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. Author manuscript; available in PMC: 2026 Oct 27.
Published in final edited form as: Mol Biol Cell. 2026 Oct 26;37(11):ar105. doi: 10.1091/mbc.E25-08-0391

Regulation of TIPIN and replication fork dynamics by USP37

Benjamin R Stromberg 1,2, Luke Scarberry 1,2, Irene S Cho 1, Matthew K Summers 1,*
PMCID: PMC13704607  NIHMSID: NIHMS2213187  PMID: 42747997

Abstract

The deubiquitinating enzyme, USP37, is associated with control of replication and the replication stress response. Cells with oncogene-induced replication stress harbor elevated levels of USP37 and are more sensitive to its loss. These cells experience increased replication stress and delayed progression of the replication fork in the absence of USP37. Consistent with these observations, we find that USP37-deficient cells more frequently fail to restart stalled forks and that these forks are more susceptible to nuclease activity. Underlying these phenomena, we show that USP37 resides at the replication fork and associates with the replisome where it is able to act on TIPIN to promote its stable association with chromatin. These findings suggest the regulation of proteins at the replication fork contributes to fork function and underlies the requirement of cancer cells for USP37.

INTRODUCTION

DNA replication occurs at the replication fork and is carried out by a large protein complex, termed the replisome. Within this molecular machine are the key enzymatic activities necessary to duplicate DNA. These include the CMG (CDC45, MCM2-7, and GINS1-4) helicase, which unwinds the DNA and the replicative polymerases (Polε, Polα, and Polδ) which generate the nascent DNA. Replisomes may encounter numerous challenges to replication, including dNTP deficiencies, DNA lesions, and collisions with transcriptional machinery. Such replication stress and alterations in response to it are associated with numerous human diseases. For example, the elevated expression or activating mutation of oncogenes such as Cyclin E drive proliferation and elevate the basal level of replication stress by inducing inappropriate S-phase entry, deregulating origin firing patterns and altering transcriptional programs. (Bartkova et al., 2005; Gorgoulis et al., 2005; Jones et al., 2013; Kotsantis et al., 2016; Macheret and Halazonetis, 2018). The high levels of replication stress initially function as a barrier to transformation and cells that evolve to form tumors must develop means to alleviate the deleterious effects of abnormal replication programs.

Replication stressors often cause slowing or stalling of polymerases while the helicase continues to unwind the DNA leaving spans of exposed single-stranded DNA (ssDNA), which triggers activation of the ATR stress response kinase. ATR then phosphorylates a host of substrates, including the effector kinase CHK1, to promote cell cycle arrest and limits general firing of origins while promoting the firing of dormant origins surrounding the stalled fork (Kumagai and Dunphy, 2000; Liu et al., 2000; Shimuta et al., 2002; Chini and Chen, 2003; Jeong et al., 2003; Jin et al., 2003; Kumagai and Dunphy, 2003; Sorensen et al., 2003; Xiao et al., 2003; Kumagai et al., 2004; Lee et al., 2005; Sorensen et al., 2005; Syljuasen et al., 2005; Zachos et al., 2005; Chini et al., 2006; Liu et al., 2006; Petermann and Caldecott, 2006; Petermann et al., 2006; Wang et al., 2006; Maya-Mendoza et al., 2007; Cimprich and Cortez, 2008; Ge and Blow, 2010; Merry et al., 2010; Petermann et al., 2010). In addition to initiating replication at dormant origins, cells employ multiple mechanisms to rescue and restart stalled replication forks. A prominent mechanism for restart is fork reversal where the stalled fork undergoes remodeling by enzymes such as SMARCAL1 and HTLF, and the ssDNA is reannealed to form a Holliday junction. However, the newly annealed DNA junction resembles a double stranded break and is susceptible to nuclease activity (e.g., MRE11, DNA2, EXO1, MUS81) (Thangavel et al., 2015; Vallerga et al., 2015; Lemacon et al., 2017). A host of factors (e.g., BRCA2, FANCD2, ABRO1, 53BP1) contribute to the protection and stabilization of the remodeled fork to allow subsequent restart (Ciccia et al., 2012; Berti et al., 2013; Betous et al., 2013; Couch et al., 2013; Wan et al., 2013; Fugger et al., 2015; Kile et al., 2015; Thangavel et al., 2015; Vallerga et al., 2015; Zellweger et al., 2015; Bhat and Cortez, 2018; Bai et al., 2020; Berti et al., 2020; Liu et al., 2020).

The Fork Protection Complex (FPC), a module within the replisome made up of TIPIN, TIMELESS, CLASPIN, and WDHD1, promotes multiple aspects of replication including replication stress tolerance (Lee et al., 2003; Lee et al., 2005; Gotter et al., 2007; Unsal-Kacmaz et al., 2007; Yoshizawa-Sugata and Masai, 2007; Lou et al., 2008; Scorah and McGowan, 2009; Smith et al., 2009; Kemp et al., 2010; Izawa et al., 2011; Leman and Noguchi, 2012; Smith-Roe et al., 2013; Hao et al., 2015; Bianco et al., 2019; Lerner et al., 2020). However, WDHD1’s classification as a core FPC component remains less clear, given its non-overlapping functions and limited evidence for interaction with other FPC members within the replisome (Villa et al., 2016; Jones et al., 2021). The FPC interacts with the CMG helicase and Polε to promote stable association of the polymerase with the fork to ensure efficient DNA replication (Gotter et al., 2007; Hodgson et al., 2007; Lou et al., 2008; Errico et al., 2009; Scorah and McGowan, 2009; Numata et al., 2010; Izawa et al., 2011; Sercin and Kemp, 2011; Aria et al., 2013; Cho et al., 2013; Lerner et al., 2020; Jones et al., 2021; Baris et al., 2022). Loss of FPC proteins thus leads to diminished fork progression, increased fork stalling and collapse (Petermann et al., 2008). Recently it has been identified that WDHD1, TIMELESS, and TIPIN, can protect the nascent DNA from degradation (Abe et al., 2018; Rageul et al., 2020). Furthermore, TIMELESS and TIPIN have shown to be necessary for restart of stalled forks (Cali et al., 2016; Rageul et al., 2020). In addition to these activities within the replisome, the FPC proteins facilitate the global replication stress response by mediating the phosphorylation of CHK1 by ATR (Lee et al., 2003; Lee et al., 2005; Unsal-Kacmaz et al., 2007; Lou et al., 2008; Kemp et al., 2010; Smith-Roe et al., 2013; Hao et al., 2015). Importantly, overexpression of TIMELESS and CLASPIN is common in cancer and promotes replication stress tolerance (Bianco et al., 2019).

