Persistent protein obstacles on genomic DNA, such as DNA-protein crosslinks (DPCs) and tight nucleoprotein complexes, can block replication forks. DPCs can be removed by the proteolytic activities of the metalloprotease SPRTN or the proteasome in a replication-coupled manner; however, additional proteolytic mechanisms may exist to cope with the diversity of protein obstacles. Here, we show that FAM111A, a PCNA-interacting protein, plays an important role in mitigating the effect of protein obstacles on replication forks. This function of FAM111A requires an intact trypsin-like protease domain, the PCNA interaction, and the DNA-binding domain that is necessary for protease activity in vivo. FAM111A, but not SPRTN, protects replication forks from stalling at poly(ADP-ribose) polymerase 1 (PARP1)-DNA complexes trapped by PARP inhibitors, thereby promoting cell survival after drug treatment. Altogether, our findings reveal a role of FAM111A in overcoming protein obstacles to replication forks, shedding light on cellular responses to anti-cancer therapies.
Collision of the DNA replication machinery with replication fork obstacles causes replication fork stalling and double-strand DNA breaks (DSBs), posing a great threat to genome integrity1,2,3. In particular, protein obstacles such as DNA- protein crosslinks (DPCs) have deleterious effects on replication forks due to their bulky nature. DPCs can form by various mechanisms, with differing sizes and DNA-crosslink structures4,5,6. For example, crosslinking chemicals, such as formaldehyde and reactive oxygen species, covalently link proteins to DNA non-specifically. In contrast, abortive enzymatic reactions can produce specific DPCs as represented by topoisomerase 1 cleavage complexes (TOP1ccs), which result from trapping of TOP1 covalently bound to DNA in an intermediate step of the enzyme reaction. Because of the severe consequences of blocking replication forks, induction of DPCs or nucleoprotein complexes is a common mechanism for anti-cancer drugs. TOP1 inhibitors, such as topotecan and its prototype camptothecin (CPT), poison TOP1, thereby inducing cytotoxic levels of TOP1ccs7. On the other hand, poly(ADP-ribose) polymerase (PARP) inhibitors (PARPis) induce nucleoprotein complexes by trapping PARP1 protein at DNA single-strand breaks (SSBs) during the SSB repair process8,9,10. Through strong DNA binding that mimics a DPC, PARP1-DNA complexes block DNA replication and cause DSBs11,12,13. Since blocked replication forks and DSBs require the homologous recombination (HR) pathway for repair, PARPis are particularly toxic to HR-deficient tumors with BRCA1/2 mutations14,15.
The discovery of the metalloproteases, Wss1 in yeasts and SPRTN in higher eukaryotes, revealed a previously unrecognized contribution of proteolysis to DPC repair16,17,18,19,20,21,22,23,24,25. The SprT metalloprotease domain of SPRTN contains a Zn2+-binding subdomain that binds single-strand DNA (ssDNA), which activates SPRTN26. SPRTN localizes to replication forks20,27 and degrades DPC proteins upon replication fork collisions28,29. SPRTN deficiency causes progeria and an increased incidence of liver cancer in humans and mice, highlighting the importance of DPC repair27,30,31,32. In addition to SPRTN, DPCs can also be resolved using the proteasome, which can degrade DPCs following replication-coupled polyubiquitination of DPC proteins29. Furthermore, replication-independent sumoylation of DPC proteins has also been reported29,33 and ACRC/GCNA family SprT proteases are involved in DPC repair in germline cells33. On the other hand, it is unknown whether these DPC repair mechanisms play a role in preventing replication fork stalling at non-covalently linked protein–DNA complexes. Indeed, we previously reported that Sprtn hypomorphic mouse embryonic fibroblasts (MEFs) are not hypersensitive to PARPis, implying that SPRTN is unlikely to be involved in removal of PARP1-DNA complexes27. Therefore, it is currently unknown whether a cellular mechanism exists to remove trapped PARPs, but such a system would be expected to undermine the therapeutic effect of PARPis.
In this study, we demonstrate that the putative serine protease FAM111A (Family with sequence similarity 111 member A) protects replication forks from stalling at PARP1-DNA complexes and TOP1ccs. FAM111A was originally identified as a replication fork protein by nascent chromatin capture proteomics and shown to interact with PCNA through its PCNA-interacting peptide (PIP) box34. Although it was proposed that FAM111A plays a role in PCNA loading, its function at ongoing replication forks was unknown. The presence of a trypsin-like protease domain in FAM111A and its localization to replication forks prompted us to investigate whether FAM111A is involved in DPC removal during DNA replication. Our findings uncover a critical role of FAM111A in promoting DNA replication fork progression not only at DPCs, but also at PARP1-DNA nucleoprotein complexes.
FAM111A KO cells are sensitive to PARP and TOP1 inhibitors
To investigate whether FAM111A is important for mitigating the effect of protein obstacles to replication forks, we examined whether FAM111A depletion affects sensitivities to a PARPi and a TOP1 poison. FAM111A-knockout (KO) by CRISPR/Cas9 (Fig. 1a, Supplementary Fig. 1, and Supplementary Table 1) rendered cells hypersensitive to the PARPi niraparib, which exhibits strong PARP-trapping effect9, and the TOP1 inhibitor CPT (Fig. 1b). Similar results were also obtained with additional FAM111A KO clones obtained using CRISPR/Cas9 targeting different sequences of the gene (Supplementary Figs. 1, 2a, b and Supplementary Table 1). Furthermore, FAM111A KO cells were sensitive to another PARPi with a strong PARP-trapping effect10, talazoparib (Supplementary Fig. 2c). In addition, FAM111A KO cells showed mild sensitization to etoposide (a topoisomerase II inhibitor that covalently traps TOP2), 5-aza-dC (a DNMT inhibitor that traps DNMT isoenzymes on DNA), and formaldehyde (Supplementary Fig. 2d). On the other hand, FAM111A KO cells were not sensitized to ionizing radiation (IR) or cisplatin (a DNA crosslinker) (Fig. 1b and Supplementary Fig. 2d).
The hypersensitivity of FAM111A KO cells to niraparib was also associated with a marked increase in apoptotic cell death as shown by Annexin V staining (Fig. 1c, d). Activation of the HR pathway was intact in FAM111A KO cells when they were treated with PARPi as shown by RAD51 focus formation (Supplementary Fig. 2e, f), indicating that an HR defect does not account for the PARPi hypersensitivity of FAM111A KO cells. This conclusion is also supported by the fact that FAM111A KO cells were not sensitive to IR (Fig. 1b), which relies on an intact HR pathway for optimal repair. Taken together, these results suggest that FAM111A protects cells from several anti-cancer drugs, especially PARP and TOP1 inhibitors.
FAM111A directly binds to ssDNA through its central region
Because FAM111A localizes to replication forks34, we hypothesized that FAM111A might have DNA-binding activity. We tested this possibility by electrophoretic mobility shift assays (EMSA) using purified full-length FAM111A protein (Supplementary Fig. 3a) and various forms of labeled DNA oligonucleotides, including ssDNA, double-strand (ds) DNA, Y fork, and dsY fork (Supplementary Fig. 3b). FAM111A displayed robust binding to ssDNA and Y-fork DNA, weaker binding to dsDNA, and undetectable binding to dsY fork DNA (Fig. 2a). These results suggest that FAM111A exhibits preferential binding to ssDNA, as ssDNA structures were common between the ssDNA and Y fork DNA oligos used in these experiments. Supershift assays using anti-FAM111A antibody verified that the mobility shift was due to FAM111A rather than possible contaminants in the protein preparation (Fig. 2b). Binding of FAM111A to ssDNA was also confirmed using fluorescence polarization assays, which showed an equilibrium dissociation constant (Kd) of ~60 nM for ssDNA binding (Fig. 2c).
We next sought to map the DNA-binding domain of FAM111A. EMSA using various fragments of FAM111A (Fig. 2d and Supplementary Fig. 3c) revealed that the central region (176–282) was sufficient to bind to ssDNA (Fig. 2e). On the other hand, the ssDNA-binding activity of the N-terminal FAM111A fragment (1–282) was diminished with varying degrees when a part of the central region was deleted (Supplementary Fig. 3d–f). These results suggested that the central region (176–282) is necessary and sufficient for the ssDNA binding of FAM111A.
