Report

ZATT (ZNF451)–mediated resolution of topoisomerase 2 DNA-protein cross-links

See allHide authors and affiliations

Science  29 Sep 2017:
Vol. 357, Issue 6358, pp. 1412-1416
DOI: 10.1126/science.aam6468

Resolving a DNA-protein cross-link

Topoisomerase 2 (TOP2) creates DNA double-strand breaks to regulate DNA topology and is critical for processes such as replication and transcription. A covalent complex between TOP2 and DNA (TOP2cc) is an intermediate in the reaction that can be trapped by drugs. Schellenberg et al. show how the SUMO ligase ZATT promotes the resolution of TOP2cc by means of tyrosyl-DNA phosphoesterase 2 (TDP2), both by enhancing recruitment of TDP2 to SUMOylated TOP2 and by enhancing the hydrolase activity of TDP2.

Science, this issue p. 1412

Abstract

Topoisomerase 2 (TOP2) DNA transactions proceed via formation of the TOP2 cleavage complex (TOP2cc), a covalent enzyme-DNA reaction intermediate that is vulnerable to trapping by potent anticancer TOP2 drugs. How genotoxic TOP2 DNA-protein cross-links are resolved is unclear. We found that the SUMO (small ubiquitin-related modifier) ligase ZATT (ZNF451) is a multifunctional DNA repair factor that controls cellular responses to TOP2 damage. ZATT binding to TOP2cc facilitates a proteasome-independent tyrosyl-DNA phosphodiesterase 2 (TDP2) hydrolase activity on stalled TOP2cc. The ZATT SUMO ligase activity further promotes TDP2 interactions with SUMOylated TOP2, regulating efficient TDP2 recruitment through a “split-SIM” SUMO2 engagement platform. These findings uncover a ZATT-TDP2–catalyzed and SUMO2-modulated pathway for direct resolution of TOP2cc.

Topoisomerase 2 (TOP2α and TOP2β in mammalian cells) regulates DNA topology through the production of transient DNA double-strand breaks (DSBs) (1). The key intermediate in the TOP2 reaction is the covalent protein-DNA TOP2 cleavage complex (TOP2cc), a protein-DNA cross-link that forms between the active-site TOP2 tyrosine and the 5′ terminus of the incised DNA duplex (2) (Fig. 1A). TOP2 is critical for facilitating DNA processes such as replication and transcription (35). However, it can be trapped on DNA by poisons, including frontline anticancer drugs such as etoposide, or by binding to existing DNA damage. Poisoning results in stable protein-blocked DNA breaks that impede elongation of RNA and DNA polymerases and cause cell death (14, 6, 7).

Fig. 1 TDP2 binds SUMOylated TOP2 and ZNF451.

(A) Cellular mechanisms that regulate TDP2-catalyzed phosphotyrosyl bond hydrolysis in poisoned TOP2cc are unknown. (B) Immunoblotting (IB) of soluble cell lysates and IPs from cells expressing YFP, YFP-TDP2, or YFP-TDP2H351N (H-N). (C) Resolution of γH2AX foci in MEFs after etoposide exposure (20 μM). Data are means ± SEM (N = 3); **P < 0.01, ***P < 0.001 [two-way analysis of variance (ANOVA) with Bonferroni posttest]. (D) Silver-stained SDS–polyacrylamide gel electrophoresis (PAGE) of YFP-TDP2 (Y-T)–associated SUMO2-modified proteins isolated as in fig. S2A. (E) Left: Immunoblotting of ZNF451 in whole-cell extracts or chromatin fraction. Right: Quantification of ZNF451 levels in chromatin. Data are means ± SEM (N = 3); *P < 0.05 (two-way ANOVA with Bonferroni posttest).

Vertebrate tyrosyl-DNA phosphodiesterase 2 (TDP2, also known as VpG unlinkase, TTRAP, or EAPII) directly resolves the protein-DNA linkages (5′-phosphotyrosyl) characteristic of TOP2-induced DSBs (711). In this context, TDP2 modulates cellular (7, 10) and organismal (12) survival after TOP2-targeting anticancer drug treatments, and TDP2 inhibitors hold promise for chemotherapy (13, 14). A critical question in TOP2 biology is how TDP2 accesses the TOP2-DNA phosphotyrosyl chemical bond, which is protected within the TOP2 protein shell (15, 16) (Fig. 1A). Because etoposide treatment can trigger TOP2 degradation by the proteasome (1719), it is hypothesized that TDP2 processes TOP2-DNA oligopeptides after TOP2cc proteolytic degradation (20). However, the molecular basis for regulation, coordination, and control of TOP2cc metabolism remains enigmatic.