The deubiquitinating enzyme, USP37, is emerging as a key regulator of replication. USP37 has been shown to enhance the expression of Cyclin A and CDT1 to promote replication initiation(Huang et al., 2011; Hernandez-Perez et al., 2016). In addition, USP37 promotes replication stress response by stabilizing CHK1 (Stromberg et al., 2021). Similarly, by stabilizing CDT1 USP37 regulates early S phase and origin licensing(Hernandez-Perez et al., 2016). Interestingly, USP37 knockdown restricts fork progression and increases origin firing (Hernandez-Perez et al., 2016; Chauhan et al., 2023). The increase in origin firing may result from diminished CHK1 activity and could also be caused by decreased fork progression or diminished fork restart. However, it isn’t clear how USP37 is promoting fork function. USP37 has been identified at ongoing replication forks, and it has been shown to interact with PCNA (Dungrawala et al., 2015; Chauhan et al., 2023). Notably, USP37 did not deubiquitinate PCNA which suggests that USP37 could be regulating other components of the replication fork.

Here, we show that cells with oncogene-induced replication stress harbor elevated USP37 levels and are more dependent on USP37 to mitigate replication stress caused by dysfunctional replication forks. These results suggest that cancer cells overexpress USP37 to enhance the tolerance of replication stress. Knockdown of USP37 reduces DNA replication fork progression, fork restart, and protection of nascent DNA. Furthermore, we find that USP37 binds to, deubiquitinates, and stabilizes the FPC component, TIPIN. This ability of USP37 to stabilize TIPIN and promote replication fork protection explains, at least in part, the alterations in DNA replication caused by the loss of USP37 observed by us and others(Hernandez-Perez et al., 2016; Chauhan et al., 2023; Tang et al., 2025).

RESULTS

USP37 promotes tolerance of oncogene-induced replication stress

Consistent with reports of elevated USP37 expression across multiple cancers, we find that USP37 protein levels are dramatically higher in cancerous epithelial cells from breast, colon, and lung compared to immortal, but non-transformed, epithelial cells from these tissues (Figures 1A-C). As we, and others, have shown that USP37 promotes efficient replication and replication stress tolerance, these data suggest that the elevated expression of USP37 in cancerous cells reflects the elevated replication stress inherent to transformed cells. We began exploring this idea further by looking at Cyclin E, an established driver of oncogenic replication stress. USP37 expression shows a positive correlation with expression of both CCNE1 and CCNE2 (Figure 1D) (Barretina et al., 2012; Cancer Cell Line Encyclopedia and Genomics of Drug Sensitivity in Cancer, 2015; DepMap, 2024). While consistent with the idea that elevated USP37 may reflect a need to buffer Cyclin E driven replication stress, it is possible that the similar expression profiles of these genes simply reflect that all three are involved in S-phase. We therefore tested the idea that cells with a gain or amplification of CCNE1 or CCNE2 would be more dependent on USP37-mediated replication stress tolerance. Examining the gene effect score of USP37 knockout in the DepMap data set revealed increased sensitivity to loss of USP37 in cell lines with 3 or more copies of either CCNE1 or CCNE2 compared to those without gain/amplification (Figure 1E) (Barretina et al., 2012; Cancer Cell Line Encyclopedia and Genomics of Drug Sensitivity in Cancer, 2015; Tsherniak et al., 2017; DepMap, 2024; Arafeh et al., 2025). Similar trends were observed in BRCA1 mutant breast cancer cell lines, another setting of replication stress (Figure S1), in this dataset.

Figure 1.

Figure 1.

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Cyclin E overexpression causes an increase and dependency on USP37. (A, B, C) USP37 protein expression in immortal and transformed cells of the colon, breast and lung. (D) Pearson correlation analysis of expression of USP37 versus CCNE1 and CCNE2 in the DepMap portal data. (E) USP37 gene effect score (DepMap Public 24Q4 +Score, Chronos) from the DepMap CRISPR dataset for cells with (3 or more copies – relative to ploidy) or without amplification of CCNE1 or CCNE2. Statistical significance was determined using an unpaired one-tailed t test (*, p<0.5). (F, G) Expression of USP37 and Cyclin E in HCT116 cells with the indicated genotypes. (F) and quantitation of USP37 protein (G), Statistical significance was determined using an unpaired two-tailed t test (*, p<0.5), n=3. (H, I) USP37 protein expression was analyzed in a cycloheximide (CHX) chase assay (H) and quantitation of USP37 protein (I), n=3. (J, K) Representative images (J) and quantification (K) of HCT116 and HCT116FBW7−/− colony formation after transfection with the indicated siRNAs. Bar graphs represent the mean and sd, n=3. Statistical significance was determined using two-way ANOVA and Holm-Sidak post-test (*, p<0.5).

To further test the relationship between USP37 and inherent replication stress, we turned to HCT116 cells with or without homozygous deletion of the tumor suppressor FBXW7 (Rajagopalan et al., 2004). As part of the SCFFBXW7 E3 ubiquitin ligase, FBXW7 suppresses protein levels of the replication stress driving oncoproteins Cyclin E and c-MYC (Koepp et al., 2001; Strohmaier et al., 2001; Welcker et al., 2004; Yada et al., 2004; van Drogen et al., 2006; Dominguez-Sola et al., 2007; Jones et al., 2013; Macheret and Halazonetis, 2018). Thus, with this isogenic model, we are able to directly compare effects on basal (HCT116) or elevated replication stress (HCT116FBXW7−/−). We first confirmed the overexpression of Cyclin E in the FBXW7-deficient cells (Figure 1F). In addition to elevated Cyclin E protein levels, we also observed significantly elevated USP37 levels (Figures 1F-G). To determine whether elevated USP37 levels might stem from a loss of SCFFBXW7 E3 ligase activity toward USP37, we measured the stability of USP37 in HCT116 and HCT116FBXW7−/− cells. No significant difference in the stability of endogenous USP37 was observed between the cell lines (Figures 1H-I). These results suggest the possibility that cells with elevated replication stress require additional USP37 and upregulate its expression.

We next assessed whether cells with inherently elevated replication stress were more sensitive to loss of USP37. We measured the impact of USP37 knockdown on the growth of cells with basal (HCT116) or elevated replication stress (HCT116FBXW7−/−). Consistent with our previous findings, loss of USP37 caused a growth defect in parental HCT116 cells. However, depleting USP37 in HCT116FBXW7−/− cells had a significantly greater impact on growth compared to the parental HCT116 cells (Figures 1J-K), consistent with a greater requirement for USP37 in cells with inherent replication stress.

Dysregulation of either Cyclin E or USP37 is detrimental to DNA replication. We therefore hypothesized that the increased sensitivity to loss of USP37 in the HCT116FBXW7−/− cells reflected enhanced replication stress in these cells. We analyzed replication stress by measuring foci of 53BP1 and phospho-RPA S33 (pRPA) by immunofluorescence. In unperturbed cells we measured no significant difference in 53BP1 or pRPA foci when either cell line was depleted of USP37 (data not shown), consistent with our prior observations of 53BP1 foci in USP37-depleted cells and a recent analysis of 53BP1 foci in another FBXW7-deficient model, which found only modest increase in foci (O'Brien et al., 2023). Therefore, we enriched the replication stress response by an acute 30-minute pulse with hydroxyurea (HU). Depletion of USP37 or the absence of FBXW7 caused modest, but not statistically significant, increases in 53BP1 foci formation in HCT116 cells with varying effects on pRPA accumulation. However, depleting USP37 in HCT116FBXW7−/− cells caused significantly increased accumulation of both 53BP1 and pRPA foci compared to loss of USP37 in parental cells or the absence of FBXW7 alone (Figures 2A-D).

