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Molecular and Cellular Biology, June 2007, p. 4283-4292, Vol. 27, No. 12
0270-7306/07/$08.00+0     doi:10.1128/MCB.02196-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

DNA Structure-Induced Recruitment and Activation of the Fanconi Anemia Pathway Protein FANCD2{triangledown}

A. Sobeck, S. Stone, and M. E. Hoatlin*

Oregon Health and Science University, Portland, Oregon 97239

Received 23 November 2006/ Returned for modification 15 December 2006/ Accepted 28 March 2007


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ABSTRACT
 
The Fanconi anemia (FA) pathway proteins are thought to be involved in the repair of irregular DNA structures including those encountered by the moving replication fork. However, the nature of the DNA structures that recruit and activate the FA proteins is not known. Because FA proteins function within an extended network of proteins, some of which are still unknown, we recently established cell-free assays in Xenopus laevis egg extracts to deconstruct the FA pathway in a fully replication-competent context. Here we show that the central FA pathway protein, xFANCD2, is monoubiquitinated (xFANCD2-L) rapidly in the presence of linear and branched double-stranded DNA (dsDNA) structures but not single-stranded or Y-shaped DNA. xFANCD2-L associates with dsDNA structures in an FA core complex-dependent manner but independently of xATRIP, the regulatory subunit of xATR. Formation of xFANCD2-L is also triggered in response to circular dsDNA, suggesting that dsDNA ends are not required to trigger monoubiquitination of FANCD2. The induction of xFANCD2-L in response to circular dsDNA is replication and checkpoint independent. Our results provide new evidence that the FA pathway discriminates among DNA structures and demonstrate that triggering the FA pathway can be uncoupled from DNA replication and ATRIP-dependent activation.


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INTRODUCTION
 
Fanconi anemia (FA) belongs to the group of chromosomal instability syndromes including ataxia telangiectasia, Bloom syndrome, and hereditary breast cancer. FA cells are hypersensitive towards DNA interstrand cross-link (ICL)-inducing agents and have highly elevated spontaneous chromosomal breakage rates, suggesting a role for FA proteins in the DNA damage response. The FA pathway consists of an upstream FA "core complex" containing at least eight proteins (FANCA, -B, -C, -E, -F, -G, -L, and -M) that is required for the DNA damage-induced activation (monoubiquitination) of its downstream target, FANCD2 (9, 12, 13, 18, 27, 37). Accumulating evidence supports a role for the FA pathway in DNA double-strand break (DSB) repair via homologous recombination (HR). Indeed, FA cells are defective in the repair of ICL-induced DNA DSBs mediated by HR (44). Activated FANCD2 interacts with major components of HR repair, such as BRCA1 (breast cancer-associated protein 1), Rad51, and the downstream FA protein BRCA2/FANCD1 (8, 12, 17, 22, 47, 49); moreover, the BRCA2 interactor PALB2 was recently identified as a Fanconi protein, FANCN (42, 43, 50). In addition, involvement of the FA proteins in DNA DSB repair via single-strand annealing and nonhomologous end joining has been suggested (10, 11, 26, 36). The recently identified FANCM and FANCJ proteins indicate a direct involvement of FA proteins at sites of DNA repair: FANCM is an FA core complex member (30, 31, 34) homologous to the archaeal protein Hef (helicase-associated endonuclease for fork-structured DNA), which resolves stalled replication forks (20). FANCJ/BRIP1, which is thought to play a role downstream in the FA pathway (23, 24, 39), is a 5'-to-3' DNA helicase with substrate specificity towards splayed DNA duplexes (Y-shaped DNA) (14). In support of a role for FA proteins in the processing of DNA structures, a recent study demonstrated that recombinant FANCD2 has direct DNA binding activity in vitro (38). Moreover, Ciccia et al. reported that recombinant FANCM is directed to branched DNA structures by a novel FA core complex member, FAAP24 (4). Accumulating evidence suggests that the FA proteins act specifically during the replication stage of the cell cycle. Rothfuss and Grompe showed that ICL-induced formation of DNA DSBs is a replication-associated event and that repair of such breaks is defective in FA cells (44). Our laboratory recently established cell-free assays in Xenopus laevis egg extracts to show that FA proteins are recruited to chromatin in a replication-dependent, DNA damage-stimulated manner, where they function to prevent accumulation of chromosomal breaks during each round of replicative DNA synthesis (46). Thus, the current model suggests that FA proteins are recruited to DNA lesions generated from replication fork stalling at sites of damaged DNA. However, which types of DNA lesions trigger the recruitment and activation of the FA proteins has not yet been investigated, in part because the FA proteins are functionally interdependent and not all the proteins in the FA pathway have been identified. We developed a new strategy that allows us to bypass the requirement for ongoing replication to activate the FA pathway by mimicking irregular DNA structures and specific replication intermediates in cell-free assays. Using this strategy, we investigated which types of DNA structures induce monoubiquitination and recruitment of FANCD2 in the presence of a fully functional FA pathway.


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MATERIALS AND METHODS
 
Production of xFA antibodies. Generation of antibodies to xFANCD2 and xFANCA was described previously (46). Antibodies to xMre11 were a kind gift of J. Gautier, and antibodies to xATRIP were a kind gift from K. Cimprich.

Preparation of Xenopus egg extracts. Extracts were prepared from Xenopus eggs according to the method of Murray (35). Tautomycin (3 µM), caffeine (4 mM), and recombinant Xenopus geminin (5 ng/µl) (7) were added as indicated.

DNA replication assay. Replication of sperm chromatin in S-phase egg extracts was monitored as described previously (46).