To identify amino acid residues in the FAM111A central region that are crucial for ssDNA binding, we introduced point mutations in aromatic or positively charged amino acid residues based on a previous report showing that those amino acid residues are involved in ssDNA binding35. We particularly focused on amino acids conserved among species or stretches of positively charged amino acids (Supplementary Fig. 3g). EMSA using MBP-FAM111A 1–282 with point mutations revealed that the F231A mutation had the largest effect on ssDNA binding among the mutants tested (Supplementary Fig. 3h, i). The F231A mutation decreased DNA binding to both ssDNA and Y-fork substrates (Fig. 2f), and similar results were obtained with the minimum DNA-binding domain (176–282) (Supplementary Fig. 3j-l). All of these data suggest that Phe231 plays a key role in FAM111A binding to ssDNA.
Disease-causing mutations enhance FAM111A cleavage in vivo
FAM111A contains a trypsin-like domain at its C-terminus with an intact catalytic triad consisting of His385, Aps439 and Ser541, a characteristic feature of the S1 serine protease family36,37. Given that proteases often exhibit autocleavage activity, we set out to find processed fragments of FAM111A in vivo to study the putative protease activity. When exogenous FAM111A was expressed in 293T cells, which lack endogenous FAM111A expression for an unknown reason (Supplementary Fig. 4a), a small FAM111A fragment (~40 kDa), as well as full-length protein, was detected on the immunoblot using the anti-FAM111A antibody that recognizes the N-terminal region (Fig. 3a). This fragment was not observed with the mutant that contains a substitution of the active site serine to an alanine (S541A). These results raised the possibility that FAM111A might have autocleavage activity in vivo. Interestingly, germline mutations have been found in FAM111A in patients with Kenny–Caffey syndrome (KCS) and osteocraniostenosis (OCS)38,39,40,41, and it has been postulated that these mutations may cause gain-of-function FAM111A activity because the mutations are always heterozygous and no loss-of-function mutations (i.e., deletions and truncations) have been reported38. We therefore tested whether the mutations found in KCS and OCS patients produce constitutively active FAM111A. For all the patient mutants tested (R569H and Y511H for KCS; S342del and D528G for OCS), the levels of full-length protein were reduced concomitant with a large increase in the levels of the processed fragment (Fig. 3b, lanes 4, 6, 8, and 10). Addition of an active site mutation (Ser541, S541A) to these patient mutants restored full-length protein expression and eliminated the processed fragment (Fig. 3b, lanes 5, 7, 9, and 11). These results suggest that the FAM111A patient mutants are constitutively active forms that result in the conversion of the full-length protein to smaller fragments through a possible autocleavage mechanism.
FAM111A activity exhibits chymotrypsin-like specificity
The S1 family serine proteases exhibit three major types of substrate specificities, which are determined by recognition of the P1 substrate residue (the amino acid residue preceding the cleavage site) by the S1 pocket of the enzyme. The trypsin-type prefers positively charged residues at the P1 position, whereas the chymotrypsin-type and the elastase-type prefer large and small hydrophobic residues, respectively36,37. To gain more information on the substrate specificity of the in vivo FAM111A cleavage, we set out to determine the in vivo cleavage site of FAM111A. We expressed recombinant FAM111A proteins in insect cells and purified with the C-terminal Strep-tag. As was the case with FAM111A expressed in human cells, recombinant FAM111A produced in insect cells displayed a processed product that was more prominent with the FAM111A R569H mutant and absent when the active site was mutated (Fig. 3c). N-terminal peptide sequencing of the C-terminal fragment from the FAM111A R569H mutant identified the cleavage site to be C-terminal to Phe334 (P1) (Fig. 3d). Cleavage at this site is predicted to produce 38.4 kDa N-terminal and 31.5 kDa C-terminal fragments, consistent with the observed fragment sizes (Fig. 3a–c). Substitution of Phe334 with glycine or arginine diminished the in vivo cleavage of FAM111A (Fig. 3e), suggesting that Phe334 contributes to the specificity of the cleavage site. The hydrophobic residue (Phe334) at the P1 position implies chymotrypsin-type specificity for this cleavage.
Chymotrypsin-like proteases typically contain a hydrophobic S1 pocket36,37. In examining the structure model of the FAM111A trypsin-like domain based on the plant Deg1 protease38,42, we indeed found a notable hydrophobic feature, Phe536, which is highly conserved among species, in the predicted S1 pocket (Supplementary Fig. 4b). In silico docking experiments using peptide sequences with the native cleavage site RTTFGKV as well as the mutant cleavage sites RTTRGKV and RTTGGKV (underlines indicate the P1 residue) showed the P1 Phe substrate displayed by far better docking score (−12.9 kcal mol−1) compared with either the P1 Arg or P1 Gly substrates (−10.5 and −8.5 kcal mol−1, respectively). Close examination of the S1 pocket revealed that the docked Phe peptide resulted in an edge-to-face ring stacking interaction between the P1 Phe334 and conserved Phe536 (Fig. 3f). By contrast, in the docking experiment with the P1 Arg mutant peptide, the Arg side chain was expelled from the S1 pocket during the energy minimization and relaxation protocol. These observations indicate that FAM111A is a serine protease with chymotrypsin-like specificity and strongly suggest that FAM111A exhibits autocleavage activity in vivo.
FAM111A autocleavage can occur in trans
Autocleavage can occur intramolecularly (in cis) and intermolecularly (in trans). Co-immunoprecipitation assays using FAM111A with different tags (3xFlag or EGFP) showed interaction between 3xFlag-FAM111A and EGFP-FAM111A (Fig. 3g), raising the possibility of an intermolecular mechanism. To directly assess this possibility, we co-expressed the constitutively active form of FAM111A (R569H) and its inactive counterpart (R569H/S541A) with different tags and tested whether constitutively active FAM111A causes cleavage of inactive FAM111A. Indeed, cleavage of the inactive FAM111A occurred when the constitutively active FAM111A was co-expressed (3xFlag-FAM111A R569H/S541A with EGFP-FAM111A R569H and vice versa) (Fig. 3h, lanes 6 and 7), demonstrating that autocleavage of FAM111A can occur intermolecularly.
FAM111A autocleavage requires an intact DNA-binding domain
We next determined whether domains other than the trypsin-like domain are required for the FAM111A autocleavage. Introduction of the F231A mutation, which disrupts ssDNA binding of FAM111A (Fig. 2f, Supplementary Fig. 3j–l), but not YFAA, which diminishes PCNA interaction34, prevented autocleavage of the constitutively active form of FAM111A (R569H) (Fig. 3i). Co-immunoprecipitation assays ruled out the possibility that the F231A and S541A mutations disrupted intermolecular interaction (Supplementary Fig. 4c). These data suggest that FAM111A activity in vivo is dependent on an intact ssDNA-binding domain.
Trapping-dependent PARPi hypersensitivity after FAM111A KO
In principle, PARPis can affect cell proliferation and survival by (i) diminishing formation of poly(ADP-ribose) polymer (inhibition mechanism) and (ii) blocking DNA replication through PARP1-DNA complex formation (trapping mechanism). The inhibition mechanism would be mimicked by PARP1 KO, while the trapping mechanism would be eliminated by PARP1 KO. To test which mechanism underlies the PARPi hypersensitivity conferred by FAM111A KO, we knocked out PARP1 by CRISPR/Cas9 in the FAM111A WT and FAM111A KO backgrounds (Fig. 4a, Supplementary Fig. 1 and Supplementary Table 1). FAM111A/PARP1 double KO (DKO) cells, as well as PARP1 KO cells, were viable and indistinguishable from parental cells in proliferation assays (Fig. 4b). These data argue against PARP1 inhibition being the major mechanism of the hypersensitivity to PARPis in FAM111A KO cells. On the other hand, PARP1 knockout conferred complete niraparib resistance to both FAM111A WT and FAM111A KO cells (Fig. 4c). Taken all together, these data suggest that sensitization to PARPis by FAM111A KO is dependent on PARP1 trapping.
