harvested by centrifugation, washed in cold water, and incubated in spheroplasting buffer (1 M sorbitol, 100 mM EDTA [pH 8.0], 0.1% ␤-mercaptoethanol, and 100 U zymoliase/ml) for 45 min at 30⬚C. 2 ml water, 200 ␮l RNase A (10 mg/ml), and 2.5 ml Solution I (2% w/v cetyl-trimethyl-ammonium-bromide [CTAB], 1.4 M NaCl, 100 mM Tris HCl [pH 7.6], and 25 mM EDTA [pH 8.0]) were sequentially added to the spheroplast pellets and samples were incubated 30 min at 50⬚C. 200 ␮l Proteinase K (20 mg/ml) were then added and the incubation was prolonged at 50⬚C for 1 hr 30 min, and at 30⬚C overnight. The sample was then centrifuged at 6000 g for 10 min: the cellular debris pellet was kept for further extraction, while the supernatant was extracted with 2.5 ml chloroform/isoamylalcohol (24/1) and the DNA in the upper phase was precipitated by addition of 2 volumes Solution II (1% w/v CTAB, 50 mM Tris-HCl [pH 7.6], and 10 mM EDTA) and centrifugation at 12000 ⫻ g for 10 min. The pellet was resuspended in 2 ml Solution III (1.4 M NaCl, 10 mM Tris HCl [pH 7.6], and 1 mM EDTA). Residual DNA in the cellular debris pellet was also extracted by resuspension in 2 ml Solution III and incubation at 50⬚C for 30 min, followed by extraction in 1 ml chloro-

Sister Chromatid Junctions at Replication Origins 1509

form/isoamylalcohol (24/1). The upper phase was pooled together with the main DNA prep. Total DNA was then precipitated with 1 volume isopropanol, washed with 70% ethanol, air dried, and finally resuspended in TE 1X. In Figure 2A, two different DNA extraction procedures were used in addition to the one described above (Method 1): in one case (Method 2), total genomic DNA was extracted according to the protocol of the “QIAGEN genomic DNA Handbook,” using genomic-tip 100/G columns; in the last case (Method 3), nuclei were isolated by addition of glass beads to intact harvested cells, 15–20 cycles of vortexing in the presence of Nuclei Isolation Buffer at 4⬚C (17% glycerol, 50 mM MOPS, 150 mM potassium acetate, 2 mM MgCl2, 500 ␮M spermidine, and 150 ␮M spermine [pH 7.2]) and centrifugation of the extract at 8000 ⫻ g for 10 min at 4⬚C. Nuclei pellets were then lysed in QIAGEN G2 buffer, in the presence of RNase A and Proteinase K, and from this step on the “QIAGEN genomic DNA Handbook” protocol was followed, using QIAGEN genomic-tip 100/G columns to isolate DNA. Two-Dimensional Electrophoresis and Hybridization 2D gel electrophoresis was carried out as originally described by Brewer and Fangman (1987). DNA was blotted onto Nylon Gene Screen Plus membrane (NEN). Membranes were initially probed with the BamHI-NcoI 3.0 kb fragment, spanning ARS305, gel purified from plasmid A6C-110 (kindly provided by C. Newlon, UMDNJ, Newark, NJ). If necessary, membranes were stripped and sequentially hybridized for the detection of fragments C, A, B, and D. The corresponding probes were amplified from yeast genomic DNA by PCR and gel purified. Oligonucleotides sequences are available upon request. Quantification of Autoradiograms All signals were quantified using a PhosphorImager Molecular Dynamics Storm 820 and ImageQuant as analysis program. Each signal was normalized in respect to the total amount of signals present in that gel, including linear monomers. “Object Average” mode of background correction was used. To help readability, all values were then reported using arbitrary units, but keeping the scale among different time points and different fragments in the same experiment. In Figure 2A, the internal proportion (%) of different replication intermediates is presented.