Preparation of DNA substrates. (i) Plasmid-derived DNA substrates. Circular plasmid DNA (pBSKS) was prepared from Escherichia coli cultures using a QiaFilter Plasmid Maxi kit (QIAGEN). Plasmid DNA was linearized with NotI (Fermentas) or fragmented (14 DNA fragments) with HaeIII (Roche) restriction enzyme. Plasmid DNA was nicked using DNase I (Invitrogen). Linearized, nicked, and intact circular (supercoiled) plasmid forms were distinguished by agarose gel electrophoresis (see Fig. 4D, inset). M13mp18 single-stranded plasmid DNA was obtained from Bayou Biolabs.


Figure 4
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  		FIG. 4. dsDNA ends are not a major trigger for xFANC2-L formation. (A) Induction of xFANCD2-L occurs independently of dsDNA ends. Egg extracts were incubated for 30 min with 45 ng/µl of circular plasmid dsDNA (lane 1), fragmented plasmid dsDNA (lane 2), or linearized plasmid dsDNA (lane 3). DNA-free extract served as a negative control (lane 4). Following incubation, 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting. (B) Linearized plasmid dsDNA induces xFANCD2-L earlier and at lower DNA concentrations than fragmented plasmid dsDNA. Egg extracts were incubated with three different concentrations of linearized or fragmented plasmid dsDNA (50 ng/µl, lanes 2, 5, and 8; 25 ng/µl, lanes 3, 6, and 9; 10 ng/µl, lanes 4 and 7). Aliquots were taken at 5 min, 15 min, and 25 min, and 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting. One microliter of DNA-free extract was used as a negative control (lane 1). (C) High concentrations of circular plasmid dsDNA induce both xFANCD2-L and xMre11-PPP. Egg extracts were incubated for 30 min with increasing concentrations of circular plasmid dsDNA as indicated. Following incubation, 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting. DNA-free extract incubated for 30 min served as a negative control (lane 5). (D) Low concentrations of nicked plasmid DNA induce xFANCD2-L and xMre11-PPP. Egg extracts were incubated for 30 min with 30 ng/µl (lanes 2 to 4) or 60 ng/µl (lanes 5 to 7) of linearized plasmid dsDNA (lanes 2 and 5), nicked plasmid dsDNA (lanes 3 and 6), or circular plasmid dsDNA (lanes 4 and 7). DNA-free extract served as a negative control (lane 1). Following incubation, 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting. (Inset) For quality control of circular plasmid dsDNA (lanes 2 and 3), linear plasmid dsDNA (lanes 4 and 5), and nicked plasmid dsDNA (lanes 6 and 7), DNA samples (2.5 µg/lane) were analyzed by agarose gel electrophoresis. M, dsDNA size marker. (E) xFANCD2-L is induced by circular ssDNA. Egg extracts were incubated for 30 min with 160 ng/µl of circular plasmid dsDNA (lane 2) or circular plasmid ssDNA (lane 3). DNA-free extract served as a negative control (lane 1). Following incubation, 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting.

(ii) Oligonucleotide-derived substrates. The oligonucleotide 5'-(dG)40-(dA)30-3' (ssDNA70) was annealed to oligonucleotide 5'-(dT)30-(dC)40-3' to generate dsDNA70. Y-DNA70 and forkDNA70 were generated as described previously (25). Oligonucleotides 5'-GGTTGACGTTCTAGTGTGACCGCAATACGGATAAGGGCTGAGCACGCCGACGAACATAC-3' (Holliday junction [HJ] strand 1), 5'-GTATGTTCGTCGGCGTGCTCAGCCCTTATCCCAGAATGCACCAACAGTTCCTCAAGATAGAGACTC-3' (HJ strand 2), 5'-GAGTCTCTATCTTGAGGAACTGTTGGTGCATTCTGGGATAAGGGCACTATGGCTCCACTGATGTCGTAAGCATCC-3' (HJ strand 3), and 5'-GGATGCTTACGACATCAGTGGAGCCATAGTGCCCTTATCCGTATTGCGGTCACACTAGAACGTCAACC-3' (HJ strand 4) were annealed to form HJ-DNA68 (134 bp) as previously described (45). HJ strand 4 was annealed to HJ strand 5, 5'-GGTTGACGTTCTAGTGTGACCGCAATACGGATAAGGGCACTATGGCTCCACTGATGTCGTAAGCATCC-3', to generate a linear double-stranded DNA (dsDNA) fragment (dsDNA68 [68 bp]) with sequence similarity to the HJ. HJ-DNA68 and dsDNA68 structures were labeled with [{gamma}-32P]ATP at the 5' end of HJ strands 3 and 5, respectively, and purified by electrophoresis through a nondenaturing 4% polyacrylamide gel. DNA structures were excised from the gel and eluted into 40 µl of elution buffer using the QIAEX gel extraction kit (QIAGEN).

(iii) Bead-DNA. Streptavidin-conjugated magnetic beads (Dynal, Inc.) coupled to biotinylated DNA structures were incubated for 35 min in egg extracts. The biotin group was attached to the 3' end of 5'-(dG)40-(dA)30-3' (in ssDNA70, dsDNA70, Y-DNA70, and forkDNA70), the 3' end of HJ strand 3 (HJ-DNA68), or the 3' end of HJ strand 5 (dsDNA68). Following incubation, DNA-coupled beads were washed five times with XB buffer (46) by collection with a magnetic rack.

Immunodepletion. Immunodepletions were performed essentially as described previously (46). In brief, 200 µl of Sepharose 4B beads (50% slurry; Amersham) was rotated overnight at 4°C with 500 µl of phosphate-buffered saline and 100 µl of anti-xFANCA, -xFANCD2, -xMre11, or -xATRIP affinity-purified antisera or the corresponding preimmune sera. The beads were pelleted from solution and washed three times in XB buffer. For depletion, 100 µl of extract was added to the beads and the extract-bead mix was rotated for two rounds at 4°C for 40 min.