FAM111A deficiency causes TOP1cc accumulation
The hypersensitivity of FAM111A KO cells to CPT (Fig. 1a, b) prompted us to hypothesize that FAM111A might also be involved in DPC repair in vivo. We tested this possibility by examining the levels of TOP1cc in FAM111A KO cells in the absence of CPT treatment. We found that FAM111A KO cells displayed increased TOP1cc foci compared with wild-type cells (Fig. 4d–f). Increased TOP1cc foci were also observed in U2OS cells after knockdown of FAM111A by RNAi (Supplementary Fig. 5a–c). TOP1cc accumulation in the FAM111A KO cells was diminished by exogenous expression of wild-type FAM111A, but not the active site (S541A) or the PIP-box (YFAA) mutants (Fig. 4d–f), indicating the importance of these domains to prevent TOP1cc accumulation. These results indicate that FAM111A is important for the repair of TOP1ccs, providing an explanation for the hypersensitivity of FAM111A KO cells to CPT.
FAM111A prevents fork stalling during PARPi or CPT treatment
Given that FAM111A localizes to replication forks34, we assessed the importance of FAM111A in protecting replication forks from protein obstacles imposed by CPT or PARPis. For these experiments, we employed DNA combing assays and examined the effect of the drugs on replication fork movement. Nascent DNA was sequentially labeled with CldU and then with IdU in the presence or absence of drugs (Fig. 5a). In the absence of drug treatment, replication fork movement was largely unaffected by FAM111A KO as seen in the comparable CldU tract lengths in FAM111A WT and KO cells (Fig. 5b). Although it has been reported that FAM111A may be involved in the PCNA loading34, the amounts of PCNA on chromatin were largely unaffected in FAM111A KO cells (Supplementary Fig. 6a). In contrast, replication fork movement was severely affected by niraparib or CPT treatment in FAM111A KO cells as evidenced by the reduced ratios of IdU- and CldU-labeled tract lengths (Fig. 5c, d). Exogenous expression of wild-type FAM111A, but not the S541A, or YFAA, rescued the effect of niraparib treatment in FAM111A KO cells (Fig. 5e), suggesting that the putative protease activity and PIP box of FAM111A are important to mitigate the effect of trapped PARPs. In addition, the F231A mutant did not rescue the replication fork defects during niraparib or CPT treatments (Fig. 5f, g), demonstrating the importance of the FAM111A DNA-binding domain. Importantly, the percentage of collapsed forks induced by hydroxyurea (HU) treatment was not increased in FAM111A KO cells (Supplementary Fig. 6b, c), suggesting that FAM111A does not regulate the stability of stalled replication forks. In addition, the S541A and F231A mutants of FAM111A localized properly to replication forks as assessed by iPOND assays (Supplementary Fig. 6d). These results demonstrate the importance of FAM111A in overcoming protein obstacles imposed by CPT and PARPis.
We reported previously that Sprtn hypomorphic MEFs are hypersensitive to CPT but not niraparib27. Together with the drug sensitivity data presented above (Fig. 1a, b), the previous observations raised the possibility that SPRTN might engage with TOP1cc but not PARP1-DNA complexes, while FAM111A might act on both. To investigate this possibility, we targeted SPRTN with CRISPR/Cas9 in HAP1, the same human cell line as FAM111A KO cells. This resulted in a SPRTN hypomorphic cell line (indicated as SPRTN CRISPR), which exhibited greatly reduced SPRTN levels due to KO in one allele and an eight-amino-acid deletion in the metalloprotease domain in the other allele (Fig. 5h, Supplementary Fig. 1 and Supplementary Table 1). Consistent with our previous study in mouse cells27, the SPRTN CRISPR cells were sensitive to CPT, validating impairment of SPRTN function, yet they were not hypersensitive to niraparib (Supplementary Fig. 6e). DNA combing assays revealed that SPRTN CRISPR cells exhibited increased replication fork stalling with CPT but not with niraparib, whereas FAM111A KO displayed fork stalling with both drugs (Fig. 5i). These results demonstrate the unique function of FAM111A in processing PARP1-DNA complexes.
FAM111A KO causes enhanced DNA damage after PARPi treatment
Finally, we investigated how FAM111A KO confers increased PARPi sensitivity and cell death (Fig. 1). If increased replication fork stalling (Fig. 5c) is the underlying cause, FAM111A KO cells are expected to experience cell-cycle defects and increased levels of DNA damage after PARPi treatment. Consistent with this prediction, FAM111A KO cells, which showed a modest reduction of G1 phase and increase in S and G2/M phases in the absence of drug treatment, exhibited profound accumulation in G2/M phase upon treatment with niraparib (Fig. 6a, b). Moreover, FAM111A KO cells exhibited higher levels of phospho-Chk1, but not phospho-Chk2, than wild-type cells after niraparib treatment (Fig. 6c), indicating augmented activation of the ATR/Chk1 pathway, which typically responds to DNA replication stress. In addition, higher levels of DNA damage were observed in FAM111A KO cells than in wild-type cells after treatment with niraparib, as shown by immunostaining with the DNA damage marker γH2AX (Fig. 6d, e). Collectively, these results suggest that the absence of FAM111A causes replication fork stalling in S phase after PARPi treatment, leading to DNA damage, cell-cycle arrest in G2/M phases, and eventually cell death.
Advancing DNA replication forks face many obstacles due to proteins that are covalently crosslinked or tightly bound to DNA. In this study, we discovered that the replication fork protein FAM111A is important for resolving replication stalling caused by protein obstacles. We provide evidence for the protease activity of FAM111A in vivo, which is dependent on the DNA-binding domain identified in this study (Figs. 2 and 3). FAM111A KO cells exhibit TOP1cc accumulation and PARPi hypersensitivity that is dependent on PARP-trapping (Fig. 4). In the absence of FAM111A, protein obstacles induced by a TOP1 poison or a PARPi causes increased replication stalling (Fig. 5). The TOP1cc removal and fork protection by FAM111A require the PIP box and an intact trypsin-like domain (Figs. 4 and 5). Collectively, we propose a model that the FAM111A PIP box tethers the trypsin-like protease domain to replication forks to protect them from protein obstacles (Fig. 6f).
The role of FAM111A in resolving protein obstacles at replication forks is reminiscent of SPRTN’s function in replication-coupled DPC repair. Both proteins contain a protease domain, a PIP box, and a DNA-binding domain that is necessary for protease activation. In addition, both proteins localize to replication forks and facilitate DNA replication at DPCs, including TOP1ccs induced by CPT. Despite these similarities, however, a striking difference was noted between FAM111A and SPRTN in the response to PARPis. Whereas CPT induces protein obstacles covalently crosslinked to DNA, PARPis are generally thought to trap PARPs non-covalently9,10,11,12,13, although covalent PARP1-DNA complexes have also been reported43,44. Interestingly, it was recently demonstrated in Xenopus egg extracts that SPRTN activation requires polymerase collision with DPCs followed by helicase bypass of DPCs29,45. Therefore, the difference between SPRTN and FAM111A in the response to PARPis might be attributable to whether the fork-blocking proteins are covalently crosslinked to DNA or not. SPRTN might be specialized for protein obstacles covalently bound to DNA, while FAM111A might have a broader role in responding to protein obstacles regardless of the DNA-crosslinking status. Although further investigations are necessary to understand how different types of protein obstacles are cleared at replication forks, our study suggests that cells are equipped with multiple mechanisms to deal with various protein obstacles during DNA replication.