Figure 2.

Figure 2.

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Depletion of USP37 impairs fork progression and triggers replication stress response. (A, B, C, D) Representative images (A, B) and quantification (C, D) of phosphor-S33 RPA (pRPA) foci (A, C) or 53BP1 foci (B, D) in in HCT116 and HCT116FBW7−/− cells transfected with the indicated siRNAs and challenged with 1mM HU for 30min. Red lines signify the mean foci number. Statistical significance was determined by one-way ANOVA and the Kruskal-Wallis multiple comparisons test. (E, F) Representative images (E) and quantification (F) of DNA fiber length in HCT116 and HCT116FBW7−/− cells transfected with the indicated siRNAs. Red lines signify mean fiber length. Statistical significance was determined with the Mann-Whitney test. Scale bars in A, C, and D indicates 3μm. (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001).

The results above are consistent with USP37’s role in promoting replication stress tolerance. Because alterations in fork progression may not induce a detectable replication stress response (e.g., 53BP1 foci), we sought additional evidence that exacerbated replication stress underlies the growth defects in USP37-depleted cells. We performed DNA fiber assays to directly analyze replication fork dynamics. Consistent with prior findings, loss of FBXW7 or the depletion of USP37 significantly diminished fiber lengths in the absence of any additional perturbation (Figures 2E-F). In keeping with our findings above, we observed that forks in USP37-depleted HCT116FBXW7−/− cells exhibited significantly decreased replication fiber lengths compared to either USP37-depleted HCT116 cells or cells with FBXW7 loss alone. Together these data suggest that USP37 enhances replication fork function under conditions of stress.

USP37 promotes replication fork dynamics during replication stress.

Data from iPOND-mass spectrometry analyses indicate that USP37 interacts with newly synthesized DNA (Dungrawala et al., 2015; Wessel et al., 2019). In addition, USP37 proximity-labeling in conjunction with proteomic analyses identified potential USP37 associations with components of the replication machinery along with numerous known interactors (Yeh et al., 2015). Notably, FZR1, which directly interacts with USP37, was clearly enriched in pulldowns from BirA*-USP37 fusion-expressing cells over BirA* expressing cells alone in these analyses but did not score as a high confidence interactor. We therefore included proteins that showed clear enrichment in the BirA*-USP37 pulldowns in our analysis even if not reaching statistical significance in the original study (Figure 3A). Consistent with these data, we find that USP37 associates with chromatin during S-phase (Figure 3B). We next confirmed that USP37 resides at the fork using iPOND-western analyses in (Figure 3C). Interestingly, the association of USP37 with nascent DNA was diminished upon induction of stress with HU after the EdU pulse, similar to the behavior of PCNA in these analyses (Figure 3C), suggesting that USP37 may travel with replication forks. Analyses of existing iPOND-proteomic data revealed that the association of USP37 with nascent DNA resembles that of replisome components (Figure 3D) (Dungrawala et al., 2015). And, similar to replisome components PCNA and TIMELESS, USP37 remained chromatin-bound upon HU-induced replication stress despite reduced association with nascent DNA (Figure 3E).

Figure 3.

Figure 3.

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USP37 localizes to replication forks and promotes fork stability. (A) USP37-associated proteins identified by BioID-USP37 (Yeh et al., 2015). Proteins showing enrichment over control are shown. Significance as determined by the original study is indicated. Note that not all known interactors scored as significant. (B) T98G cells synchronized in G1 or S-phase after release from serum-withdrawal were subject to biochemical fractionation. The fractions were probed for the indicated proteins. (C) iPOND immunoblot detecting USP37 and other fork components at nascent DNA in unperturbed HCT116 cells and in cells treated with 4mM HU for 2 hours. No click serves as negative control. (D) Analysis of iPOND-mass spec data from (Dungrawala et al, 2015). Protein abundance for components of the replisome, replication stress response proteins and USP37 are shown. For protein complexes the mean abundance of all components is shown. (E) Immunoblot analysis of the indicated proteins in the chromatin fraction with and without replication stress. (F) DNA fiber assay measuring fork restart by adding 4mM HU for 3hrs between Idu and Cldu pulses then calculating the percentage of stalled forks (Idu alone). The bar graph represents the mean and sd percent stalled forks, n>100 forks per condition. Statistical significance was determined by one-way ANOVA with Dunnet’s multiple comparisons test (**p<0.01, ***p<0.001). (G) DNA fiber assay measuring replication fork resection by following Idu and Cldu pulses with 4mM HU for 3hrs and then normalizing Cldu length to Idu. Red lines signify mean CldU:IdU, n>150 forks per condition. Statistical significance was determined by the Mann-Whitney test (***p<0.0001).

Our data thus far suggest that USP37 promotes fork function. Given its role in promoting replication stress tolerance we asked whether USP37 promotes the ability of stalled forks to restart. We therefore performed DNA fiber assays on USP37-depleted cells that had been challenged with HU between pulses of the nucleotide analogs. USP37-deficient cells exhibited a marked increase in the number of forks that were unable to restart as indicated by failure to incorporate CldU during the second pulse (Figure 3F). Failure to restart stalled forks is often associated with nucleolytic degradation of the nascent DNA. We therefore analyzed resection of nascent DNA by DNA fiber analysis. After challenge with HU, USP37-depleted cells showed pronounced resection as indicated by decreased CldU:IdU ratios (Figure 3G).