Immunoblotting. Protein samples were separated on gradient gels (Invitrogen) and transferred to Immobilon P membranes (Millipore). After being blocked in 5% milk for 1 h, membranes were incubated with the following primary antibodies: anti-XFANCD2 (1:1,000), xFANCA (1:1,000), xATRIP (1:1,000), and xMRE11 (1:5,000). Horseradish peroxidase-conjugated rabbit secondary antibody (Jackson Labs) was used (1:10,000). Protein bands were visualized using an ECL Plus system (Amersham).

Ubiquitination assay. The ubiquitination assay was conducted as previously described (51) using nickel-nitrilotriacetic acid agarose beads (QIAGEN).


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RESULTS
 
DNA substrates stimulate monoubiquitination of xFANCD2. Although little is known about how the FA proteins are activated and recruited to DNA, the mobility shift of FANCD2 (FANCD2-L) during S phase and following DNA damage is widely used as a measure for FA pathway activation. To test whether FANCD2-L can be directly induced in Xenopus egg extracts in response to DNA substrates, we generated dsDNA fragments by a restriction digest of a dsDNA plasmid. A similar strategy was used by Costanzo et al., who demonstrated that the ends of these dsDNA fragments mimic DNA DSBs that activate the DNA DSB repair enzyme xMre11 by hyperphosphorylation in egg extracts (6). Addition of the dsDNA fragments to egg extracts resulted in the appearance of a lower-mobility form of xFANCD2 (xFANCD2-L) (Fig. 1A, middle). As expected, the dsDNA fragments also induced hyperphosphorylation of xMre11 (xMre11-PPP, Fig. 1A, top). In contrast, the FA core complex protein xFANCA did not undergo a detectable size change in the DNA-containing extract (Fig. 1A, bottom). Taken together, dsDNA fragments stimulated the activation of xFANCD2 and Mre11.


Figure 1
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  		FIG. 1. Monoubiquitination of xFANCD2 in response to dsDNA structures. (A) Presence of dsDNA fragments triggers formation of xFANCD2-L. dsDNA fragments were incubated in egg extracts for 30 min. One microliter of egg extract containing no DNA (circle with diagonal line) or 150 ng/µl dsDNA fragments was analyzed for xMre11, xFANCD2, and xFANCA by immunoblotting. Mobility shift of xMre11 was used as a positive control (6). Egg extracts contained the phosphatase inhibitor tautomycin to stabilize phosphorylated isoforms of xMre11. (B) xFANCD2-L induced by dsDNA fragments represents monoubiquitinated xFANCD2. DNA-free extracts (lanes 1, 3, 5, and 7) or extracts containing 160 ng/µl dsDNA fragments (lanes 2, 4, 6, and 8) were supplemented with either His-tagged human ubiquitin (His-Ub) (lanes 3, 4, 7, and 8) or an equal volume of H2O (lanes 1, 2, 5, and 6). Following incubation for 30 min, 1 µl of extract was directly analyzed for xFANCD2-L (lanes 1 to 4). His-tagged, ubiquitinated proteins were reisolated from DNA-containing or DNA-free extracts using Ni beads and analyzed for the presence of xFANCD2 by immunoblotting (lanes 7 and 8). Ni beads reisolated from DNA-free or DNA-containing extracts that did not contain His-Ub (lanes 5 and 6) were used as a negative control for direct binding of xFANCD2-S or -L to Ni beads. (C) xFANCD2-L is induced by linear and forked dsDNA structures. DNA structures ssDNA70 (lanes 2 and 3), dsDNA70 (lanes 4 and 5), Y-DNA70 (lanes 6 and 7), and forkDNA70 (lanes 8 and 9) were incubated in egg extracts for 30 min, and 1 µl of extract was analyzed for induction of xFANCD2-L or xMre11-PPP by immunoblotting. Symbols for each DNA substrate are explained in Table 1. For stabilization of xMre11-PPP, the phosphatase inhibitor tautomycin (T) was added to egg extracts where indicated (lanes 3, 5, 7, and 9).

The mobility shift of human FANCD2 following DNA damage has been shown to be due to monoubiquitination (12). To test if the dsDNA fragment-induced mobility shift of xFANCD2 in egg extracts (Fig. 1A, middle) is due to monoubiquitination, we added His-tagged ubiquitin (His-Ub) to DNA-free or dsDNA fragment-containing extracts, followed by reisolation of His-Ub-tagged proteins using nickel beads. As shown in Fig. 1B, His-Ub-xFANCD2 was highly enriched in extracts containing dsDNA fragments compared to DNA-free extracts, demonstrating that the dsDNA fragment-induced xFANCD2-L represents ubiquitinated xFANCD2.