DNA that restrains branch migration of Holliday junctions. Nucleic Acids Res. 28, e6. Bell, S.P., and Dutta, A. (2002). DNA replication in eukaryotic cells. Annu. Rev. Biochem. 71, 333–374. Be´nard, M., Maric, C., and Pierron, G. (2001). DNA replicationdependent formation of joint DNA molecules in Physarum polycephalum. Mol. Cell 7, 971–980. Bianchi, M., DasGupta, C., and Radding, C.M. (1983). Synapsis and the formation of paranemic joints by E. coli RecA protein. Cell 34, 931–939. Borde, V., Goldman, A.S., and Lichten, M. (2000). Direct coupling between meiotic DNA replication and recombination initiation. Science 290, 806–809. Brewer, B.J., and Fangman, W.L. (1987). The localization of replication origins on ARS plasmids in S. cerevisiae. Cell 51, 463–473. Brewer, B.J., Sena, E.P., and Fangman, W.L. (1988). Analysis of replication intermediates by two-dimensional agarose gel electrophoresis. In Cancer Cells, 6: Eukaryotic DNA Replication (Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press), pp. 229–234. Cimprich, K.A. (2003). Fragile sites: breaking up over a slowdown. Curr. Biol. 13, R231–R233. Collins, I., and Newlon, C.S. (1994). Meiosis-specific formation of joint molecules containing sequences from homologous chromosomes. Cell 76, 65–75. Delidakis, C., and Kafatos, F.C. (1989). Amplification enhancers and replication origins in the autosomal chorion gene cluster of Drosophila. EMBO J. 8, 891–901. Deshpande, A.M., and Newlon, C.S. (1996). DNA replication fork pause sites dependent on transcription. Science 272, 1030–1033. Dijkwel, P.A., Vaughn, J.P., and Hamlin, J.L. (1991). Mapping of replication initiation sites in mammalian genomes by two-dimensional gel analysis: stabilization and enrichment of replication intermediates by isolation of the nuclear matrix. Mol. Cell. Biol. 11, 3850– 3859. Duckett, D.R., Murchie, A.I., Diekmann, S., von Kitzing, E., Kemper, B., and Lilley, D.M. (1988). The structure of the Holliday junction, and its resolution. Cell 55, 79–89. Duckett, D.R., Murchie, A.I., and Lilley, D.M. (1990). The role of metal ions in the conformation of the four-way DNA junction. EMBO J. 9, 583–590.

In Vitro Resolution of X-Shaped Molecules Following first dimension gel electrophoresis, the slices of agarose were incubated for branch migration either in TNE (without Mg2⫹) (10 mM Tris-Hcl [pH 8.0], 100 mM NaCl, and 0.1 mM EDTA), or in TNM (with Mg2⫹) (10 mM Tris-Hcl [pH 8.0], 50 mM NaCl, 10 mM MgCl2, and 0.1 mM EDTA) Branch Migration Buffer as described in Panyutin and Hsieh, 1994, for 1 to 4 hr at 65⬚C. Agarose lanes were then processed for second-dimension electrophoresis.

Gerbi, S.A., and Bielinsky, A.K. (2002). DNA replication and chromatin. Curr. Opin. Genet. Dev. 12, 243–248.

Acknowledgments

Haering, H.C., Lowe, J., Hochwagen, A., and Nasmyth, K. (2002). Molecular architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773–788.

We thank F. Antequera, M. Bianchi, N. Cozzarelli, J. Haber, I. Hickson, C. Lucca, G. Maffioletti, K. Nasmyth, C. Newlon, A. Pellicioli, R. Rothstein, J. Sogo, R. Wellinger, and all members of our laboratory for helpful discussions and sharing results before publication, I. Hickson and R. Rothstein for reagents, and D. Facchinetti and Y. Docksani for technical support. This work was supported by Associazione Italiana per la Ricerca sul Cancro, and partially by European Union (grant RTN2-2001-00177), Telethon (grant GGP030412), and Ministero della Salute. Received: July 16, 2003 Revised: September 5, 2003 Accepted: October 8, 2003 Published: December 18, 2003 References Aguilera, A. (2002). The connection between transcription and genomic instability. EMBO J. 21, 195–201. Allers, T., and Lichten, M. (2000). A method for preparing genomic

Gonzales-Barrera, S., Cortes-Ledesma, F., Wellinger, R., and Aguilera, A. (2003). Equal sister chromatid exchange is a major mechanism of double strand break repair in yeast. Mol. Cell 11, 1661–1671.