Sensitivity to loss of USP37 correlates with loss of replisome function

The data above suggest that USP37 acts on replisome components to maintain fork function. To gain insight into how USP37 might regulate the fork, we compared cellular sensitivities to loss of USP37 with loss of other genes in CRISPR-knockout data in the DepMap dataset (Tsherniak et al., 2017; DepMap, 2024; Arafeh et al., 2025). We reasoned that sensitivity to loss of USP37 would correlate with loss of proteins that it regulates. We examined genes that were screened across all 1178 cell lines in the Public 24Q4+Score, Chronos dataset. Seven of the top ten genes correlating with USP37 loss are involved in replication (Figures 4A, Table S1). Among these are genes encoding three replisome components, TIPIN, POLA2 and CLSPN. TIPIN shows the strongest and most significant correlation to USP37, while POLA2 and CLSPN rank 4th and 6th, respectively. Similarly, USP37 has the highest correlation with TIPIN while it ranks 6th with CLASPIN. In contrast, USP37 does not show a strong reciprocal ranking with POLA2 (Table S1). Several additional replisome components are among the top USP37-correlating genes, but many do not show such a relationship (Table S1). Interestingly, TIPIN and CLASPIN, along with TIMELESS and, potentially, WDHD1, comprise the Fork Protection Complex (FPC), within the replisome. USP37 exhibits significant correlation with all four FPC proteins (Figure 4A). The strength of the correlation between USP37 and TIPIN is similar to that between TIPIN and TIMELESS, which are obligate binding partners (Gotter et al., 2007; Unsal-Kacmaz et al., 2007; Yoshizawa-Sugata and Masai, 2007). Accordingly, TIPIN is the third highest correlating gene with TIMELESS with USP37 ranking 8th (Table S1). WDHD1 shows a weaker correlation with USP37 and generally has a lower correlation with TIPIN, TIMELESS, and CLASPIN than they do with each other (Figure 4A). These results likely reflect unique functions of WDHD1 within the replisome and may support the idea that it is not a component of the FPC (Villa et al., 2016). The FPC promotes multiple aspects of fork function including fork stability as well as promoting the replication stress response via CHK1 activation (Lee et al., 2003; Lee et al., 2005; Gotter et al., 2007; Unsal-Kacmaz et al., 2007; Yoshizawa-Sugata and Masai, 2007; Lou et al., 2008; Kemp et al., 2010; Hao et al., 2015). Interestingly, knockout of two other genes involved in ATR-CHK1 activity, TOPBP1 and TICCR, also correlate strongly with USP37. ATR and CHEK1 show stronger correlations with USP37 than the majority of replisome components (Table S1 and S2). These similar gene dependencies, potential interactions with TIMELESS and WDHD1 (Figure 3A), our observations in altered fork dynamics (Figures 2E-F and 3F-G) and our previous demonstration that USP37 promotes CHK1 function, suggest that USP37 may regulate the FPC (Stromberg et al., 2021).

Figure 4.

Figure 4.

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USP37 binds to TIPIN. (A) Pearson correlation of gene effect scores (DepMap Public 24Q4+Score, Chronos) of the indicated genes in the DepMap CRISPR data (**p<0.01, ****p<0.0001). (B, C) 293T cells were transfected with plasmids encoding USP37-Flag and STag-TIPIN and the expression of these exogenous proteins was measured with immunoblotting, and an interaction between the two proteins was determined by S-Tag (B) or Flag (C) pulldown/immunoprecipitation. (D, E, F) Immunoblot analysis (D) and quantification (E, F) of chromatin-bound protein levels in cells transfected with the indicated siRNAs. The bar graph represents mean and sd of relative protein levels, n=3. Statistical significance was determined by one-way ANOVA with Holm-Sidak post-test (*p<0.05). TPS = total protein stain, WCL = whole cell lysate

USP37 binds to and stabilizes TIPIN

Given the potential interaction with TIMELESS and correlated impacts to cells upon loss of USP37, TIPIN, or TIMELESS we asked whether USP37 could interact with these proteins. We conducted an immunoprecipitation (IP) experiment between USP37 and different components of the fork protection complex. We transfected 293T cells with plasmids encoding USP37-Flag and GFP-S-Tag-TIPIN. USP37 immunoprecipitates contained both TIPIN and TIMELESS while TIPIN was able to reciprocally pull down USP37 as well (Figures 4B-C). These results altogether suggest that USP37 binds to the TIPIN-TIMELESS complex.

TIPIN-TIMELESS localizes to chromatin during S-phase, consistent with its roles in promoting replication and the replication stress response (Yoshizawa-Sugata and Masai, 2007). Moreover, the chromatin-bound complex is subject to ubiquitin-mediated regulation (Rageul et al., 2020). To determine whether USP37 might contribute to regulation of this complex we examined TIPIN and TIMELESS levels in chromatin-enriched fractions of control or USP37-depleted cells. Interestingly, we found that USP37-depleted cells exhibited a loss of TIPIN from chromatin, with minimal impact on TIMELESS suggesting that USP37 specifically regulates TIPIN in chromatin (Figures 4D-F). The soluble pools of TIPIN and TIMELESS were not significantly altered by the absence of USP37 (Figure S2).

We therefore further examined the relationship between USP37 and TIPIN. Given that USP37 prevents degradation of its substrates, we investigated whether it stabilizes TIPIN. 293T cells were transfected with Flag-TIPIN with or without Myc-USP37 or the catalytically dead USP37 mutant (Myc-USP37C350A). Translation was then blocked with CHX, and cells were harvested every 2 hours to measure TIPIN protein levels. Flag-TIPIN alone was unstable and almost completely lost by 6 hours, but in the presence of Myc-USP37 it became significantly more stable (Figures 5A-B). Importantly, the catalytically inactive Myc-USP37C350A was unable to rescue TIPIN stability (Figures 5A-B). We also observed that increasing USP37 expression correlates with TIPIN protein levels (Figure 5C). Furthermore, treating cells with 10 μM MG132 enhanced TIPIN protein levels in a similar manner to USP37 over expression (Figure 5C) suggesting USP37 was preventing proteasome mediated loss of TIPIN. Given that TIMELESS is known to promote TIPIN stability, we asked whether the presence of TIMELESS would impact these observations (Gotter et al., 2007; Unsal-Kacmaz et al., 2007; Yoshizawa-Sugata and Masai, 2007; Kemp et al., 2010). As expected, co-expression of TIMELESS enhanced TIPIN stability (Figures S3A-B), while TIPIN has less impact on TIMELESS similar to prior study (Kemp et al., 2010). The addition of USP37 enhanced the stability of both proteins (Figures S3C-D), consistent with results examining TIPIN alone.

Figure 5.

Figure 5.

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USP37 stabilizes and deubiquitinates TIPIN. (A, B) Immunoblot analysis of 293T cells transfected with Flag-TIPIN and either empty vector, Myc-USP37C350A, or Myc-USP37 and analyzed by immunoblot after treatment with 100 ug/ml cycloheximide (CHX) (A) and quantification of TIPIN expression (B). Graph represents the mean and sd, n=3. Statistical significance was determined a two-way ANOVA with Holm-Sidak post-test compared to vector (*p<0.05, **p<0.01). (C) Immunoblot analysis of 293T cells transfected with Flag-TIPIN and an increasing amount of Myc-USP37. One sample was treated with 10 μM MG132 for 4hrs. (D) 293T cells were transfected with His-Ubiquitin (Ub), Myc-TIPIN, and USP37-Flag or USP37C350A-Flag, as indicated, and treated with 10 μM MG132 for 4hrs. Ubiquitin conjugated proteins were enriched by Ni+2 affinity purification followed by immunoblot for Myc or His epitopes. TPS = total protein stain, WCL = whole cell lysate.

To determine whether USP37 is stabilizing TIPIN by deubiquitinating it, we transfected 293T cells with His-ubiquitin, Myc-TIPIN, and either Flag-USP37 or Flag-USP37C350. Ubiquitination of proteins was enhanced by inhibiting the proteasome with MG132 and then ubiquitinated proteins were purified from lysate with Ni+2 agarose beads. Myc-TIPIN by itself was shown to be highly ubiquitinated, but ubiquitination of TIPIN was drastically reduced by Myc-USP37 (Figure 5D). Conversely, the ubiquitination of TIPIN was unchanged in the presence of Myc-USP37C350.