Linear and branched dsDNA substrates trigger activation of xFANCD2-L. To further map the structural characteristics of DNA capable of triggering formation of xFANCD2-L, we compared four DNA substrates mimicking DNA structures known to arise during replication: single-stranded linear DNA (ssDNA70; length, 70 nucleotides), double-stranded linear DNA (dsDNA70; length, 70 bp), Y-shaped DNA (Y-DNA70; length, 70 bp), and forked dsDNA (forkDNA70; length, 70 bp) (25) (Table 1). These DNA structures were recently used to stimulate the interaction between topoisomerase III{alpha} and the Bloom syndrome helicase in Xenopus egg extracts (25). Following addition of the DNA substrates, extracts were analyzed for the presence of xFANCD2-L and xMre11-PPP. The dsDNA70 substrate induced both xFANCD2-L and xMre11-PPP (Fig. 1C, lanes 4 and 5), similar to what we observed in response to plasmid-derived dsDNA fragments (Fig. 1A). In addition, forkDNA70 (Fig. 1C, lanes 8 and 9), but not ssDNA70 (Fig. 1C, lanes 2 and 3) or Y-DNA70 (Fig. 1C, lanes 6 and 7), induced xFANCD2-L. Interestingly, forkDNA70 did not induce xMre11-PPP (Fig. 1C, lanes 8 and 9) despite the presence of three dsDNA ends in this structure. The FA core complex protein xFANCA did not show a detectable mobility shift in response to any of the four DNA structures tested (data not shown). Taken together, xFANCD2-L induction was triggered by linear and forked dsDNA but not ssDNA or Y-shaped DNA.


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  		TABLE 1. Overview of DNA structures used in this studya

xFANCD2-L associates with linear and branched dsDNA substrates. Following monoubiquitination, human FANCD2 is thought to associate with sites of DNA damage (12, 33, 47, 49). Thus, we hypothesized that xFANCD2-L associated with those DNA structures that triggered its formation in egg extracts. To test this, we coupled ssDNA70 and forkDNA70 to magnetic beads prior to incubation in egg extracts. After incubation, the bead-DNA structures were pelleted from the extract and the pellet and soluble fractions were analyzed separately. As shown in Fig. 2A, xFANCD2-L was induced in the presence of bead-forkDNA70 but not in the presence of bead-ssDNA70 (compare lanes 2 and 3). Reisolation of the bead-DNA structures from the extract revealed that xFANCD2-L was associated with the forkDNA70 (Fig. 2A, lane 9), whereas no free xFANCD2-L was detected in the postincubation extract (Fig. 2A, lane 6). We also observed that xFANCA was enriched on the bead-DNA structures (Fig. 2A, lanes 8 and 9), consistent with previous results suggesting close DNA association of the FA core complex proteins (32, 41). Next, we directly compared xFANCD2-L recruitment to bead-bound ssDNA70, dsDNA70, Y-DNA70, and forkDNA70. As shown in Fig. 2B, xFANCD2-L associated with dsDNA70 (lane 2) and forkDNA70 (lane 4), but not with ssDNA70 (lane 1) or Y-DNA70 (lane 3), consistent with the results described for the soluble DNA structures. Interestingly, a portion of the nonubiquitinated form of xFANCD2, xFANCD2-S, was associated with all four bead-DNA substrates (Fig. 2A, lanes 8 and 9; Fig. 2B, lanes 1 to 4). xFANCA associated with all tested bead-DNA structures but did not exhibit any detectable mobility changes (Fig. 2B, lanes 1 to 4). Together, these results show that nonmodified xFANCD2 associated with DNA substrates irrespective of their composition, whereas monoubiquitinated xFANCD2-L was found associated selectively with linear and forked dsDNA. We also tested whether xFANCD2-L formation was triggered by an HJ, a cruciform structure generated as a major intermediate during HR. We compared the cruciform structure HJ-DNA68 to a linear dsDNA68 structure of similar length and sequence (Table 1). For the comparison, we took into account the fact that the bead-bound HJ-DNA68 structure contains twice the number of base pairs and three times the number of free dsDNA ends that the bead-bound dsDNA68 structure contains. Following coupling to beads, the DNA structures were each incubated separately in 10 µl egg extract at the following concentrations: 10 pmol HJ-DNA68, 10 pmol dsDNA68, 20 pmol dsDNA68, and 30 pmol dsDNA68. This strategy allowed us to test three different molar ratios of HJ-DNA68 to dsDNA68 (Fig. 2C), 1:1 (equal numbers of DNA molecules), 1:2 (equal numbers of total DNA base pairs), and 1:3 (equal numbers of free dsDNA ends). We found that similar amounts of xFANCD2-L bound to the HJ-DNA68 and to dsDNA68 at a molar ratio of 1:1 between the two structures (Fig. 2C, compare lanes 1 and 2). In addition, xFANCD2-L recruitment did not occur proportionally to the total number of DNA base pairs present in the DNA structures (Fig. 2C, compare lanes 1 and 3) or in proportion to the number of DNA ends presented by the DNA structures (Fig. 2C, compare lanes 1 and 4). The marked increase in dsDNA68-bound xFANCD2-L between extracts containing 10 pmol dsDNA68 (lane 2) and 20 pmol dsDNA68 (lane 3) is a phenomenon that we consistently observe, indicating that above a critical DNA concentration in the egg extracts, binding of xFANCD2-L to activating DNA structures increases in a nonlinear manner. In summary, xFANCD2-L was induced by—and recruited to—an HJ but did not show a preference for this structure over a linear dsDNA structure.