To identify modulators of TDP2-dependent TOP2cc repair, we stably expressed yellow fluorescent protein (YFP)–tagged TDP2 in human embryonic kidney (HEK) 293F cells, purified TDP2-containing protein complexes by means of anti-GFP/YFP single-domain camelid nanobody (sdAb) resin, and identified copurifying proteins by liquid chromatography–tandem mass spectrometry (fig. S1, A to C). We observed robust enrichment of TOP2α, TOP2β, and SUMO2 (small ubiquitin-like modifier 2), but not SUMO1 peptides, in YFP-TDP2 immunoprecipitates (IP) (tables S1 and S2). Western blotting revealed that TDP2 interacts with a ladder of intact (nonproteolyzed) TOP2α and TOP2β (Fig. 1B, lane 5), which is posttranslationally modified with SUMO2 (fig. S1D, lane 4).

TOP2 is conjugated with SUMO2 during mitosis or in response to TOP2 poisons (2123), and we found that etoposide treatment prior to TDP2 IP increased the amount of high–molecular weight SUMO2 and the extent of modification of TOP2α and TOP2β (fig. S1E, lanes 8 to 12). Intriguingly, IP conducted with a catalytically inactive variant of TDP2H351N (8) was nearly devoid of SUMO2-TOP2 (Fig. 1B, lane 6), which suggests that TDP2 catalysis is required to liberate intact SUMOylated TOP2 from the insoluble chromatin fraction. This prompted us to evaluate the untested link between the proteasome and TDP2 in repair of poisoned TOP2cc by monitoring the resolution of phosphohistone H2AX (γH2AX) foci after etoposide treatment. As reported previously, Tdp2 knockout (Tdp2−/−) mouse embryonic fibroblasts (MEFs) display delayed resolution of etoposide-induced γH2AX foci relative to Tdp2+/+cells (11), and we found that this was severely impaired by proteasome inhibition with MG132 (Fig. 1C and fig. S1F). These data are consistent with a TDP2 SUMO2-dependent TOP2cc resolution mechanism that acts independently of, or parallel to, proteasome-mediated TOP2cc repair.

To identify factors that regulate SUMO2- and TDP2-dependent TOP2cc repair, we conducted tandem affinity purification (TAP) from HEK293F cells expressing YFP-TDP2 and His6-tagged SUMO2 (Fig. 1D and fig. S2A). TAP samples contained a ladder of SUMO2-modified proteins (Fig. 1D, lane 5). Ubiquitin-like specific protease 1 (ULP1) treatment of this ladder uncovered four proteins (Fig. 1D, lane 8): TOP2α, TOP2β, SUMO2, and ZNF451 (a prototypical member of a recently identified class of SUMO2 E3/E4 ligases) (fig. S2B) (24, 25). Similar to TDP2, IP samples of endogenous ZNF451 or GFP-ZNF451 were enriched with SUMO2, TOP2α, and TOP2β (fig. S2, C and D, and tables S1 and S3), which suggests that these proteins form a functional complex in cells. Furthermore, recombinant ZNF451 bound to TOP2α and TOP2β (fig. S2E). ZNF451 was recruited to the cellular chromatin fraction after etoposide treatment when the proteasome was inhibited (Fig. 1E), but not by ionizing radiation–induced DSBs (fig. S2F). These data indicate ZNF451 directly binds to TOP2 and is recruited to chromatin after TOP2 poisoning.