TIPIN attenuates S-phase progression and replication defects in USP37-depleted cells

We previously reported that USP37-deficient cells exhibit delayed progression through S-phase (Stromberg et al., 2021). To gain insight into the importance of TIPIN regulation by USP37, we asked whether TIPIN expression would attenuate this delayed transit of cells through S-phase. Cells were transfected with empty vector, Flag-S-TIPIN, Flag-TIMELESS, or TIPIN +TIMELESS followed by depletion of USP37. The cells were pulsed with EdU to label replicating cells and were harvested 7 hours later. To examine the progression of replicating cells through S-phase and the cell cycle, we analyzed the EdU-positive (EdU+) cells for the presence of Cyclin A by immunofluorescence. Cells that were replicating at the time of the EdU pulse and had transited beyond early mitosis (M/G1) were identified as Cyclin A-negative (CycA-) and EdU+. Consistent with our prior report, USP37-depleted cells exhibited a reduced fraction of CycA- cells within the EdU+ population (CycA-, EdU+) (Figures 6A-B), demonstrating that S-phase is prolonged upon loss of USP37 (Stromberg et al., 2021). Expression of TIPIN, TIMELESS, or the combination had no impact on the fraction of CycA-,EdU+, cells in control-depleted cells. However, expression of either TIPIN or TIMELESS increased the fraction of CycA-,EdU+, cells in USP37-depleted populations while the fraction of CycA-,Edu+ cells in USP37-deficient populations expressing both proteins was essentially indistinguishable from control cells (Figures 6A-B).

Figure 6.

Figure 6.

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Expression of TIPIN attenuates S-phase progression and replication defects in USP37-deficient cells. (A) Representative images of cells transfected with the indicated siRNAs and either empty vector or FLAG-S-TIPIN 7 hours after a 10 μM EdU pulse. (B) Quantification of the Cyclin A-negative fraction of Edu-positive cells (CycA-,EdU+) cells, treated as in A. The bar graph represents the mean and sd, n=3. Statistical significance was determined a two-way ANOVA with Holm-Sidak post-test compared to vector (*p<0.05, **p<0.01). (C) Quantification of DNA fiber length in Hela cells after transfection with the indicated siRNAs and doxycycline-induced expression of the indicated transgenes. Red lines signify mean fiber length. Statistical significance was determined with a one-way ANOVA and Holm-Sidak post-test. (**p<0.01, ****p<0.0001).

The improved progression of USP37-depleted cells through S-phase upon expression of TIPIN raises the possibility that the fork progression defect observed in the absence of USP37 (Figure 2F) is also attenuated. To investigate this possibility, we generated Hela cells with inducible expression of GFP-S-TIPIN or GFP and performed DNA fiber analyses. Similar to our results with HCT116 cells, depletion of USP37 from Hela cells led to diminished replication tract lengths in cells expressing GFP. Consistent with our results above, expression of TIPIN significantly improved, but did not fully rescue the effect of USP37 loss on fork progression (Figure 6C).

DISCUSSION

USP37 expression is deregulated in numerous cancers, including breast, colon, lung, brain, and blood (Sun et al., 2006; Pyeon et al., 2007; Ding et al., 2008; Finak et al., 2008; Skrzypczak et al., 2010; Huang et al., 2011; Yang and Shih, 2013; Pan et al., 2015; Typas et al., 2015; Hernandez-Perez et al., 2016; Qin et al., 2018; Cai et al., 2019; Xiao et al., 2019; Hong et al., 2020; Qin et al., 2021; Wu et al., 2021a; Wu et al., 2021b; Cao et al., 2022; Kamran et al., 2023). Increased USP37 expression is associated with poor prognosis (Bianchi et al., 2007; Qin et al., 2018; Wu et al., 2021a; Cao et al., 2022) and has been shown to promote transformation (Yang and Shih, 2012). USP37 is also required for embryonic development in zebrafish and mice (IMPC; Tse et al., 2009; Stromberg et al., 2021). The biological role(s) of USP37 underlying these phenotypes is poorly understood.

We show that cells with increased levels of Cyclin E, a driver of replication stress, are particularly susceptible to the loss of USP37. Furthermore, we have shown that the increase in Cyclin E is likely to drive an increase in USP37. This is supported by our prior work which has shown that expression of Cyclin E induces elevated endogenous USP37 levels (Huang et al., 2011). Cyclin E/CDK2 can activate USP37 by phosphorylation which stabilizes USP37 due to its ability to deubiquitinate itself (Huang et al., 2011). However, we did not detect any difference in USP37 stability between cells with normal or high levels of Cyclin E (Figures 1H-I). This data suggests that the increase in USP37 is not due to enhanced activity caused by Cyclin E/CDK2-mediated activation of USP37. Additionally, it shows that USP37 is not an FBXW7 substrate because the loss of FBXW7 would increase the stability of its substrates. It is more likely that Cyclin E increases USP37 transcription indirectly by enhancing the activity of E2F (Duronio et al., 1996). However, as USP37 expression has also been shown to correlate with c-MYC, another driver of replication stress, it is possible that elevated expression of USP37 is selected for as a result of the replication stress in these cells.

Both Cyclin E and USP37 are highly active during S phase and regulate DNA replication and replication stress response, respectively (Dulic et al., 1992; Lukas et al., 1997; Coverley et al., 2002; Huang et al., 2011; Hernandez-Perez et al., 2016; Stromberg et al., 2021). We therefore hypothesize that the replication stress induced by overexpression of Cyclin E is being mitigated by USP37. This idea is supported by the increase in replication stress response, measured by 53BP1 and pRPA, following depletion of USP37 in the HCTFBXW7−/− cells (Figures 2A-D). The loss of USP37 also caused a modest, but insignificant, increase in 53BP1 and pRPA foci in the parental HCT116 cells. It had also been shown that depleting USP37 levels reduces DNA replication in U2OS cells (Hernandez-Perez et al., 2016; Chauhan et al., 2023). We saw a similar impact in both the HCT116 and HCTFBXW7−/− as well as Hela cells. Interestingly, the cells that accumulated the most replication stress had the least efficient fork progression. It could be that the cells are straining to replicate DNA which leads to replication stress response. We and others have been unable to detect an increase in replication stress markers in USP37-deficient cells without induction of additional stress. However, the decreased fork dynamics in the absence of USP37 clearly demonstrate that these cells are experiencing replication stress. This may reflect an inability of IF to detect low levels of replication stress response; a similar effect to low doses of aphidicolin which restrict DNA replication without eliciting a robust stress response (Koundrioukoff et al., 2013).