Figure 2
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  		FIG. 2. xFANCD2-L associates with linear and branched dsDNA structures. (A) xFANCD2-L is recruited to DNA structures that trigger its activation. Bead-coupled ssDNA70 and forkDNA70 structures were incubated in egg extracts for 30 min, and 1 µl of extract was directly analyzed for induction of xFANCD2-L (lanes 1 to 3). Bead-DNA substrates were separated from the extract followed by analysis of bead-DNA (lanes 7 to 9) and extract supernatant (lanes 4 to 6) for xFANCD2 and xFANCA by immunoblotting. Empty beads were used as a negative control (lanes 1, 4, and 7). (B) xFANCD2-L associates with linear and branched dsDNA structures. Bead-coupled ssDNA70, dsDNA70, Y-DNA70, and forkDNA70 were incubated in egg extracts for 30 min, reisolated, and analyzed for bound xFANCD2 and xFANCA by immunoblotting. (C) xFANCD2-L associates with an HJ structure. Bead-coupled HJ-DNA68 or dsDNA68 was incubated in egg extract for 30 min, reisolated, and analyzed for xFANCA and xFANCD2 by immunoblotting. DNA substrate concentrations were as follows: HJ-DNA68 was incubated at 10 pmol/10 µl extract (lane 1), and dsDNA68 was incubated at 10 pmol/10 µl extract (lane 2), 20 pmol/10 µl extract (lane 3), or 30 pmol/10 µl extract (lane 4). The three different molar ratios between HJ-DNA68 and dsDNA68 are 1:1 (compare lanes 1 and 2), 1:2 (compare lanes 1 and 3), and 1:3 (compare lanes 1 and 4), respectively. One microliter of DNA-free extract was used as a size control for xFANCD2-S (lane 5).

DNA structure-bound xFANCD2-L is controlled by the FA core complex. During DNA replication and after DNA damage, monoubiquitination of FANCD2 depends on the FA core complex proteins (12, 28-30, 46, 47). Thus, we tested if monoubiquitination of xFANCD2 in response to the DNA structures in egg extracts depended on the FA core complex. We depleted the core complex protein xFANCA from extracts followed by incubation with plasmid-derived dsDNA fragments. As shown in Fig. 3A, xFANCD2-L was not induced in xFANCA-depleted extracts compared to mock-depleted extracts (compare lanes 2 and 3). Interestingly, levels of activated xMre11-PPP were reduced in xFANCA-depleted extracts compared to the control extracts (compare lanes 2 and 3), whereas depletion of xFANCD2 did not inhibit activation of xMre11-PPP or vice versa (Fig. 3A). Thus, the FA core complex regulates formation of xFANCD2-L and—at least partially—xMre11-PPP in response to dsDNA fragments, but the two events occur independently. Next we tested whether association of xFANCD2 (-S or -L) with DNA structures was FA core complex dependent. We incubated bead-coupled ssDNA70 and forkDNA70 in xFANCA-depleted or mock-depleted extracts. We found that association of xFANCD2-L with forkDNA70 was abrogated in xFANCA-depleted extracts (Fig. 3B, compare lanes 3 and 4). In contrast, binding of xFANCD2-S to ssDNA70 or forkDNA70 was not affected in the absence of xFANCA (Fig. 3B, compare lanes 1 and 2 and lanes 3 and 4). Thus, formation and DNA binding of xFANCD2-L were strictly core complex dependent, whereas xFANCD2-S bound the DNA structures independently of the FA core complex. We also tested whether the absence of xMre11 might affect xFANCD2-L binding to the forked DNA substrate. As expected from the results shown in Fig. 3A, depletion of xMre11 did not affect induction or binding of xFANCD2-L to forkDNA70 (data not shown).


Figure 3
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  		FIG. 3. DNA-induced formation and recruitment of xFANCD2-L are dependent on xFANCA but not on xMre11 or the xATR/xATRIP complex. (A) Formation of xFANCD2-L and xMre11-PPP in response to dsDNA fragments is xFANCA controlled. Egg extracts were depleted of xFANCA (lane 3), xFANCD2 (lane 4), or xMre11 (lane 1) or mock depleted (lane 2) and incubated with dsDNA fragments for 30 min. One microliter of extract was subsequently analyzed for xFANCA, xFANCD2, or xMre11 by immunoblotting. The inset (upper right corner) shows that xFANCD2 and xMre11 protein levels are not affected in an xFANCA-depleted extract (lane 2) compared to a mock-depleted extract (lane 1). (B) Recruitment of xFANCD2-L to forked DNA is dependent on xFANCA but not xATRIP. Egg extracts were depleted of xFANCA (upper panel, lanes 2 and 4) or xATRIP (lower panel, lanes 2 and 4). Mock-depleted extract was used as a control in all depletion experiments (lanes 1 and 3). Depleted extracts were incubated with bead-coupled ssDNA70 (lanes 1 and 2) or bead-coupled forkDNA70 (lanes 3 and 4) for 30 min and analyzed for the indicated proteins. One microliter of DNA-free egg extract was used as a control for protein size (lane 5).

DNA structure-associated xFANCD2-L is xATRIP independent. Apart from the FA core complex, ATR and its regulatory subunit, ATRIP, are also suspected to control FANCD2 function. Recently, we showed that the replication-associated, chromatin-bound form of xFANCD2 (xFANCD2-L) is absent in xATRIP-depleted extracts (46). Moreover, in human cells with ATR deficiency, monoubiquitination of FANCD2 is inhibited (1). We tested whether association of xFANCD2 with bead-bound ssDNA70 or forkDNA70 in egg extracts required the presence of xATRIP. Interestingly, neither binding of xFANCD2-S to either DNA substrate nor association of xFANCD2-L with forkDNA70 was affected in xATRIP-depleted extracts (Fig. 3B, lower panel, compare lanes 1 and 2 and lanes 3 and 4). Thus, xATRIP appears to be dispensable for the induction and recruitment of monoubiquitinated xFANCD2 in response to the DNA structures tested in this assay.