TDP2 is unable to hydrolyze intact, recombinant Saccharomyces cerevisiae TOP2 DNA-protein cross-links (ScTOP2cc) in vitro, but the activity is enabled by heat denaturation of ScTOP2cc (20), which suggests that TOP2cc resolution could be regulated by proteasome-independent mechanisms. Given the direct binding of ZNF451 to TOP2 and TDP2 (figs. S2E and S3A), we hypothesized that ZNF451 could regulate the activity of TDP2 on TOP2cc. To test this, we generated reconstituted TOP2cc via reaction of TOP2α or TOP2β with a suicide oligonucleotide substrate (20) (Fig. 2A) and assayed the result for TDP2-dependent TOP2cc resolution (Fig. 2B and fig. S3, B to D). We found that mammalian TOP2cc is generally refractory to direct resolution by TDP2 except at high concentrations of TDP2, where we observed liberation of a small amount of 15-nucleotide DNA product diagnostic of TDP2-catalyzed hydrolysis of TOP2cc (Fig. 2B, lanes 7 and 8, and fig. S3B). Strikingly, TDP2 was more active on TOP2cc (α and β) in the presence of ZNF451 by more than three orders of magnitude, with nanomolar concentrations of ZNF451 and TDP2 being sufficient to catalyze TOP2cc hydrolysis (Fig. 2B and fig. S3, B to E). Neither ZNF451 alone nor a catalytically impaired TDP2H351N mutant supported this reaction (fig. S3F, lanes 4 and 6), indicating that ZNF451 stimulates TOP2cc hydrolysis catalyzed by TDP2. Probing of the TOP2cc structure suggests that ZNF451 alters the conformation of TOP2cc to facilitate the TDP2 direct-resolution reaction (fig. S3G).

Fig. 2 ZNF451 promotes phosphotyrosyl bond hydrolysis by TDP2.

(A) Synthesis and purification of stalled TOP2cc. (B) TOP2βcc (0.2 nM) hydrolysis by TDP2 is enhanced by ZNF451. A representative gel from three experiments is shown. (C) Cellular proliferation of HEK293F or CRISPR knockout cells measured by area under the curve (AUC) of cell confluency after 6 days of growth with etoposide. IC50, 50% confluence from a four-parameter fit of log[etoposide] versus AUC; data are means ± SD (N = 4). (D) Clonogenic survival of MEFs in the indicated concentrations of etoposide; nt, nontargeting. Data are means ± SEM (N ≥ 4); *P ≤ 0.05 (F test, linear quadratic model). (E) Resolution of DSBs marked by γH2AX foci in MEFs after etoposide exposure in the presence or absence of MG132. Data are means ± SEM (N = 3); *P < 0.05, **P < 0.01 (two-way ANOVA with Bonferroni posttest); ns, not significant.

To assess the contribution of ZNF451 to TOP2cc repair in mammalian cells, we examined the cellular sensitivity of ΔZNF451, ΔTDP2, and ΔTDP2/ΔZNF451 double knockout HEK293F cell lines to etoposide (Fig. 2C and fig. S4, A and B). ZNF451 deletion conferred an even more severe etoposide sensitivity than did TDP2 deletion (Fig. 2C). Similarly, short hairpin RNA (shRNA)–mediated ZNF451 knockdown sensitized HEK293F cells to etoposide, whereas GFP-ZNF451 overexpression decreased etoposide sensitivity, indicating that ZNF451 expression directly correlates with etoposide resistance in a dose-dependent manner (fig. S4, C and D). This effect was specific to TOP2 drugs, as ZNF451 knockdown did not affect sensitivity to camptothecin, methyl methanesulfonate, or phleomycin B1 (Zeocin) (fig. S4D). TDP2 deletion further increased etoposide sensitivity in ΔZNF451, but to a lesser degree than in wild-type cells, suggesting both collaborative and unique functions of these proteins in the response to TOP2 damage.

Knockdown of the murine ZNF451 homolog (ZFP451) also decreased cell survival after etoposide treatment in both wild-type and Tdp2−/−-transformed MEFs (Fig. 2D and fig. S4E). Although ZFP451 depletion alone did not confer a significant defect in γH2AX foci resolution, it caused a delay in the repair kinetics when combined with Tdp2 deletion (Tdp2−/−) or proteasome inhibition (Fig. 2E). In line with protein-protein interaction results, ZFP451 and TDP2 appear to act in the same proteasome-independent TOP2cc repair pathway, as ZFP451 depletion in Tdp2−/− cells did not further impair resolution of etoposide-induced γH2AX foci in MG132-treated cells. These effects were not the result of global impairment of DSB repair by proteasome inhibition, as MG132 did not cause major defects in γH2AX foci resolution after ionizing radiation treatment (fig. S4F). Overall, our results identify ZNF451 as a component in the cellular response to TOP2-induced damage that operates through both TDP2-dependent and TDP2-independent mechanisms.