Our study strongly suggests that FAM111A functions as a protease in vivo. A mutation in the putative active site serine abolishes its function in TOP1cc repair and replication during niraparib treatment (Figs. 4d–f and 5e). Although wild-type FAM111A shows limited protease activity when expressed in cells, the FAM111A mutants found in KCS or OCS patients are constitutively active and display increased autocleavage activity in vivo (Fig. 3b). We speculate that FAM111A might undergo activation only when necessary, and the KCS/OCS mutants might have bypassed the required activation mechanism. Homology-based structure modeling suggests that the FAM111A trypsin-like domain is similar to the Deg/HtrA family proteases38, which contain one or two PDZ domains that play regulatory roles during protease activation in response to various cellular stresses, including high temperature, abnormal pH, or unfolded proteins42,46,47. Although FAM111A lacks a PDZ domain, we instead identified a DNA-binding domain in FAM111A that is necessary for FAM111A activity in vivo (Figs. 2 and 3i). This raises the possibility that FAM111A might undergo activation by sensing ssDNA structures as has been demonstrated for SPRTN. However, ssDNA-binding alone might not be a sufficient mechanism to restrict FAM111A activation to events of replication fork collisions with protein obstacles. Consistent with this notion, we have not been able to demonstrate protease activities in vitro using purified recombinant FAM111A even in the presence of multiple DNA structures. One possibility is that FAM111A might undergo additional regulation in vivo to become fully active, for example, through post-translational modifications, cofactor binding, or conformational changes. Interestingly, many Deg/HtrA proteases dramatically change their oligomer status during activation, in some cases forming cages with the protease active sites facing inward47,48. Indeed, our results showed that FAM111A proteins interact with each other in vivo (Fig. 3g). While the activation mechanism is still unclear, our study suggests that FAM111A might be a stress-responsive protease similar to the Deg/HtrA family proteases, but one that responds to replication stress imposed by protein obstacles.
The present study also identifies FAM111A deficiency as a mechanism of hypersensitivity to PARPis. Because FAM111A deficiency augments DNA damage induced by PARPis (Fig. 6d, e), FAM111A inhibition might be a viable strategy to enhance PARPi killing in HR-deficient tumors and to expand PARPi therapies to HR-proficient tumors. Moreover, our data showed that FAM111A levels vary among cancer cell lines (Supplementary Fig. 4a). Therefore, determining the FAM111A status in tumors might be useful because a therapeutic advantage can be gained through the selective vulnerability of FAM111A-deficient cells to PARPis.
Finally, the present results shed important new light on previous studies of FAM111A. Our data suggest that FAM111A mutations found in KCS or OCS patients lead to constitutive protease activity (Fig. 3b). This observation raises two possibilities for the potential consequences of disease-associated mutations. One is that hyperactivation of FAM111A causes abnormal degradation of DNA-binding proteins in KCS and OCS, and the other is that hyperactivation causes a FAM111A deficiency through hyper-autocleavage. In fact, FAM111A forms oligomers and can degrade FAM111A proteins in trans (Fig. 3g, h), so that it is possible that the heterozygous mutations in KCS and OCS cause depletion of total FAM111A protein through a dominant-negative effect. Another implication of our study is that it might help understand the role of FAM111A as an antiviral defense mechanism. It has been demonstrated that FAM111A is a host range restriction factor for SV40 and mutant orthopoxviruses49,50. Given the recent report showing that FAM111A localizes to SV40 viral replication centers51, it is possible that FAM111A might restrict viral replication by recognizing and degrading viral nucleoprotein complexes.
In conclusion, we demonstrated that FAM111A is important for DNA replication at protein obstacles. This study not only sets new frameworks for future studies on DPC repair, replication fork regulation, and anti-cancer therapies, but also provides new insight into the role of FAM111A in genetic diseases and viral defense mechanisms.
Human chronic myelogenous leukemia cell line HAP1 was purchased from Horizon Discovery. Human embryonic kidney cell line 293T, human osteosarcoma cell line U2OS, human proximal tubular cell line HK2, human lung fibroblast cell line MRC-5, human cervical adenocarcinoma cell line HeLa, human colorectal carcinoma cell line HCT116, human mesothelioma cell line H226, human fibrosarcoma cell line HT1080, and human hepatocellular carcinoma cell line HepG2 were obtained from American Type Culture Collection. These cell lines were cultured in respective medium: Iscove’s Modified Dulbecco’s Medium for HAP1, Dulbecco’s modified Eagle’s medium for 293T and HeLa, McCoy’s 5A medium for U2OS and HCT116, Eagle’s minimum essential medium for MRC-5, HT1080, and HepG2. All of these media were supplemented with 10% fetal bovine serum (FBS). For HT1080 and HepG2, the medium was also supplemented with MEM non-essential amino acids (Corning) and sodium pyruvate (Corning). HK2 cell line was cultured in keratinocyte serum free medium (Invitrogen).
For mammalian cell expression, a cDNA fragment encoding human FAM111A was cloned in the following vectors: pLVX2-IRES-puro (no epitope tag), pLVX3-IRES-puro (for an N-terminal 3xFlag tag), and pLVX6-IRES-puro (for an N-terminal EGFP tag). For insect cell expression, a DNA fragment encoding FAM111A with tandem Strep tags (FAM111A-strep) was synthesized with codon-optimization for expression in S. frugiperda (Thermo Fisher Scientific) (Supplementary Table 2). A DNA fragment encoding FAM111A with an N-terminal tandem Strep-tag (Strep-FAM111A) was generated by PCR. The ORFs were cloned into pDEST20 (for an N-terminal GST tag) and pFastBac NT-B. For bacterial expression, a DNA fragment encoding FAM111A 1–282 was generated by gene synthesis with codon-optimization for expression in E. coli (Thermo Fisher Scientific) (Supplementary Table 2) and used as a PCR template to amplify DNA fragments for FAM111A 1–99, 100–175, and 176–282. A DNA fragment encoding FAM111A 283–611 was amplified from human FAM111A cDNA. These PCR fragments were cloned in pMALII-c2x for expression of MBP fusion proteins in E. coli. Point mutations were introduced using the Quickchange mutagenesis kit (Agilent) or by PCR. All plasmids were confirmed by Sanger sequencing. Cloning primers are summarized in Supplementary Table 3.
Transfection and viral infection
Plasmid transfection was performed using Lipofectamine 2000 (Thermo Fisher Scientific) or Turbofectin 8.0 (OriGene) according to suppliers’ instructions. Lentiviruses were produced by transfecting 293T cells with a lentiviral and two packaging plasmids (psPax2 and pMD2.G). Culture media containing lentiviral particles were harvested after 48 h and used to infect cells in the presence of 4 µg ml−1 polybrane. After selection with 2 µg ml−1 puromycin for 48 h, cells were cultured in the absence of puromycin for at least 24 h. siRNA transfection was performed using RNAiMAX (Thermo Fisher Scientific). The target sequences of siRNAs are as follows: siControl, 5′-CGUACGCGGAAUACUUCGA-3′; siFAM111A.271, 5′-CUAAAGAGCAACAGAAUAA-3′; siFAM111A.1213, 5′-CGAUUAAAGUAGUGAAACU-3′. siRNA oligo sequences are shown in Supplementary Table 3.
Western blotting and co-immunoprecipitation
Cells were lysed using NP-40 lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 0.1% NP-40, 10% glycerol, 5 mM EDTA, 50 mM NaF, 1 mM Na3VO4) supplemented with protease inhibitor cocktail (Sigma). For FAM111A autocleavage assays, cells were lysed in SDS lysis buffer (PBS supplemented with 1% SDS and protease inhibitor cocktail). Cell lysates containing 30 µg protein were run on an SDS-PAGE gel and analyzed by Western blotting. Primary antibodies used were rabbit anti-FAM111A (Abcam, ab184572, 1:2000), rabbit anti-FAM111A (Sigma, HPA040176, 1:500), mouse anti-PARP1 (C-2-10, kind gift from Guy Poirier, 1:1000), mouse anti-SPRTN52, mouse anti-β-actin (Sigma, A5316, 1:5000), rabbit anti-Chk1 (Epitomics, 1740–1, 1:2000), rabbit anti-phospho-Chk1 (S345) (Cell Signaling Technology, #2348, 1:1000), mouse anti-Chk2 (BD Bioscience, 611570, 1:1000), rabbit anti-phospho-Chk2 (T68) (Cell Signaling Technology, #2197, 1:1000), mouse anti-PCNA (Santa Cruz, sc-56, 1:2000), rabbit anti-Histone H3 (Cell Signaling Technology, #9715, 1:1000), mouse anti-Flag (Sigma, F1804, 1:1500), rabbit anti-Flag antibody (Cell Signaling Technology, #2368, 1:1000), and mouse anti-GFP (Santa Crus, sc-9996, 1:1000). Chemiluminescence signals were imaged by X-ray film exposure or ChemiDoc (BioRad).