Induction of xFANCD2-L does not require the presence of dsDNA ends. We demonstrated above that xFANCD2 is activated by linear and branched dsDNA substrates; however, it was not evident from these studies whether the DNA structures per se or their respective dsDNA ends triggered xFANCD2 activation. To determine whether dsDNA ends are required to activate xFANCD2, we compared induction of xFANCD2-L in extracts containing equal amounts of (i) circular plasmid DNA (no dsDNA ends), (ii) linearized plasmid DNA (two dsDNA ends per plasmid molecule), and (iii) HaeIII-fragmented plasmid DNA (28 dsDNA ends per plasmid molecule). Interestingly, xFANCD2-L was induced in response to all three DNA substrates (Fig. 4A, lanes 1 to 3). In all three cases, induction of xFANCD2-L was abrogated in xFANCA-depleted extracts, showing that these responses are part of the FA pathway (data not shown). In contrast, xMre11—known to be stimulated by dsDNA ends—was activated in proportion to the number of DNA ends present in the extract (HaeIII-fragmented plasmid >> linearized plasmid; Fig. 4A, lanes 2 and 3) with no Mre11-PPP being induced by circular plasmid DNA (lane 1). We further compared the timing and DNA dose dependency of xFANCD2-L and xMre11-PPP formation in response to linearized and fragmented plasmid dsDNA. As shown in Fig. 4B, xFANCD2-L was detected as early as 15 min following addition of linearized plasmid DNA (at 50 and 25 ng/µl, upper panel, lanes 5 and 6, respectively), whereas only a very weak induction of xMre11-PPP was detectable at a later time point (25 min, 50 and 25 ng/µl, upper panel, lanes 8 and 9). In contrast, in the presence of fragmented plasmid dsDNA, xMre11-PPP was detectable at 15 min at all three DNA concentrations (50, 25, and 10 ng/µl, lower panel, lanes 5 to 7). xFANCD2-L was detected later and only at the highest DNA concentration (25 min, 50 ng/µl, lower panel, lane 8). In summary, activation of xFANCD2 occurred in response to dsDNA but—in contrast to xMre11—did not require the presence of dsDNA ends; moreover, longer dsDNA molecules appeared to be a better stimulus for formation of xFANCD2-L than shorter ones.

Robust induction of xMre11-PPP was not observed at the low concentrations of circular plasmid DNA that stimulated FANCD2-L, but Mre11-PPP was observed at high concentrations of DNA (160 ng/µl) (Fig. 4C). There are at least two possible explanations for this observation. First, the presence of circular dsDNA alone might induce xMre11-PPP, although much less efficiently than dsDNA ends. Alternatively, the circular DNA preparation could contain an undetectable minor amount of nicked or linearized plasmid sufficient to activate xMre11. No nicked or linearized plasmid DNA was detected by agarose gel electrophoresis in the circular plasmid DNA preparation used in the experiment (Fig. 4D, inset, lanes 2 and 3). As shown in Fig. 4D, low concentrations (30 to 60 ng/µl) of nicked and linearized but not circular plasmid DNA induced xMre11-PPP, whereas xFANCD2 was activated by all three DNA substrates (Fig. 4D). Taken together, minor amounts of nicked and/or linearized plasmid DNA contained in circular plasmid preparations might be responsible for the induction of xMre11-PPP when higher concentrations of the circular plasmid DNA are incubated in egg extracts (as seen in Fig. 4C).

Our analysis of small DNA structures incubated in egg extracts showed that an ssDNA substrate (70 nucleotides) did not activate xFANCD2. However, recent work by Zhu and Dutta demonstrated that FANCD2 localizes to ssDNA regions that are generated during illegitimate chromosomal rereplication in the cell (53, 54). To test whether longer stretches of ssDNA might be able to activate xFANCD2, we incubated a circular ssDNA plasmid (M13 phage ssDNA, 7,249 nucleotides) in egg extracts and assayed for induction of xFANCD2-L. As shown in Fig. 4E, the ssDNA plasmid induced activation of xFANCD2 but not xMre11 (lane 3), even at the high DNA concentration used in this experiment (160 ng/µl). This result suggests that longer ssDNA regions can stimulate xFANCD2-L formation. However, since ssDNA molecules are known to fold back on themselves to form irregular double-helical hairpin structures (15), such "pseudo"-dsDNA regions on the M13 ssDNA plasmid might contribute to the stimulation of xFANCD2-L formation.

Circular plasmid-induced activation of the FA pathway is checkpoint and replication independent. One possible mechanism for the circular plasmid induction of FANCD2-L is that plasmid DNA activates a checkpoint in the egg extract, which in turn might induce xFANCD2-L (1, 5, 16). To examine this possibility, we added the checkpoint kinase inhibitor caffeine to extracts containing circular plasmid dsDNA (160 ng/µl). As shown in Fig. 5A, activation of xFANCD2-L was not affected in the presence of caffeine (compare lanes 2 and 4 and lanes 3 and 5). In contrast, induction of xMre11-PPP was inhibited (compare lanes 2 and 4 and lanes 3 and 5), consistent with a previous study showing that caffeine blocks the induction of xMre11-PPP in response to dsDNA fragments (6). In addition, depletion of xATRIP, which functionally blocks the checkpoint mediator xATR (19), had no effect on xFANCD2-L formation (Fig. 5B, compare lanes 1 and 2 and lanes 3 and 4). Thus, activation of xFANCD2-L by circular plasmid DNA is unlikely to be part of a checkpoint response.