Using iPOND-western analysis we have confirmed that USP37 resides at the replication fork as implicated by several prior studies (Dungrawala et al., 2015; Yeh et al., 2015; Wessel et al., 2019). Interestingly, our studies showed that USP37 dissociates from nascent DNA upon the induction of stress, which mimics the behavior of replisome components observed by a prior quantitative iPOND-mass spectrometry analysis (Dungrawala et al., 2015). Indeed, analysis of USP37 in that dataset confirmed that its presence at nascent DNA with and without replication stress closely resembles that of the replisome (Figure 3D). These data strongly suggest that USP37 is functioning at the replication fork but do not explain how or to what extent. We therefore further investigated how the loss of USP37 impacts replication fork function. We measured the stability of forks in the absence of USP37. By measuring the ratio of CldU to IdU with a DNA fiber assay after stalling forks with HU we observed an increase in fork resection without USP37. This observation suggests that USP37-deficient cells are unable to protect the double stranded ends of remodeled replication forks from the nucleases such as MRE11 and DNA2 (Thangavel et al., 2015; Vallerga et al., 2015; Lemacon et al., 2017). Interestingly, a recent report found that USP37 limits nucleolytic degradation of forks in cells with compromised fork stability due to loss of BRCA1 or inhibition of ATR, whereas our findings suggest this fork-protective function of USP37 extends to additional replication stress contexts(Tang et al., 2025). Consistent with the increased fork resection in USP37-deficient cells, we detected an increased failure to restart forks after stalling. Altogether this data shows that without USP37, cells are less efficient at DNA replication, have increased DNA resection at stalled forks, and are less able to restart after stalling.

A prior study utilizing proximity ligation established that USP37 is in the vicinity of numerous replisome components (Yeh et al., 2015). Examination of data from the DepMap CRISPR screen revealed that a cellular sensitivity to the loss or USP37 is more similar to the loss of members of the Fork Protection Complex (FPC), namely TIPIN, TIMELESS, and CLASPIN, than to the replisome components in general (Tsherniak et al., 2017; DepMap, 2024; Arafeh et al., 2025). The FPC contributes to multiple aspects of DNA replication, including fork restart, protection of nascent DNA and activation of CHK1; all of which are impacted by the loss of USP37 (Figures 3F-G) (Stromberg et al., 2021). Furthermore, USP37 and the FPC are markedly displaced from nascent DNA under replication stress whereas the MCM and GINS subcomplexes of the replisome display a less pronounced loss (Figure 3D). For these reasons we investigated the relationship between USP37 and the FPC.

Immunoprecipitation experiments in 293T cells with exogenously expressed proteins identified that USP37 interacts with TIPIN (Figures 4B-C). A complimentary IP that pulled down TIPIN and probed for USP37 was able to confirm these findings. Although little is known about the regulation of their protein levels, TIPIN and TIMELESS show S-phase specific localization to chromatin and do undergo proteasome-dependent loss from chromatin (Rageul et al., 2020). We show that TIPIN, and to a much lesser extent TIMELESS, proteins were lost from chromatin in USP37-deficient cells suggesting that USP37 regulates TIPIN at the replication fork. Consistent with this idea, USP37 stabilizes TIPIN in a cycloheximide (CHX) chase assay in an activity dependent manner alone or in the presence of TIMELESS. Ectopic USP37, but not the catalytically inactive mutant, is able to deubiquitinate TIPIN in vivo. As USP37 was able to promote the stability of both TIPIN and TIMELESS, it is curious that a greater impact was observed on TIPIN in the chromatin than on TIMELESS upon loss of USP37. It is likely that another factor, such as TIMELESS binding SDE2, which has been shown to stabilize TIMELESS in chromatin, is maintaining this pool in the absence of USP37. Altogether these results strongly suggest that TIPIN is a USP37 substrate, and that USP37 knockdown reduces TIPIN levels at replication forks causing decreased DNA replication, an increase in nascent DNA resection, and impaired fork restart.

This idea is further supported by our finding that exogenous TIPIN mitigates the delays in fork progression and transit through S-phase we observe in USP37-deficient cells. Interestingly, TIMELESS had a similar ability to rescue S-phase progression while the combination of TIPIN and TIMELESS completely restored progression. The ability of TIMELESS to rescue progression may seem unexpected. However, in addition to stabilizing TIPIN, TIMELESS promotes localization of TIPIN to chromatin, suggesting that the exogenous TIMELESS recruits the soluble pool of TIPIN to chromatin and is consistent with the improved rescue when coexpressed with TIPIN (Yoshizawa-Sugata and Masai, 2007; Kemp et al., 2010). The FPC is important for both fork function (efficient replication rates, fork stability during stalling) and for efficient CHK1 activation (Lee et al., 2003; Lee et al., 2005; Hodgson et al., 2007; Unsal-Kacmaz et al., 2007; Lou et al., 2008; Petermann et al., 2008; Voineagu et al., 2008; Errico et al., 2009; Scorah and McGowan, 2009; Smith et al., 2009; Kemp et al., 2010; Numata et al., 2010; Izawa et al., 2011; Sercin and Kemp, 2011; Leman and Noguchi, 2012; Aria et al., 2013; Cho et al., 2013; Smith-Roe et al., 2013; Akamatsu and Kobayashi, 2015; Hao et al., 2015; Cali et al., 2016; Abe et al., 2018; Lerner et al., 2020; Jones et al., 2021; Baris et al., 2022). Importantly, these activities are separable. In fact, as interaction of TIPIN with RPA is required for CHK1 activity, it has been suggested that FPC recruited to ssDNA by RPA activates CHK1 rather than replisome associated FPC) (Kemp et al., 2010; Vipat and Moiseeva, 2023). Thus, our data support a model in which regulation of TIPIN stability and the FPC at the fork/RPA by USP37 maintains both normal rates of replication and efficient CHK1 activation (Figure 6). CHK1 also regulates replication fork dynamics (Petermann et al., 2006; Maya-Mendoza et al., 2007; Scorah and McGowan, 2009; Petermann et al., 2010; Gonzalez Besteiro et al., 2019). Further investigations will be necessary to define the regulation TIPIN pools by USP37 and determine the contributions of TIPIN and CHK1 to altered fork dynamics of USP37-deficient cells.

Several recent reports also show the association of USP37 with the replisome and implicate USP37 in preventing disassembly of the CMG complex by antagonizing ubiquitination of MCM7, which triggers extraction and disassembly of CMG from chromatin by the p97 AAA+ ATPase (Dewar et al., 2017; Le et al., 2021; Villa et al., 2021; Xia et al., 2021; Bolhuis et al., 2025; Kochenova et al., 2025; Villa et al., 2025) USP37 prevents unscheduled replisome unloading. MCM7 ubiquitination is catalyzed by two E3 ligases; CRL2LRR1 at terminating forks and TRAIP at interstrand cross-links and protein-DNA cross-links (Dewar et al., 2017; Sonneville et al., 2017; Wu et al., 2019; Fan et al., 2021; Jenkyn-Bedford et al., 2021; Le et al., 2021; Villa et al., 2021). These studies support a role for USP37 in antagonizing TRAIP-mediated MCM7 ubiquitination when topoisomerase is inhibited (Kochenova et al., 2025; Villa et al., 2025). While premature CMG disassembly triggered by CRL2LRR1 would certainly contribute to the altered fork dynamics that we observe in the absence of USP37, the DNA structure of the replication fork has been indicated to prevent ubiquitination of MCM7 by CRL2LRR1, suggesting that MCM7 deubiquitination is not required at active replication forks (Deegan et al., 2020; Jenkyn-Bedford et al., 2021). Yet, LRR1 deficiency improves the clonogenic growth of USP37-deficient cells under replication stress (Villa et al., 2025). This finding is consistent with regulation of MCM7-ub but does leave open the possibility that CRL2LRR1 activity toward other substrates contributes to the effect as well.