For co-immunoprecipitation experiments, 293T cells were transfected with expression plasmids for 48 h and lysed with NP-40 lysis buffer supplemented with proteinase inhibitor cocktail (Sigma) and 41 U ml−1 Benzonase (Novagen). Cell lysates containing ~1 mg protein were incubated with anti-Flag beads (Sigma) for 4 h at 4 °C and the beads were washed five times with NP-40 lysis buffer. Precipitated proteins were analyzed by Western blotting.
Generation of knockout cell lines
CRISPR/Cas9-based gene knockout was performed using pX335 (kind gift from Feng Zhang, Addgene #42335) for FAM111A and PARP1, and eSpCas9(1.1) (kind gift from Feng Zhang, Addgene #71814) for SPRTN. For CRISPR/Cas9 using the nickase Cas9 (pX335), the following pairs of gRNAs were used: FAM111A, set 1 (gRNA1 and gRNA2) and set 2 (gRNA3 and gRNA4); PARP1, gRNA1, and gRNA2. The target sequences of gRNA are as follows: FAM111A gRNA1, 5′-ATCAACCCACTAAGTGGCAC-3′; FAM111A gRNA2, 5′-CATATTATTGGCCATCCATAA-3′; FAM111A gRNA3, 5′-ATTATGGTTGCCCACTTACT-3′; FAM111A gRNA4, 5′-CAATGTGTAAGGGTGACATT-3′; PARP1 gRNA1, 5′-CCACCTCAACGTCAGGGTGC-3′; PARP1 gRNA2, 5′-TGGGTTCTCTGAGCTTCGGT-3′; SPRTN gRNA5, 5′-AAGGGGTTCGCTGAGACGGA-3′. Indel mutations were analyzed by Surveyor assays (Surveyor Mutation Detection Kit, Integrated DNA Technologies) and Sanger sequencing of PCR fragments cloned using TOPO TA Cloning (Thermo Fisher). Sequencing results for cell clones used in this study are summarized in Supplementary Fig. 1 and Supplementary Table 1. All HAP1 clones were diploid based on FACS analyses. DNA oligos for cloning and PCR amplification of target regions are summarized in Supplementary Table 3.
Clonogenic survival assays
Cells were seeded in triplicate in six-well plates, allowed to attach on plates for 4 h, and treated with various agents. For CPT (Sigma), niraparib (Merck), etoposide (Sigma), cisplatin (Teva Generics), and 5-aza-2′-deoxycytidine (Sigma), drug containing media were replenished every other day. For talazoparib (Selleckchem), media were not changed during treatment. For formaldehyde (Sigma), cells were treated for the first 24 h and the medium was changed. Cells were cultured for 6 days, fixed, and stained with Coomassie Blue. Colonies containing more than 50 cells were counted manually.
For detection of cell death, cells were stained first with Annexin V-APC (BD Bioscience, 550474) in binding buffer (10 mM HEPES-NaOH pH 7.4, 140 mM NaCl, 2.5 mM CaCl2) for 15 min at room temperature, and then with 100 µg ml−1 of PI in binding buffer. For analyses of DNA content, cells were fixed with 70% EtOH and stained with PI solution (50 µg ml−1 PI, 10 µg ml−1 RNase A, 0.05% NP-40). Two diploid clones of HAP1 (WT #2 and WT #3) were used as controls for cell-cycle analyses. Stained cells were analyzed by BD FACS Canto II (BD Biosiences). Percentages of cells in G2/M phase were estimated using ModFit LT (Verity Software House).
Immunofluorescence and microscopy
For γH2AX and RAD51 immunofluorescence, cells were fixed with 4% paraformaldehyde for 10 min, permeabilized with 0.2% Triton X-100 in PBS for 10 min, and blocked with 1% normal goat serum (for γH2AX) or 3% BSA (for RAD51) in PBS for 1 h. For TOP1cc staining, cells were fixed with 4% paraformaldehyde for 15 min at 4 °C, permeabilized with 0.25% Triton X-100 in PBS for 15 min at 4 °C and treated with 1% SDS in PBS for 5 min at room temperature. Cells were washed five times with wash buffer (0.1% Triton X-100, 0.1% BSA in PBS) and blocked with 10% milk in 10 mM Tris-HCl pH 7.4 and 150 mM NaCl. Primary antibodies used were mouse anti-phospho-Histone H2A.X (Ser139) (Millipore, 05-636, 1:500), rabbit anti-RAD51 (Calbiochem, PC130, 1:300), and mouse anti-TOP1cc antibody53 (1:100). Secondary antibodies used were goat anti-mouse IgG Alexa fluor 488 (Invitrogen, A11029, 1:2000) and goat anti-rabbit IgG Alexa fluor 488 (Invitrogen, A11034, 1:2000). Images were captured using a Zeiss Axio Scope.A1 fluorescent microscope equipped with a CCD camera (Jenoptik). Foci were scored using ImageJ on images captured with a 63× objective (for γH2AX) or counted manually (for RAD51 and TOP1cc). At least 100 cells were scored for focus formation by blinded observers.
DNA combing assay
Cells were labeled with 100 µM CldU (Sigma) for 30 min, washed three times with pre-warmed PBS, and labeled with 100 µM IdU (Sigma) for an additional 30 min. After labeling cells with IdU, cells were immediately washed three times with ice-cold PBS to inhibit DNA replication. For studying DNA replication under genotoxic stress, the IdU labeling was performed in the presence of a drug (3 µM Niraparib or 30 nM CPT). To measure collapsed forks, cells were treated with 5 mM HU for 30 min after CldU labeling (30 min) and released in IdU-containing media for 60 min. Where indicated, 5 µM ATR inhibitor (VE-821) was added during CldU labeling and HU treatment. DNA fibers were extracted in agarose plugs with FiberPrep DNA Extraction kit (Genomic Vision) and stretched onto silanized coverslips using the FiberComb Molecular Combing System (Genomic Vision). Combed DNA was dehydrated in an oven at 65 °C for 4 h and denatured with 0.5 M NaOH and 1 M NaCl for 8 min. Samples were then washed three times with PBS for 5 min each on a shaker, dehydrated sequentially in 70, 90, and 100% ethanol for 5 min each and dried at room temperature for 10 min. Samples were blocked with Block Aid (Invitrogen, B10710) at 37 °C for 30 min and incubated with rat anti-CldU antibody (Abcam, ab6326, 1:25) and anti-IdU antibody (BD Biosciences, 347580, 1:5) in Block Aid at 37 °C for 1 hr. After washing with PBS-T (0.05% Tween-20), samples were incubated with goat anti-rat IgG Alexa 488 (Invitrogen, A11006, 1:200) and goat anti-mouse IgG Alexa 594 (Invitrogen, A11032, 1:200) in Block Aid at 37 °C for 30 min. After washing with PBS-T, samples were mounted with ProLong Gold (Invitrogen). DNA fibers were photographed on a Zeiss Axio Scope.A1 fluorescent microscope and the length of CldU and IdU tracts were measured by blinded observers using ImageJ.