Figure 5
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  		FIG. 5. Formation of xFANCD2-L in response to circular dsDNA is replication and checkpoint independent. (A and B) Circular dsDNA induces xFANCD2-L in a checkpoint-independent manner. (A) Egg extracts were incubated with 160 ng/µl circular plasmid dsDNA for 30 min in the absence (lanes 2 and 3) or presence (lanes 4 and 5) of caffeine. Where indicated, extracts were supplemented with the phosphatase inhibitor tautomycin to stabilize xMre11-PPP. Following incubation, 1 µl of extract was analyzed for xFANCD2 and xMre11 by immunoblotting. DNA-free extract was used as a negative control (lane 1). (B) Egg extracts depleted of xATRIP (lanes 2 and 4) or mock-depleted extracts (lanes 1 and 3) were incubated with 160 ng/µl circular plasmid dsDNA. Aliquots were taken at 20 min (lanes 1 and 2) and 60 min (lanes 3 and 4) and assayed for xFANCD2 by immunoblotting. DNA-free extract was used as a size control for xFANCD2 (lane 5). (C and D) Circular dsDNA induces xFANCD2-L in a replication-independent manner. (C) Egg extracts were incubated with 160 ng/µl of circular plasmid dsDNA for 30 min in the absence (lane 2) or presence (lane 3) of geminin and assayed for xFANCD2 and xMre11 by immunoblotting. DNA-free extract was used as a size control for xFANCD2 and xMre11 (lane 1). The efficiency of replication inhibition was measured in a parallel replication assay using an aliquot of the geminin-treated extract supplemented with Xenopus sperm chromatin (inset). (D) Nonactivated, M-phase egg extracts or CaCl2-activated, S-phase egg extracts were incubated with 160 ng/µl circular plasmid dsDNA. Extract aliquots were taken at the indicated time points and analyzed for xFANCD2 and xMre11.

Next, we asked whether replicative DNA synthesis on the circular plasmid template was responsible for inducing xFANCD2-L. When directly added to an S-phase extract, plasmid DNA can be remodeled into a functional nucleus and replicated, although replication efficiency is very low under these conditions (2, 3, 52). We added the replication inhibitor geminin to egg extracts containing circular plasmid DNA (160 ng/µl). As shown in Fig. 5C, geminin did not affect formation of xFANCD2-L or xMre11-PPP in the presence of plasmid DNA. Moreover, plasmid-induced formation of xFANCD2-L and xMre11-PPP was comparable between a replication-competent S-phase extract and an extract unable to support replication (M-phase extract) (Fig. 5D, compare lanes 3 and 5 and lanes 4 and 6), further supporting the idea that formation of xFANCD2-L and xMre11-PPP was not induced by replication of the plasmid DNA.


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DISCUSSION
 
In this study, we showed that the FA pathway is directly triggered in Xenopus egg extracts by DNA structures that mimic DNA intermediates occurring in replicating chromatin. We measured FA pathway activation by monitoring the modified form of xFANCD2, xFANCD2-L, which we determined to be monoubiquitinated xFANCD2. We did not detect a mobility change of xFANCA in response to the DNA structures tested, whereas Thomashevski et al. reported a phosphorylation-induced mobility shift of chromatin-bound human FANCA in cells treated with a DNA interstrand cross-linker (48). However, the authors report that detection of phosphorylated FANCA required several purification steps for cellular chromatin extracts (including chromatography), suggesting that phosphorylated FANCA might not be readily detectable in egg extracts.

We showed that plasmid-derived dsDNA fragments trigger monoubiquitination of xFANCD2. Further analysis revealed that linear or forked dsDNA substrates, both structures with high dsDNA contents, induced formation of xFANCD2-L. In contrast, only the linear but not the forked dsDNA substrate induced formation of xMre11-PPP. Since xMre11-PPP is known to be induced by the presence of dsDNA ends (6), the three ends presented by the forked DNA structure were apparently not recognized by xMre11, possibly because the fork arms are too short to mimic dsDNA breaks recognizable by this repair enzyme. The fact that the forked DNA triggered xFANCD2-L but not xMre11-PPP suggests that the forked or the double-stranded nature of this substrate, but not its DNA ends, activated xFANCD2. The finding that ssDNA and a Y-shaped DNA substrate, structures with no or little dsDNA content, failed to activate xFANCD2-L further supports the idea that the presence of linear or branched dsDNA regions is sufficient to trigger the FA pathway.

The fact that newly formed xFANCD2-L associated with its activating dsDNA structures is an important finding because it provides us with a simple strategy to isolate xFANCD2 in its monoubiquitinated, DNA-associated form from egg extracts. In addition, this new cell-free assay can be adapted to a small-molecule screen for inhibitors or activators of the FA pathway.

Whether xFANCD2-L binds DNA directly is uncertain; however, the fact that recombinant nonubiquitinated human FANCD2 binds DNA in vitro (38) suggests direct DNA binding for the activated FANCD2-L form as well. The forked dsDNA substrate, mimicking a stalled replication fork, reproducibly recruited xFANCD2-L more efficiently than the linear dsDNA substrate (Fig. 2B), suggesting that forked DNA structures might be a preferred target for the FA proteins. Another candidate structure for FA pathway activation was the HJ, a key repair intermediate in HR-mediated repair. The fact that Hef, the archaeal homolog of human FANCM, can unwind and cleave HJs (20, 21) suggested an involvement of the FA pathway in HJ processing. Indeed, xFANCD2-L was induced by—and recruited to—the HJ structure; however, it did not show a preference for this substrate compared to linear dsDNA, hinting that this HJ structure might not be a specific target for xFANCD2-L.

As expected, induction of xFANCD2-L and its subsequent association with the linear and branched dsDNA structures were strictly FA core complex dependent. Surprisingly, the nonmodified xFANCD2-S form associated with all DNA structures that we tested, even the ones that did not trigger FANCD2-L formation. This is unexpected since current evidence, including results from our laboratory, suggests that only FANCD2-L is recruited to sites of damaged DNA in chromatin (12, 33, 46, 47, 49). One possible explanation is that xFANCD2-S can associate with small, naked DNA structures incubated in egg extracts whereas its interaction with DNA is more tightly controlled at the chromatin level. The fact that xFANCD2-S associates with DNA structures even in the absence of a functional FA core complex suggests that xFANCD2-S has low affinity to DNA but becomes specifically activated by monoubiquitination via the FA core complex at dsDNA structures. Somewhat in contrast with our data, Park et al. (38) reported that human recombinant FANCD2-S showed specificity towards certain DNA structures, e.g., an HJ, in vitro. However, observational differences are likely due to the fact that Park et al. studied direct DNA binding of nonmodified recombinant hFANCD2 whereas the Xenopus extract assay monitors DNA association of monoubiquitinated endogenous xFANCD2 in a core complex-controlled context.