Heck, M.S., and Spradling, A.C. (1990). Multiple replication origins are used during Drosophila chorion gene amplification. J. Cell Biol. 110, 903–914. Hickson, I.D. (2003). RecQ helicases: caretakers of the genome. Nat. Rev. Cancer 3, 169–178. Higgins, N.P., Kato, K., and Strauss, B. (1976). A model for replication repair in mammalian cells. J. Mol. Biol. 101, 417–425. Ivessa, A.S., Zhou, J.Q., and Zakian, V.A. (2000). The Saccharomyces Pif1p DNA helicase and the highly related Rrm3p have opposite effects on replication fork progression in ribosomal DNA. Cell 100, 479–489. Kadyk, L.C., and Hartwell, L.W. (1992). Sister chromatids are preferred over homologs as substrates for recombinational repair in Saccharomyces cerevisiae. Genetics 132, 387–402. Kolodner, R.D., Putnam, C.D., and Myung, K. (2002). Maintenance of genome stability in Saccharomyces cerevisiae. Science 297, 552–557. Kornberg, A., and Baker, J. (1992). In DNA Replication, 2nd ed., (New York: W.H. Freeman & Co.), p. 280.

Molecular Cell 1510

Koshland, D., and Hartwell, L.H. (1987). The structure of sister minichromosome DNA before anaphase in Saccharomyces cerevisiae. Science 238, 1713–1716. Laurie, B., Katrich, V., Sogo, J.M., Koller, T., Dubochet, J., and Stasiak, A. (1998). Geometry and physics of catenates applied to the study of DNA replication. Biophys. J. 74, 2815–2822. Liang, C., Spitzer, J.D., Smith, H.S., and Gerbi, S.A. (1993). Replication initiates at a confined region during DNA amplification in Sciara DNA puff II/9A. Genes Dev. 7, 1072–1084. Lilley, D.M. (1997). All change at Holliday junction. Proc. Natl. Acad. Sci. USA 94, 9513–9515. Lopes, M., Cotta-Ramusino, C., Pellicioli, A., Liberi, G., Plevani, P., Muzi-Falconi, M., Newlon, C.S., and Foiani, M. (2001). The DNA replication checkpoint response stabilizes stalled replication forks. Nature 412, 557–561. Lucas, I., and Hyrien, O. (2000). Hemicatenanes form upon inhibition of DNA replication. Nucleic Acids Res. 28, 2187–2193. Lucca, C., Vanoli, F., Cotta-Ramusino, C., Pellicioli, A., Liberi, G., Haber, J., and Foiani, M. (2003). Checkpoint-mediated control of replisome-fork association and signalling in response to replication pausing. Oncogene, in press. Malkova, A., Ivanov, E.L., and Haber, J.E. (1996). Double-strand break repair in the absence of RAD51 in yeast: a possible role for break-induced DNA replication. Proc. Natl. Acad. Sci. USA 93, 7131– 7136. Murray, A.W., and Szostak, J.W. (1985). Chromosome segregation in mitosis and meiosis. Annu. Rev. Cell Biol. 1, 289–315. Nasmyth, K. (2001). Disseminating the genome: joining, resolving and separating sister chromatids during mitosis and meiosis. Annu. Rev. Genet. 35, 673–745. Newlon, C.S., Collins, I., Dershowitz, A., Deshpande, A.M., Greenfeder, S.A., Ong, L.Y., and Theis, J.F. (1993). Analysis of replication origin function on chromosome III of Saccharomyces cerevisiae. Cold Spring Harb. Symp. Quant. Biol. 58, 415–423. Olavarrieta, L., Martinez-Robles, M.L., Sogo, J.M., Stasiak, A., Hernandez, P., Frimer, D.B., and Schvartzman, J.B. (2002a). Supercoiling, knotting and replication fork reversal in partially replicated plasmids. Nucleic Acids. Res. 30, 656–666. Olavarrieta, L., Martinez-Robles, M.L., Herna`ndez, P., Krimer, D.B., and Schvartzman, J.B. (2002b). Knotting dynamics during DNA replication. Mol. Microbiol. 46, 699–707. Osborn, A.J., Elledge, S.J., and Zou, L. (2002). Checking on the fork: the DNA-replication stress-response pathway. Trends Cell Biol. 12, 509–516. Page`s, V., and Fuchs, R.P. (2003). Uncoupling of leading- and lagging-strand DNA replication during lesion bypass in vivo. Science 300, 1300–1303. Panyutin, I.G., and Hsieh, P. (1994). The kinetics of spontaneous DNA branch migration. Proc. Natl. Acad. Sci. USA 91, 2021–2025. Paques, F., and Haber, J.E. (1999). Multiple pathways of recombination induced by double-strand breaks in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 63, 349–404. Paulovich, A.G., and Hartwell, L.H. (1995). A checkpoint regulates the rate of progression through S-phase in S. cerevisiae in response to DNA damage. Cell 82, 841–847. Pellicioli, A., Lucca, C., Liberi, G., Marini, F., Lopes, M., Plevani, P., Romano, A., Di Fiore, P., and Foiani, M. (1999). Activation of Rad53 kinase in response to DNA damage and its effect in modulating phosphorylation of the lagging strand DNA polymerase. EMBO J. 18, 6561–6572. Schwacha, A., and Kleckner, N. (1994). Identification of joint molecules that form frequently between homologs but rarely between sister chromatids during yeast meiosis. Cell 76, 51–63. Schwacha, A., and Kleckner, N. (1995). Identification of double Holliday junctions as intermediates in meiotic recombination. Cell 83, 783–791. Segurado, M., Gomez, M., and Antequera, F. (2002). Increased recombination intermediates and homologous integration hot spots at DNA replication origins. Mol. Cell 10, 907–916.