Purification of recombinant proteins
Recombinant protein production in insect cells was performed essentially as described before19 with modifications. pDEST20/FAM111A-strep, pFastBac NT-B/FAM111A-strep and pFastBac NT-B/Strep-FAM111A were transformed into the DH10Bac E. coli. Purified bacmids were transfected into Sf21 in using Cellfectin II (Thermo Fisher Scientific) as instructed by the manufacturer. After incubation at 28 °C for 3 days, virus-containing supernatants (P1 virus) were transferred to suspension culture in Sf-900 III SFM (Thermo Fisher Scientific) and viruses were amplified for 3 days at 28 °C (P2 virus). Viral titer was measured by baculoQUANT (Oxford Expression Technologies) as instructed. If necessary, the P2 virus was further amplified (P3 virus). For protein production, Sf21 cells in Sf-900 III SFM supplemented with glucose, yeastolate, and lactalbumin were infected with virus stocks at M.O.I of 10–20 and cultured for 3 days at 28 °C. These cells were collected, washed once with PBS, frozen in liquid N2, and stored at −80 °C until protein purification. For protein purification, cell pellets were thawed and resuspended in lysis buffer (50 mM HEPES-NaOH pH 7.5, 1 M NaCl, 1 mM MgCl2, 10% glycerol, 1% NP-40) supplemented with 4 U ml−1 Benzonase (Novagen). For GST-FAM111A-strep, 1 mM PMSF and cOmplete Mini EDTA-free Protease Inhibitor Cocktail (Roche) were added to the lysis buffer. For FAM111A-strep and Strep-FAM111A, 1 mM DTT, 1 mM EDTA and 2 µM Pepstatin A were added to the lysis buffer. After 30-min incubation on ice, samples were sonicated and cell debris was removed by centrifugation (20,000 × g, 4 °C, 30 min). Strep-Tactin superflow resin equilibrated with lysis buffer was mixed with lysates by rotation at 4 °C for 2–3 h. Resins were washed twice with lysis buffer and once with wash buffer (50 mM HEPES-NaOH pH 7.5, 250 mM NaCl, 10% glycerol). For FAM111A-strep and Strep-FAM111A, 1 mM DTT was included in the wash buffer. Proteins were eluted with elution buffer (wash buffer with 10 mM d-Desthiobiotin), frozen in liquid N2, and stored at −80 °C.
For protein production in E. coli, pMALII-c2x vectors were transformed into BL21 (DE3). For MBP-FAM111A WT 1–99, 100–175, 176–282 and 283–611, cells were cultured until OD600 = 0.5–1.2 and protein expression was induced with 0.1 mM IPTG at 18 °C overnight. Other MBP-FAM111A proteins were produced at the basal level expression without IPTG at 37 °C overnight. Cells were collected, washed with PBS, frozen in liquid N2, and stored at −80 °C until protein purification. For protein purification, frozen cells were thawed and resuspended in PBS with 1 mM PMSF and 250 µg ml−1 lysozyme. After 30 min incubation on ice, cells were sonicated and cell debris was removed by centrifugation (20,000 × g, 4 °C, 30 min). Amylose resin (New England Biolab) equilibrated with PBS was mixed with lysates at 4 °C for 2–3 h. Resins were washed with MBP wash buffer (50 mM Tris-HCl pH 8.0, 500 mM NaCl, 10% glycerol, 0.1% NP-40) three times and proteins were eluted with MBP elution buffer (50 mM Tris-HCl pH 8.0, 100 mM NaCl, 10% glycerol, 10 mM maltose). Purified proteins were frozen in liquid N2 and stored at −80 °C.
Electrophoretic mobility shift assay (EMSA)
EMSA was performed essentially as described before54 with modification. 5′ IRDye700-labeled 75-nt DNA oligo (O3063) was synthesized by IDT. Double-stranded (ds), Y-fork, and ds Y-fork DNA substrates were generated by annealing labeled O3063 with other non-labeled DNA oligos (ds: O4786; Y-fork: O4787; ds Y-fork: O4787 + O3730 + O3757) in annealing buffer (10 mM Tris-HCl pH 8.0, 50 mM NaCl, 1 mM EDTA). DNA oligo sequences are shown in Supplementary Table 3. DNA oligos were separated on 6% non-denaturing polyacrylamide gels and the properly annealed DNA was extracted in ddH2O overnight at room temperature. Remaining gels were removed and the buffer was exchanged to TE by Micro Bio-spin six column (Bio-Rad). For EMSA, serially diluted recombinant FAM111A proteins were mixed with 0.7 pmol of DNA substrates in binding buffer (final 25 mM Tris-HCl pH 7.5, 5 mM MgCl2, 10% glycerol, 20 ng µl−1 BSA, 1 mM DTT, total 10 µl) and incubated on ice for 1 h. For experiments using MBP-FAM111A 1–282, protein purity was assessed and protein amounts that contain roughly equal levels of full-length proteins were used. Samples were run on 6% non-denaturing polyacrylamide gel using 0.5 × TBE (4 °C, 100–120 V, 90–100 min) and imaged with Odyssey (LI-COR) or ChemiDoc MP (Bio-Rad). Images were analyzed using Image Studio Lite (LI-COR) or Image Lab (Bio-Rad) and percentages of band shifts were calculated based on the band intensities of free and shifted DNA substrates.
Fluorescence polarization DNA-binding assay
DNA binding was measured by fluorescence polarization assays essentially as described before20 with modifications. Serially diluted FAM111A protein (Strep-FAM111A or MBP-FAM111A 176–282) or control protein (BSA or MBP) was mixed with 5′ 6-FAM-labeled 31-nt DNA oligos (IDT) (final 10 nM) in FP buffer (10 mM HEPES-NaOH pH 7.5, 150 mM NaCl, total 30 µl). DNA oligo sequences are from previous study (O1676)54 (Supplementary Table 3). Samples were transferred to black 384-well plate and fluorescence polarization was measured on a Cytation 1 imager (BioTek). Kinetic constants were calculated by four-parameter logistic curve fitting in Prism 5 (GraphPad).
Recombinant FAM111A-strep R569H protein was produced in Sf21 cells and strep-tag containing protein fragments were captured by Strep-Tactin Super Flow Plus as described above. The sample was separated by SDS-PAGE and transferred to a PVDF membrane. Proteins were stained with Coomassie Brilliant Blue R-250 and the cleaved C-terminal fragment was sequenced by Edman degradation peptide sequencing (five cycles) at Alphalyse Laboratories.
Substrate docking analyses
Homology modeling of the FAM111A trypsin-like domain (residues 343–587) was performed using SWISS-MODEL55. The homology model was constructed using as a template the crystal structure of the plant protease Deg1 in active conformation (Protein Data Bank ID: 3QO6 chain A)42, possessing 23.7% amino acid identity to the modeled region of FAM111A. The substrate peptide sequences were initially positioned in the active site by backbone alignment with corresponding residues of a peptide substrate present in the Deg1 crystal structure.
In preparation for docking, the structural models were imported using Protein-Preparation-Wizard with Maestro 2019 version 19–3 (Schrödinger, LLC) with the OPLS3 force field. Bond orders were assigned, zero-order bonds to metals were determined, disulfide bonds were created, and all hydrogens were generated. Hydrogen bond assignments were applied based on water sampling, and protonation states were predicted for pH 7.4 ± 1.0 using PROPKA56,57. Steric clashes were resolved using a convergence RMSD of 0.3 Å using the OPLS3 force field under Polak–Ribiere Conjugate Gradient (PRCG) minimization58 for 5000 steps, or until the energy difference between subsequent structures was <0.001 kJ mol-Å−156.
For molecular docking of peptide substrate sequences, substrates were docked into the binding site using Glide extra precision (XP) peptide-docking protocols (Glide, 19–3, Schrödinger, LLC), which allowed for multiple molecular conformations to be sampled59,60,61,62. The starting conformations based on the PRCG minimized state were further energy minimized with a water-based solvent to generate charges at an extended cutoff (VdW at 8.0 Å, electrostatic at 15 Å, H-bond at 4.0 Å). Soft restraints were placed on all residues >12 Å from the modeled substrate by using harmonic restraints of 100 kcal mol−1, thus allowing the residues within the cutoff to freely refine position. In addition, the Monte Carlo Molecular Dynamics (MCMD) module with Maestro was employed for sampling side chain conformations and refining atomic positions of substrate-contacting residues within the FAM111A trypsin-like domain to optimize fitting with the substrate, as previously described62,63,64,65,66,67. Docking scores were used to rank the three sequences docked, namely, RTTFGKV, RTTRGKV, and RTTGGKV60,61,62.
The degree of amino acid conservation among species was analyzed by the ConSurf server68.