Additionally, it is possible that changes in fork remodeling and nuclease activities at USP37-deficient forks lead to altered DNA structures that are permissive for CRL2LRR1-dependent MCM7-ubiquitination at stalled forks(Tang et al., 2025). Interestingly, TIPIN-TIMELESS, which can be removed from chromatin independently of MCMs, interacts with CRL2LRR1 and can be ubiquitinated by the ligase in vitro (Somyajit et al., 2017; Rageul et al., 2020; Patel et al., 2023). Thus, one possibility, consistent with rescue of USP37-deficiency phenotypes by both loss of LRR1 and expression of TIPIN-TIMELESS is that loss of TIPIN-TIMELESS leads to altered replisome function and fork stability, which leaves the replisome susceptible to CRL2LRR1. Alternatively, it is possible that USP37 acts on different proteins depending on the nature of the challenge encountered by replication forks and these observations represent distinct events. Future studies on protein alterations at forks under different conditions will be required to fully understand the function of USP37 at this location.

The ability of USP37 to promote replication fork function is another mechanism by which USP37 enables cancer growth. Other studies have shown that USP37 can stabilize proteins that lead to proliferation such MYC, ERα, and HIF2α (Pan et al., 2015; Hong et al., 2020; Cao et al., 2022). We have shown that USP37 also enables unregulated growth by promoting replication stress tolerance through the stabilization of CHK1, and now TIPIN. Without USP37, the inherent replication stress induced by proliferation can become toxic and lead to cell death. Inhibiting USP37 could be a viable therapeutic and potentially synergistic with chemotherapies which elicit replication stress.

MATERIALS AND METHODS

Reagents and Resources

See Table S3.

Cell lines, tissue culture, and transfections

293T, T98G, CCD-18Co, MCF10A, MCF7, MDA-MB-231, MDA-MB-468, T47D, NuLi-1, HBEC-3KT and Hela cell lines were obtained from ATCC. HT29 and SW480 cells were gifts from Dr. Janet Houghton. H1299 and A549 cells were gifts from Dr. Anne Strohecker. Parental and FBWX7−/− HCT116 cells were generously provided by Dr. Bert Vogelstein (Rajagopalan et al., 2004). HeLa Flp-In T-REx cells were a gift from Dr. Stephen S. Taylor (Westhorpe et al., 2011). All cell lines used were routinely tested for mycoplasma. MCF10A cells were grown in DMEM/F12 media supplemented with 5% FBS, 20 ng/mL EGF, 0.5 mg/mL hydrocortisone, 1 mg/mL cholera toxin, 10 μg/mL insulin and 1% Pen/Strep. NuLi-1 and HBEC-3KT cells were grown in Airway Epithelial Cell Basal Medium and Bronchial Epithelial Cell Growth Kit (ATCC, additives. All cell other lines were cultured in DMEM (Corning) with 10% FBS (Avantor) and 1% Pen/Strep (ThermoFisher). All cells were kept at 37°C and 5% CO2. Lipofectamine RNAiMAX (ThermoFisher) was used for siRNA transfection, and plasmid transfections were performed using TransIT-LT1 (Mirus Bio) following the manufacturer’s protocols. All siRNAs used have been previously validated by other publications (Huang et al., 2011; Stromberg et al., 2021). For the generation of cells with inducible expression of GFP or GFP-TIPIN, HeLa Flp-In T-REx cells were maintained in DMEM containing tetracycline-free FBS (Takara) and co-transfected with pGLAP1 or pGLAP1 containing TIPIN cDNA or empty vector and pOG44 in a 1:9 ratio. After 48 hours, 200 μg/mL of hygromycin B (Fisher) in DMEM was added to the cells for selection. After 10 days, the stable cells were maintained in 50 ug/mL of hygromycin B in DMEM.

Plasmids

Plasmids encoding USP37 and 6xHIS-Ubiquitin were previously described (Huang et al., 2011; Burrows et al., 2012; Stromberg et al., 2021). pENTR223 TIPIN (clone HsCD00368791) was obtained from the Harvard PlasmID repository, and a stop codon was inserted by Q5 site-directed mutagenesis. The resulting cDNA was then moved into pCS2-MYC, pGLAP1 or pGLAP2 destination vectors by Gateway cloning (Torres et al., 2009). pcDNA4-Flag-Timeless was a gift from Aziz Sancar (Addgene plasmid # 22887; http://n2t.net/addgene:22887; RRID: Addgene 22887) (Unsal-Kacmaz et al., 2005). cDNA inserts were verified by sequencing.

Drug treatments

Hydroxyurea (HU) was dissolved in water at a concentration of 1M. For immunofluorescence experiments HU was used at 1mM for 30 minutes and for the DNA fiber assays HU was used at 4mM for 3 hours. Idu and Cldu were dissolved in DMSO and used at 50 uM and 200 uM respectively. Cycloheximide (CHX) was dissolved in DMSO and used at a concentration of 100μg/ml. MG132 was dissolved in DMSO and used at a concentration of 10μM for 4 hours.

Immunoblotting

Cells were lysed with EBC buffer (50mM Tris pH 8.0, 150mM NaCl, 0.5% NP40, 1mM DTT, with protease and phosphatase inhibitor tablets (ThermoFisher), and the concentration was measured using Pierce bicinchoninic acid assay (ThermoFisher). Lysates were denatured with Laemmli buffer and boiling for 5 min, then resolved with a hand-cast SDS-PAGE gel. The proteins were then transferred to polyvinylidene difluoride membrane (MilliporeSigma) and stained with Fast Green (MilliporeSigma) to measure total protein loaded. The membrane was blocked with 5% nonfat dried milk in Tris-buffered saline with Tween (0.14M NaCl, 3mM KCl, 2.5mM Tris, pH 7.4, and 1% Tween) (TBST). All primary antibody incubations were performed with shaking at at 4°C for 16 hours to detect endogenous proteins or at room temperature for 1hr to detect epitope tagged proteins. All secondary antibody incubations were performed with shaking at room temperature for 30 minutes in TBST. Washing steps were performed using TBST and protein bands were visualized using the LI-COR Odyssey CLx infrared imaging system.

Colony Formation Assays

HCT116 and HCT116 FBWX7−/− cells were transfected with siRNA and sparsely plated in triplicate into 6 well plates 24 hours post transfection. Fresh media was added every three days for 10 to 14 days. At completion, cells with fixed with methanol, stained with 0.5% w/v crystal violet in 25% methanol, and imaged using the LI-COR Odyssey CLx infrared imaging system.