ZNF451 is a SUMO2 E3/E4 ligase (24, 25), so we examined TOP2 SUMOylation in vitro using reactions containing SUMO E1 (Sae2/Aos2), E2 (Ubc9), SUMO2, and full-length ZNF451. ZNF451 exhibited robust autoSUMOylation and catalyzed polySUMOylation of recombinant TOP2α (Fig. 3A, lanes 4 to 8). ZNF451 further displayed a marked preference for SUMOylating TOP2cc over TOP2 (Fig. 3B and fig. S5, A to C), which suggests that TOP2cc is a preferred and specific target for ZNF451 SUMO2 ligase activity. In HEK293F cells, steady-state levels of TOP2α (Fig. 3, C and D) and TOP2β (fig. S5, D and E) SUMOylation with SUMO2 were largely dependent on ZNF451 (Fig. 3C, lanes 7 and 8), as was etoposide-induced stimulation of TOP2 SUMOylation (Fig. 3C, lanes 9 and 10). Intriguingly, ZNF451-dependent TOP2 SUMOylation was also triggered by treatment with ICRF-193, a drug that induces TOP2 clamping on DNA (Fig. 3C, lanes 11 and 12). These results indicate that ZNF451 regulates TOP2 SUMOylation after treatment with drugs that perturb the TOP2 reaction cycle by distinct mechanisms.

Fig. 3 ZNF451 preferentially SUMOylates TOP2cc and enhances TDP2 activity.

(A) Recombinant TOP2α was incubated with SUMO E1, E2, and increasing concentrations of SUMO2 in the presence or absence of ZNF451. (B) Reactions with 2 nM TOP2αcc (DNA-Cy5 label) and 75 nM free TOP2α were incubated with SUMO E1, E2, SUMO2, and the indicated concentration of ZNF451, then separated by SDS-PAGE. TOP2cc SUMOylation was monitored by scanning for the Cy5 label; free TOP2β SUMOylation was monitored by Western blotting. (C) Immunoblot of lysates and Ni-NTA–purified samples from cells expressing His6-SUMO2 pretreated with the indicated drugs for 20 min. (D) Quantification of SUMO2-TOP2α signal from lanes 7 to 12 in (C). Data are means ± SD (N = 3); *P < 0.05, **P < 0.001 (two-tailed t test). (E) Representative image (top) and quantification (bottom) of TOP2β or SUMO2 covalently bound to genomic DNA in MEFs after 1 hour of etoposide treatment and recovery in the presence or absence of 20 μM MG132. Data are means ± SEM (N ≥ 7); *P < 0.05 (two-way ANOVA with Bonferroni posttest).

TOP2 SUMOylation by ZNF451 also increased the efficiency of ZNF451-TDP2 TOP2cc hydrolysis by an additional ~75% (fig. S5F). Thus, we tested the role of SUMOylation in TDP2-catalyzed removal of TOP2cc with an ICE (in vivo complexes of enzyme) assay that monitors TOP2β and SUMO2-modified TOP2β DNA-protein cross-link removal from chromatin (11) (Fig. 3E, supplementary text, and fig. S6). After TOP2 poisoning and etoposide removal, the turnover of the SUMO2-modified TOP2β fraction was specifically delayed in Tdp2−/− cells, but only when the proteasome was inhibited (Fig. 3E and fig. S6E). In vitro treatment of TOP2cc from ICE samples with recombinant wild-type TDP2 also showed enhanced removal of the SUMOylated TOP2βcc fraction relative to total TOP2β (fig. S6F), consistent with a model where TOP2cc SUMOylation promotes TDP2-catalyzed TOP2cc direct resolution.