Cell fractionation was performed as described previously9. Briefly, trypsinized cells were washed with PBS and resuspended in hypotonic buffer (100 mM MES-NaOH pH 6.4, 1 mM EDTA, 0.5 mM MgCl2) supplemented with protease inhibitor cocktail. After mixing by gentle pipetting, samples were layered on hypotonic buffer containing 30% sucrose and centrifuged for 10 min at 15,000 × g at 4 °C. The pellet was isolated and proteins were extracted sequentially by resuspending the pellet in buffer A (50 mM HEPES-NaOH pH 7.5, 100 mM KCl, 2.5 mM MgCl2, 0.05% Triton X-100, protease inhibitor cocktail), buffer B (50 mM HEPES-NaOH pH 7.5, 250 mM KCl, 2.5 mM MgCl2, 0.05% Triton X-100, protease inhibitor cocktail) and buffer C (50 mM HEPES-NaOH pH 7.5, 500 mM KCl, 2.5 mM MgCl2, 0.1% Triton X-100, protease inhibitor cocktail). In each step, pellet was isolated by centrifugation (15,000 × g, 4 °C, 10 min). After these extractions, chromatin proteins were released by micrococcal nuclease (NEB) in buffer A supplemented with 5 mM CaCl2 at 37 °C for 15 min. Samples were centrifuged (15,000 × g, 4 °C, 10 min) and released proteins in supernatant were analyzed by Western blotting.
3xFlag-FAM111A (WT, S541A or F231A) were stably expressed in 293T cells by lentiviral vectors and iPOND experiments were performed essentially as described previously69. Briefly, cells were labeled with 10 µM EdU for 20 min, and chased with 10 µM thymidine for 30 min. Harvested cells were fixed with 1% formaldehyde for 20 min, permeabilized by 0.25% Triton X-100 for 30 min at room temperature followed by 90 min of click reaction, in which biotin is conjugated to EdU in the EdU-labeled DNA. Cells were lysed in RIPA lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.1% SDS, 0.5% Sodium deoxycholate) supplemented with protease inhibitor cocktail and PMSF and sonicated. Precipitates were removed by centrifugation (15,000 × g, 15 min, 4 °C) and EdU-labeled DNA-protein complexes were pulled down by streptavidin agarose beads (EMD Millipore) at 4 °C overnight. Beads were washed once with RIPA lysis buffer, once with 1 M NaCl, once with PBS, and twice with RIPA lysis buffer. Pulled down proteins were eluted in 2xLithium dodecyl sulfate (LDS) sample buffer at 70 °C for 1 h and analyzed by Western blotting.
Statistics and reproducibility
GraphPad Prism 5 was used to graph the data and statistical significance was determined by a two-tailed unpaired t-test. Experiments were repeated at least twice and similar results were obtained.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
The source data underlying Figs. 1b, d, 2a, c, e, f, 4b, c, f, 5b–e, g, i, 6b, e, and Supplementary Figs. 2b-d, f, 3f, i, k, l, 5c, 6c, and e are provided as a Source Data file. The uncropped blots underlying Figs. 1a, 2a, b, e, f, 3a–c, e, g–i, 4a, d, 5f, h, 6c, and Supplementary Figs. 2a, 3a–c, e, f, h–k, 4a, c, 5a, 6a, and d are provided in Supplementary Fig. 7. All data are available from the authors upon reasonable request.
Zeman, M. K. & Cimprich, K. A. Causes and consequences of replication stress. Nat. Cell Biol. 16, 2–9 (2014).
Cortez, D. Preventing replication fork collapse to maintain genome integrity. DNA Repair (Amst.) 32, 149–157 (2015).
Saldivar, J. C., Cortez, D. & Cimprich, K. A. The essential kinase ATR: ensuring faithful duplication of a challenging genome. Nat. Rev. Mol. Cell Biol. 18, 622–636 (2017).
Ide, H., Nakano, T., Shoulkamy, M. I. & Salem, A. M. H. Formation, repair, and biological effects of DNA–protein cross-link damage. In Advances in DNA Repair (ed. Chen, C.) 43–80 (IntechOpen, 2015).
Tretyakova, N. Y., Groehler, At & Ji, S. DNA-protein cross-links: formation, structural identities, and biological outcomes. Acc. Chem. Res. 48, 1631–1644 (2015).
Ide, H., Nakano, T., Salem, A. M. H. & Shoulkamy, M. I. DNA-protein cross-links: formidable challenges to maintaining genome integrity. DNA Repair (Amst.) 71, 190–197 (2018).
Pommier, Y. Topoisomerase I inhibitors: camptothecins and beyond. Nat. Rev. Cancer 6, 789–802 (2006).
Satoh, M. S. & Lindahl, T. Role of poly(ADP-ribose) formation in DNA repair. Nature 356, 356–358 (1992).
Murai, J. et al. Trapping of PARP1 and PARP2 by Clinical PARP Inhibitors. Cancer Res. 72, 5588–5599 (2012).
Murai, J. et al. Stereospecific PARP trapping by BMN 673 and comparison with olaparib and rucaparib. Mol. Cancer Ther. 13, 433–443 (2014).
Shen, Y., Aoyagi-Scharber, M. & Wang, B. Trapping Poly(ADP-Ribose) polymerase. J. Pharmacol. Exp. Ther. 353, 446–457 (2015).
Pommier, Y., O’Connor, M. J. & de Bono, J. Laying a trap to kill cancer cells: PARP inhibitors and their mechanisms of action. Sci. Transl. Med. 8, 362ps17 (2016).
Murai, J. & Pommier, Y. PARP trapping beyond homologous recombination and platinum sensitivity in cancers. Annu. Rev. Cancer Biol. 3, 131–150 (2019).
Bryant, H. E. et al. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature 434, 913–917 (2005).
Farmer, H. et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 434, 917–921 (2005).
Stingele, J., Schwarz, M. S., Bloemeke, N., Wolf, P. G. & Jentsch, S. A DNA-dependent protease involved in DNA-protein crosslink repair. Cell 158, 327–338 (2014).
Stingele, J. & Jentsch, S. DNA-protein crosslink repair. Nat. Rev. Mol. Cell Biol. 16, 455–460 (2015).
Stingele, J., Habermann, B. & Jentsch, S. DNA-protein crosslink repair: proteases as DNA repair enzymes. Trends Biochem. Sci. 40, 67–71 (2015).
Stingele, J. et al. Mechanism and regulation of DNA-protein crosslink repair by the DNA-dependent metalloprotease SPRTN. Mol. Cell 64, 688–703 (2016).
Vaz, B. et al. Metalloprotease SPRTN/DVC1 orchestrates replication-coupled DNA-protein crosslink repair. Mol. Cell 64, 704–719 (2016).
Lopez-Mosqueda, J. et al. SPRTN is a mammalian DNA-binding metalloprotease that resolves DNA-protein crosslinks. Elife 5, e21491 (2016).
Stingele, J., Bellelli, R. & Boulton, S. J. Mechanisms of DNA-protein crosslink repair. Nat. Rev. Mol. Cell Biol. 18, 563–573 (2017).
Vaz, B., Popovic, M. & Ramadan, K. DNA-protein crosslink proteolysis repair. Trends Biochem. Sci. 42, 483–495 (2017).
Morocz, M. et al. DNA-dependent protease activity of human Spartan facilitates replication of DNA-protein crosslink-containing DNA. Nucleic Acids Res. 45, 3172–3188 (2017).
Fielden, J., Ruggiano, A., Popovic, M. & Ramadan, K. DNA protein crosslink proteolysis repair: from yeast to premature ageing and cancer in humans. DNA Repair (Amst.) 71, 198–204 (2018).
Li, F., Raczynska, J. E., Chen, Z. & Yu, H. Structural insight into DNA-dependent activation of human metalloprotease Spartan. Cell Rep. 26, 3336–3346 e4 (2019).
Maskey, R. S. et al. Spartan deficiency causes accumulation of Topoisomerase 1 cleavage complexes and tumorigenesis. Nucleic Acids Res. 45, 4564–4576 (2017).
Duxin, J. P., Dewar, J. M., Yardimci, H. & Walter, J. C. Repair of a DNA-protein crosslink by replication-coupled proteolysis. Cell 159, 346–357 (2014).
Larsen, N. B. et al. Replication-coupled DNA-protein crosslink repair by SPRTN and the proteasome in xenopus egg extracts. Mol. Cell 73, 574–588 e7 (2019).
Ruijs, M. W. et al. Atypical progeroid syndrome: an unknown helicase gene defect? Am. J. Med. Genet. A 116A, 295–299 (2003).