Immunoprecipitation

Cells were transfected with the indicated plasmids and lysed in EBC buffer 48 hours after transfection. For flag immunoprecipitation, 1μg of the indicated antibody was mixed equal volumes lysate at 4°C overnight. Magnetic Protein A and Protein G beads (ThermoFisher) were then added and incubated at 4°C for an additional 30 minutes. For S-tag immunoprecipitation, equal volumes of lysate were mixed with S-protein agarose beads (MilliporeSigma). Beads were washed three times with EBC buffer and proteins were eluted by boiling beads in Laemmli buffer for 5 minutes and visualized by WB.

Immunofluorescence and microscopy

For detection of DNA damage HCT116 and HCT116 FBW7−/− cells were seeded on glass coverslips (that had been washed in two-parts nitric acid and one-part hydrochloric acid) then transfected with the indicated siRNAs. After 24 hours cells were treated with HU then fixed with cold methanol for 10 minutes, washed in PBS, then permeabilized and blocked with 0.2% Triton X-100 and 5% goat serum in PBS for 30 minutes. Cells were incubated with antibodies diluted in blocking buffer overnight. After washing the cells in PBS, they were incubated with Alexa Fluor secondary antibodies (ThermoFisher,1:500) in 5% BSA in PBST for 30 minutes at room temperature. Again, the cells were washed in PBS and then mounted with ProLong Glass Antifade Mountant with NucBlue (ThermoFisher) which also counter stained DNA. Cells were imaged using a Leica DM5500B fluorescent microscope. Images were analyzed and foci quantified with using FIJI.

DNA Fiber Assay

DNA fibers assays were based on a published protocol with a few alterations (Nieminuszczy et al., 2016; Quinet et al., 2017). HCT116 or Hela Flp-In T-REx cells were transfected with the indicated siRNAs and, for Hela Flp-In T-REx cells, transgene expression was induced by addition of 1 ug/mL doxycycline (ThermoFisher). After 24 hours, the cells were pulsed with IdU for 20 minutes followed by CldU for 20 minutes with three PBS washes in between pulses. For fork resection and fork restart experiments HU was added before or after the CldU pulse as previously described. Additionally, the CldU pulse was doubled to 40 minutes for the experiments studying fork restart. After CldU pulse, the cells were trypsinized, harvested, and diluted in PBS to 500 cells per microliter. To lyse the cells down the glass slide, 1.5 to 3 ul of cells suspended in PBS were added to the top of the slide and allowed to dry for 5 minutes before adding 6ul of lysis buffer (0.05% sodium dodecyl sulfate, 200mM Tris-HCl pH7.4, 50mM EDTA). After allowing the cells to lyse for 5 minutes the slides were tilted to approximately 30 degrees to allow the cells to lyse down the microscope slide. Once dry, the DNA was fixed in 3:1 methanol to acetic acid for 10min dried before adding to chilled 70% ethanol overnight. The next day the slides were removed from the ethanol, dried, and denatured with 2.5M HCl at 37°C for 30 minutes. After being denatured the slides were washed with PBST three times and allowed to dry before being blocked in 1% BSA in PBST for 30 minutes. Again, cells were dried, and primary anti-BrdU antibodies were diluted in blocking buffer, added to the slides, and incubated overnight. The next day the slides were washed in PBST three times, allowed to dry, then incubated with secondaries for 1 hour at room temperature. Again, the slides were washed with PBST three times, dried, mounted with Fluoromount G (ThermoFisher) and imaged using a Leica DM5500B fluorescent microscope.

Immunoprecipitation of nascent DNA

Immunoprecipitation of nascent DNA (iPOND) was based on published protocols with little alteration (Sirbu et al., 2012; Dungrawala and Cortez, 2015). Approximately 2x108 HCT116 cells were used per condition. Cells were pulsed with 10 μM EdU (Click Chemistry Tools) for 15min. Where indicated, 2 mM HU was added at the time of the EdU and incubated for 2 hours. EdU was omitted for the no-click reaction. The remainder of the protocol was followed as described and half the sample was loaded on a 4-15 % gradient gel (Bio-Rad) for western analysis.

Rescue of S-phase progression

Plated 30,000 HeLa cells on coverslips, allowing cells to adhere overnight. The cells were then co-transfected with pGLAP2 TIPIN and/or pCDNA4-Flag TIMELESS or empty vector for 24 hours. Fresh media was added to the plates and the cells were then transfected with siUSP37.1, siUSP37.4, or non-targeting siCTRL. The next day the cells were pulsed with 10 μM Edu for 10 minutes, washed and incubated in fresh media for 7 hours, after which they were fixed in ice-cold methanol for 10 minutes. We then proceeded with the Click-IT reaction kit (ThermoFisher). The cells were then blocked in 5% BSA for 1 hour. They were then stained using anti-Cyclin A (SantaCruz). The cells were imaged utilizing the EVOS M7000 microscope at 20x. Edu positive and Cyclin A negative cells were determined utilizing Cell Profiler software.

Statistical analysis

Statistical analyses were performed with GraphPad Prism Software using an unpaired t Test, 1- or 2-way ANOVA as indicated in the figure legends (GraphPad Software, Inc.). A p-value of <0.05 was considered significant.

Supplementary Material

Supplemental Material

Significance Statement.

  • Ubiquitination events carried out by multiple ligases allow cells to circumvent numerous challenges to faithful replication of the genome; however, the role of deubiquitinating enzymes in these processes is less defined.

  • Here we establish that the deubiquitinase USP37 resides at active replication forks promoting fork stability and function, notably under oncogene-induced replication stress.

  • These findings enhance our understanding of the basic mechanisms of replication and how cancer cells cope with intrinsically elevated replication stress.

ACKNOWLEDGEMENTS

The authors thank The Ohio State University Comprehensive Cancer Center’s Genomics Shared Resource for technical support. We also thank members of the Summers laboratory for insightful discussion and constructive comments on the manuscript. We thank Dr. Michael Kemp for providing TIPIN antibody (Yoshizawa-Sugata and Masai, 2007). This work was supported by The Ohio State University Comprehensive Cancer Center/Department of Radiation Oncology start-up funds and NIH grants R01 GM112895 to MKS, Pelotonia Graduate Student Scholar Fellowships to BRS and LS, and a Pelotonia Undergraduate Student Scholar Fellowship to ISC. Research reported in the publication was supported by The Ohio State University Comprehensive Cancer Center and the National Institutes of Health under grant number P30 CA016058. The content of this work is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Abbreviations:

TPS

total protein stain

WCL

whole cell lysate

FPC

fork protection complex

CMG

CDC45, GINS1-4, MCM2-7

USP37

Ubiquitin Specific Protease 37

HU

hydroxyurea

IP

immunoprecipitation

SCF

Skp1-Cul1-F-box

CRL2

Cullin Ring Ligase 2

iPOND

isolation of Proteins on Nascent DNA

DepMap

The Cancer Dependency Map

EdU

5-ethynyl-2`-desoxyuridine

CldU

5-chloro-2'-deoxyuridine

IdU

5-iodo-2'-deoxyuridine

Footnotes

DATA and RESOURCE AVAILABILITY

The data supporting this study are presented in the manuscript and supporting material or available through the DepMap portal. Plasmids generated in this study are available upon request.

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