We found that the levels of xMre11-PPP were reduced, though not abrogated, in the absence of a functional FA core complex, indicating that either the stability of phosphorylated xMre11 or the efficiency of its phosphorylation is partly controlled by the FA core complex. Interestingly, in human cells the localization of Mre11 into DNA-damage induced chromatin foci is also inhibited when there is a deficient FA core complex (40). A surprising finding was that induction and recruitment of xFANCD2-L to DNA structures were not affected in an xATRIP-depleted extract. This is unexpected because FANCD2-L formation is inhibited in ATR-deficient human cells (1); moreover, we recently demonstrated that chromatin-bound xFANCD2-L is absent in xATRIP-depleted egg extracts. One possible explanation is that xATRIP controls chromatin binding of xFANCD2-L but not its formation and association with the DNA structures used in this study.

Several studies suggest a role for FA proteins in the repair of DNA DSBs. However, our findings do not indicate simple DNA DSBs as the major trigger of the FA pathway. (i) Forked dsDNA activated xFANCD2 but not the DNA DSB-binding protein xMre11 (see above). (ii) Equal amounts of xFANCD2-L were recruited to an HJ structure and to a linear dsDNA structure despite the presence of three free DNA ends in the bead-bound HJ structure versus one free end in the bead-bound linear dsDNA substrate. For further investigation, we compared the xFANCD2-L responses to linear dsDNA substrates of different lengths where the relative number of dsDNA ends was indirectly measurable by the degree of xMre11 hyperphosphorylation. Comparing circular, nicked, linearized, and restriction-fragmented plasmid DNA revealed that xFANCD2-L was activated independently of the number of dsDNA ends. In fact, the linearized plasmid DNA was a stronger trigger for FANCD2-L formation than the shorter plasmid DNA fragments, whereas the opposite was the case for triggering xMre11-PPP formation. Thus, the length of the linear dsDNA substrate rather than its ends influenced the degree of xFANCD2-L induction. Interestingly, low concentrations of nicked and linearized—but not intact circular—plasmid DNA activated xMre11, hinting that nicks in dsDNA can also trigger xMre11-PPP formation.

To determine how the circular plasmid triggered activation of xFANCD2-L, we tested two hypotheses: induction of FANCD2-L could be (i) part of a plasmid-induced checkpoint in the extract or (ii) due to replication of the plasmid DNA in the extract. The induction of xFANCD2-L was unaffected by caffeine or the absence of the xATR/xATRIP checkpoint kinase complex, suggesting that plasmid-induced FA pathway activation was checkpoint independent. Likewise, geminin treatment or use of replication-incompetent M-phase extracts had no inhibitory effect on xFANCD2-L, ruling out a replicating plasmid-mediated induction of xFANCD2-L. We are currently investigating if processing or remodeling of the plasmid in the extracts is involved in triggering the ubiquitination of FANCD2.

Interestingly, a circular ssDNA plasmid was also able to trigger activation of xFANCD2, but not xMre11. Like the dsDNA plasmid, the ssDNA plasmid induced xFANCD2-L independently of replication or checkpoint activation (data not shown). Since ssDNA plasmids are known to fold into dsDNA-like hairpin structures, it is at this point not possible to determine whether the long ssDNA stretches of the M13 ssDNA plasmid or its double-stranded hairpin regions are responsible for xFANCD2 activation. However, the fact that FANCD2 localizes specifically to ssDNA regions generated by rereplication in human cells suggests that longer ssDNA regions are in fact a trigger for xFANCD2-L induction (54).

In summary, we showed that the FA pathway is inducible in a replication-free environment in the presence of defined DNA structures. Monoubiquitinated xFANCD2 associates with dsDNA structures in an xFA core complex-dependent but xATRIP-independent manner. The DNA substrate specificity of xFANCD2-L suggests that a rather broad spectrum of single- and double-stranded linear or branched DNA lesions—but not simple DNA DSBs—activates the FA pathway. Taken together, our results suggest a model where FANCD2 activation occurs during DNA replication when dsDNA is exposed at the replication fork because of either stalling or damage.


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ACKNOWLEDGMENTS
 
We thank Vincenzo Costanzo, Hua Lu, and Weidong Wang for many helpful discussions and critical reading of the manuscript. We are indebted to J. Gautier and K. Cimprich for generously sharing antibodies for Xenopus proteins.

A. Sobeck has received funding from the American Heart Association (0520117Z). M. Hoatlin has received funding from the National Institutes of Health (CA112775) and the Fanconi Anemia Research Fund.


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FOOTNOTES
 
* Corresponding author. Mailing address: Biochemistry and Molecular Biology and Molecular and Medical Genetics, Oregon Health and Science University, 3181 S. W. Sam Jackson Park Rd., Medical Research Building, Rm. 518 (mailcode L224), Portland, OR 97239. Phone: (503) 494-1123. Fax: (503) 494-8393. E-mail: hoatlinm{at}OHSU.edu Back

{triangledown} Published ahead of print on 9 April 2007. Back


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Molecular and Cellular Biology, June 2007, p. 4283-4292, Vol. 27, No. 12
0270-7306/07/$08.00+0     doi:10.1128/MCB.02196-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




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