SUMOylation enhances protein-protein associations in the DNA damage response (26, 27), and TDP2 binds SUMO2 (but not SUMO1) (28). Thus, covalent labeling of TOP2cc with SUMO2 may recruit TDP2 to poisoned TOP2cc. Maltose binding protein pulldowns map the SUMO2-binding region of TDP2 to the catalytic domain (amino acids 108 to 362, TDP2cat) (fig. S7A), which lacks a canonical SUMO interaction motif (SIM) (28). To define the molecular basis for this noncanonical SUMO interaction, we crystallized and determined x-ray structures of mouse TDP2cat as binary mTDP2cat-SUMO2 and ternary mTDP2cat-SUMO2-DNA complexes (Fig. 4A, fig. S7B, and table S4). SUMO2 binds distal to the TDP2 catalytic center through five SUMO-binding elements, SB1 to SB5 (Fig. 4, A and B). Using small-angle x-ray scattering, we confirmed that this overall architecture of mTDP2cat-SUMO2 is consistent with that in solution (fig. S7, C to H). The core of the SUMO2-TDP2 interface is composed of a “split-SIM” structure, distinct from well-characterized SUMO-SIM interfaces. In a triangular configuration, two β strands (β0 of SB1; C-terminal β14 from SB5) engage the SUMO2 β sheet (Fig. 4B). A prominent feature of this interface is the insertion of the TDP2 C-terminal Leu370 into a SUMO2 hydrophobic pocket, where the terminal carboxylate of Leu370 forms a salt bridge with SUMO2 Lys42 (Fig. 4, B and C). Loops SB2, SB3, and SB4 augment this split-SIM core, which comprises an interface (~800 Å2) that is larger than typical SUMO-SIM interfaces (500 to 600 Å2) (fig. S7, I and J). Accordingly, the dissociation constant (Kd) for the TDP2-SUMO2 interaction (fig. S7, K and L) is 880 nM, placing TDP2 among the stronger SUMO-binding proteins (28).

Fig. 4 Structure of SUMO2-mTDP2cat reveals molecular basis for recruitment of TDP2 by SUMOylation.

(A) Structure of the DNA-mTDP2cat-SUMO2 complex. SUMO2 binds the TDP2 catalytic domain with a split SIM (red) and three loops (turquoise) distal from the DNA binding site. (B) A “β triangle” is formed by strands β0 and β14 of mTDP2cat and β2 of SUMO2. Amino acid abbreviations: D, Asp; E, Glu; K, Lys; L, Leu; Q, Gln; R, Arg; S, Ser; T, Thr; V, Val. (C) The C terminus of TDP2 fits in a pocket on SUMO2. (D) Immunoblotting of IPs from cells coexpressing YFP, YFP-TDP2, or YFP-TDP2 C-AE with shRNAs. (E) HEK293F cells expressing YFP-TDP2 (wild-type or C-AE mutant) and a nontargeting (nt) or ZNF451-targeting shRNA control were microirradiated with a UV laser between the two white arrows at t = 0. Scale bar, 10 μm.

To probe functions of the TDP2-SUMO2 interaction, we mutated β0 to encode proline substitutions (TDP2PQ), and/or extended the buried C-terminal Leu370 by two residues (TDP2C-AE) to create steric blocks to SUMO2 engagement (Fig. 4C and fig. S7M). In HEK293 cells, YFP-TDP2 C-AE, PQ, or C-AE/PQ double mutants failed to colocalize with SUMO2 (fig. S8A). In vitro, TDP2C-AE blocked SUMO2 binding in size-exclusion chromatography and when monitored by nuclear magnetic resonance spectroscopy yet did not impair TDP2 phosphotyrosylase activity (fig. S8, B to E). YFP-TDP2 association with SUMO2-TOP2 was impaired by the C-AE SUMO-binding mutation, by ZNF451 knockdown, or by a combination of these defects; hence, the TDP2-SUMO2 interface is important for engagement of SUMOylated TOP2 (Fig. 4D).