Lessel, D. et al. Mutations in SPRTN cause early onset hepatocellular carcinoma, genomic instability and progeroid features. Nat. Genet. 46, 1239–1244 (2014).
Maskey, R. S. et al. Spartan deficiency causes genomic instability and progeroid phenotypes. Nat. Commun. 5, 5744 (2014).
Borgermann, N. et al. SUMOylation promotes protective responses to DNA-protein crosslinks. EMBO J. 38, e101496 (2019).
Alabert, C. et al. Nascent chromatin capture proteomics determines chromatin dynamics during DNA replication and identifies unknown fork components. Nat. Cell Biol. 16, 281–293 (2014).
Bochkarev, A., Pfuetzner, R. A., Edwards, A. M. & Frappier, L. Structure of the single-stranded-DNA-binding domain of replication protein A bound to DNA. Nature 385, 176–181 (1997).
Hedstrom, L. Serine protease mechanism and specificity. Chem. Rev. 102, 4501–4524 (2002).
Rawlings, N. D. et al. The MEROPS database of proteolytic enzymes, their substrates and inhibitors in 2017 and a comparison with peptidases in the PANTHER database. Nucleic Acids Res. 46, D624–D632 (2018).
Unger, S. et al. FAM111A mutations result in hypoparathyroidism and impaired skeletal development. Am. J. Hum. Genet. 92, 990–995 (2013).
Isojima, T. et al. A recurrent de novo FAM111A mutation causes Kenny-Caffey syndrome type 2. J. Bone Miner. Res. 29, 992–998 (2014).
Nikkel, S. M. et al. Mother-to-daughter transmission of Kenny-Caffey syndrome associated with the recurrent, dominant FAM111A mutation p.Arg569His. Clin. Genet. 86, 394–395 (2014).
Abraham, M. B. et al. Short stature and hypoparathyroidism in a child with Kenny-Caffey syndrome type 2 due to a novel mutation in FAM111A gene. Int. J. Pediatr. Endocrinol. 2017, 1 (2017).
Kley, J. et al. Structural adaptation of the plant protease Deg1 to repair photosystem II during light exposure. Nat. Struct. Mol. Biol. 18, 728–731 (2011).
Prasad, R. et al. Suicidal cross-linking of PARP-1 to AP site intermediates in cells undergoing base excision repair. Nucleic Acids Res. 42, 6337–6351 (2014).
Prasad, R., Horton, J. K., Dai, D. P. & Wilson, S. H. Repair pathway for PARP-1 DNA-protein crosslinks. DNA Repair (Amst.) 73, 71–77 (2019).
Sparks, J. L. et al. The CMG helicase bypasses DNA-protein cross-links to facilitate their repair. Cell 176, 167–181 e21 (2019).
Clausen, T., Southan, C. & Ehrmann, M. The HtrA family of proteases: implications for protein composition and cell fate. Mol. Cell 10, 443–455 (2002).
Clausen, T., Kaiser, M., Huber, R. & Ehrmann, M. HTRA proteases: regulated proteolysis in protein quality control. Nat. Rev. Mol. Cell Biol. 12, 152–162 (2011).
Krojer, T. et al. Structural basis for the regulated protease and chaperone function of DegP. Nature 453, 885–890 (2008).
Fine, D. A. et al. Identification of FAM111A as an SV40 host range restriction and adenovirus helper factor. PLoS Pathog. 8, e1002949 (2012).
Panda, D., Fernandez, D. J., Lal, M., Buehler, E. & Moss, B. Triad of human cellular proteins, IRF2, FAM111A, and RFC3, restrict replication of orthopoxvirus SPI-1 host-range mutants. Proc. Natl Acad. Sci. USA 114, 3720–3725 (2017).
Tarnita, R. M., Wilkie, A. R. & DeCaprio, J. A. Contribution of DNA replication to the FAM111A-mediated simian virus 40 host range phenotype. J. Virol. 93, e01330 (2019).
Machida, Y., Kim, M. S. & Machida, Y. J. Spartan/C1orf124 is important to prevent UV-induced mutagenesis. Cell Cycle 11, 3395–3402 (2012).
Patel, A. G. et al. Immunodetection of human topoisomerase I-DNA covalent complexes. Nucleic Acids Res. 44, 2816–2826 (2016).
Toth, A., Hegedus, L., Juhasz, S., Haracska, L. & Burkovics, P. The DNA-binding box of human SPARTAN contributes to the targeting of Poleta to DNA damage sites. DNA Repair (Amst.) 49, 33–42 (2017).
Waterhouse, A. et al. SWISS-MODEL: homology modelling of protein structures and complexes. Nucleic Acids Res. 46, W296–W303 (2018).
Sondergaard, C. R., Olsson, M. H., Rostkowski, M. & Jensen, J. H. Improved treatment of ligands and coupling effects in empirical calculation and rationalization of pKa values. J. Chem. Theory Comput. 7, 2284–2295 (2011).
Li, H., Robertson, A. D. & Jensen, J. H. Very fast empirical prediction and rationalization of protein pKa values. Proteins 61, 704–721 (2005).
Polak, E. & Ribiere, G. Note sur la convergence de méthodes de directions conjuguées. Esaim Math. Model Numer Anal 3, 35–43 (1969).
Caulfield, T. & Devkota, B. Motion of transfer RNA from the A/T state into the A-site using docking and simulations. Proteins 80, 2489–2500 (2012).
Loving, K., Salam, N. K. & Sherman, W. Energetic analysis of fragment docking and application to structure-based pharmacophore hypothesis generation. J. Comput. Aided Mol. Des. 23, 541–554 (2009).
Salam, N. K., Nuti, R. & Sherman, W. Novel method for generating structure-based pharmacophores using energetic analysis. J. Chem. Inf. Model. 49, 2356–2368 (2009).
Caulfield, T. & Medina-Franco, J. L. Molecular dynamics simulations of human DNA methyltransferase 3B with selective inhibitor nanaomycin A. J. Struct. Biol. 176, 185–191 (2011).
Caulfield, T. R. et al. Phosphorylation by PINK1 releases the UBL domain and initializes the conformational opening of the E3 ubiquitin ligase Parkin. PLoS Comput. Biol. 10, e1003935 (2014).
Abdul-Hay, S. O. et al. Selective targeting of extracellular insulin-degrading enzyme by quasi-irreversible thiol-modifying inhibitors. ACS Chem. Biol. 10, 2716–2724 (2015).
Batra, J. et al. Long-range electrostatic complementarity governs substrate recognition by human chymotrypsin C, a key regulator of digestive enzyme activation. J. Biol. Chem. 288, 9848–9859 (2013).
Lopez-Vallejo, F. et al. Integrating virtual screening and combinatorial chemistry for accelerated drug discovery. Comb. Chem. High. Throughput Screen. 14, 475–487 (2011).
Paulus, A. et al. Coinhibition of the deubiquitinating enzymes, USP14 and UCHL5, with VLX1570 is lethal to ibrutinib- or bortezomib-resistant Waldenstrom macroglobulinemia tumor cells. Blood Cancer J. 6, e492 (2016).
Ashkenazy, H. et al. ConSurf 2016: an improved methodology to estimate and visualize evolutionary conservation in macromolecules. Nucleic Acids Res. 44, W344–W350 (2016).
Sirbu, B. M. et al. Analysis of protein dynamics at active, stalled, and collapsed replication forks. Genes Dev. 25, 1320–1327 (2011).
We thank Larry M. Karnitz for critical reading of the manuscript, Kevin L. Peterson for technical assistance, and Karen S. Flatten for technical advice. This work was supported by the National Institutes of Health (R01 CA233700 to Y.J.M.) and Eagles Cancer Research Fund to Y.J.M.
S.H. Kaufmann is an inventor on U.S. patent No. 8,530,172 held by Mayo Clinic that deals with use of antibodies to protein–DNA complexes as diagnostic reagents to predict responsiveness to topoisomerase inhibitors. The remaining authors declare no competing interests.
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Kojima, Y., Machida, Y., Palani, S. et al. FAM111A protects replication forks from protein obstacles via its trypsin-like domain. Nat Commun 11, 1318 (2020). https://doi.org/10.1038/s41467-020-15170-7
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