Shirahige, K., Hori, Y., Shiraishi, K., Yamashita, M., Takahashi, K., Obuse, C., Tsurimoto, T., and Yoshikawa, H. (1998). Regulation of DNA-replication origins during cell-cycle progression. Nature 395, 618–621. Sogo, J.M., Stahl, H., Koller, T., and Knippers, R. (1986). Structure of replicating Simian Virus 40 minichromosomes. The replication fork, core histone segregation and terminal structures. J. Mol. Biol. 189, 189–204. Sogo, J.M., Lopes, M., and Foiani, M. (2002). Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defect. Science 297, 599–602. Tercero, J.S., and Diffley, J.F.X. (2001). Regulation of DNA replication fork progression through damaged DNA by the Mec1/Rad53 checkpoint. Nature 412, 553–557. Thomas, B.J., and Rothstein, R. (1989). Elevated recombination rates in trancriptionally active DNA. Cell 56, 619–629. Wach, A., Brachat, A., Pohlmann, R., and Philippsen, P. (1994). New heterologous modules for classical or PCR-based gene disruption in Saccharomyces cerevisiae. Yeast 10, 1793–1808. Wellinger, R.E., Scha¨r, P., and Sogo, J.M. (2003). Rad52 independent accumulation of DNA joint circular minichromosomes during S-phase in Saccharomyces cerevisiae. Mol. Cell. Biol. 23, 6363– 6372. Zou, H., and Rothstein, R. (1997). Holliday junctions accumulate in replication mutants via a RecA homolog-independent mechanism. Cell 90, 87–96. Zou, L., and Elledge, S.J. (2003). Sensing DNA damage through ATRIP recognition of RPA-ssDNA Complexes. Science 300, 1542– 1548.