In comparison to strong phenotypes observed with ZNF451 and TDP2 depletion (Fig. 2, C and D), we found that overexpressed TDP2C-AE complements Tdp2−/− MEFs in clonogenic survival and γH2AX repair assays after etoposide treatment (fig. S8, F and G). We reasoned that SUMOylation of TOP2 by ZNF451 may thus act secondarily to its enhancement of TOP2cc hydrolase activity, with SUMOylation acting to direct TDP2 to stalled cleavage complexes. We thus examined the kinetics of TDP2 recruitment to TOP2 DNA damage generated by ultraviolet (UV) microirradiation (10, 29, 30). YFP-TDP2 accumulated within 150 s after UV treatment (fig. S8H), and both ZNF451 knockdown and the C-AE mutation impaired TDP2 mobilization to DNA damage (Fig. 4E and fig. S8I). TDP2C-AE mutation also delayed, but did not ablate, removal of the SUMOylated fraction of TOP2 from ICE-purified TOP2cc (fig. S8J). Together, these results indicate that the TDP2 interaction with TOP2cc is enhanced by interactions between TDP2 and SUMO2 and is dependent on ZNF451 SUMO2 ligase. Furthermore, TDP2 directly binds TOP2 and ZNF451 in vitro (figs. S3A and S7A), so this array of interactions with SUMO2, TOP2, and ZNF451 likely facilitates recruitment of TDP2 to SUMO2-TOP2cc in cells.

The ZNF451-TDP2–modulated TOP2cc direct resolution pathway (fig. S9) may contribute to tumor adaptation during chemotherapy with TOP2 poisons, and thus it constitutes a new potential target for chemotherapeutic intervention. In addition to TDP2-related functions, ZNF451 displays TDP2-independent effects on the cellular response to TOP2 poisoning that will require further investigation. It will be important to elucidate the mechanics of ZNF451-TDP2’s influence on TOP2 and its potential for modulating TOP2 regulation of genome dynamics and transcription (4, 12, 31, 32). Given ZNF451’s association with TDP2-mediated TOP2 repair, we propose renaming it ZATT (i.e., zinc finger protein associated with TDP2 and TOP2) to appropriately reflect these cellular functions.

Supplementary Materials

www.sciencemag.org/content/357/6358/1412/suppl/DC1

Materials and Methods

Supplementary Text

Figs. S1 to S9

Tables S1 to S6

References (3354)

References and Notes

Acknowledgments: We thank NIEHS staff and core facilities: A. Moon, J. Krahn, and L. Pedersen (x-ray crystallography), C. Malone and B. Petrovich (anti-GFP sepharose and HEK293F cell culture), A. Janoshazi and J. Tucker (microscopy), A. Adams and K. Johnson (mass spectrometry), M. Sifre and C. Bortner (flow cytometry), and N. Gassman (UV microirradiation). Supported by NIH Intramural Research Program grants 1Z01ES102765 (R.S.W.), 1ZIAES050111-26 (R.E.L.), and ZES102488-09 (J.G.W.); Spanish and Andalusian Government grants SAF2010-21017, SAF2013-47343-P, SAF2014-55532-R, CVI-7948, and FEDER funds (F.C.-L.) and BES-2015-071672 (A.H.-R.); European Research Council grant ERC-CoG-2014-647359 (F.C.-L.); and University of Seville grant PIF-2011 (J.A.L.). The Advanced Photon Source SERCAT beamline is supported by the U.S. Department of Energy, Office of Basic Energy Sciences (DOE OBES) grant W-31-109-Eng-38. The Advanced Light Source is operated by Lawrence Berkeley National Laboratory on behalf of DOE OBES and the IDAT program, supported by the DOE Office of Biological and Environmental Research and NIH project MINOS (R01GM105404). CABIMER is supported by the Andalusian Government. Coordinates were deposited in the RCSB Protein Data Bank under 5TVP for TDP2-SUMO2-DNA and 5TVQ for TDP2-SUMO2. Author contributions: conceptualization, M.J.S., F.C.-L., and R.S.W.; methodology, M.J.S., J.G.W., G.A.M., F.C.-L., and R.S.W.; investigation, M.J.S., J.A.L., A.H.-R., L.R.B., J.G.W., A.M.M.-C., and G.A.M.; writing (original draft), M.J.S., F.C.-L., and R.S.W.; writing (reviewing and editing), M.J.S., J.A.L., J.G.W., G.A.M., F.C.-L., and R.S.W.; funding acquisition, F.C.-L., R.E.L., and R.S.W.; supervision, R.E.L., F.C.-L., and R.S.W. The authors declare no competing financial interests.
View Abstract

Navigate This Article