MAPK Pathway Suppression Unmasks Latent DNA Repair Defects and Confers a Chemical Synthetic Vulnerability in BRAF-, NRAS-, and NF1-Mutant Melanomas

Melanomas can be classified into four genomic subtypes based on the presence or absence of mutations in RAS pathway genes: BRAF, NRAS, NF1, and triple wild-type (WT; ref. 1). Fortunately, selective BRAF and MEK inhibitors, and more recently BRAF/MEK inhibitor combinations, have improved prognosis and overall survival in patients with metastatic BRAF-mutant disease (2). Nevertheless, all individuals ultimately relapse, and do so, on average, in 11 months (3, 4). A fraction of NRAS-mutant tumors (15%) exhibit partial responses to MEK inhibitors, albeit with shorter durations (5). The MEK pathway is also hyperactivated in NF1-mutant melanomas; however, the clinical activity of MEK inhibitors in this subtype is not known. Regardless, these observations suggest that although RAF/MEK pathway inhibition will remain an important cornerstone of melanoma treatment, improved combinations and/or sequential therapies are needed. Accordingly, additional meaningful targets must be identified.

Agents that target epigenetic enzymes are increasingly being developed as potential cancer therapies (6). Histone deacetylase (HDAC) inhibitors are one such class of compounds, and various drugs have been approved for use in hematopoietic malignancies (7). Although single-agent efficacy in solid tumors has not been observed, HDAC inhibitors are currently being evaluated in combination with other targeted agents in several diseases (8). However, the key to discovering successful combinations, if they exist, will likely lie in (i) identifying the most selective/potent agent for the specified target to minimize potential toxicities; (ii) elucidating the mechanism of action of the combination identified; and (iii) using this insight to prospectively identify patients who are most likely to respond. Although these criteria are important for developing any successful combination therapy, they may be critical for developing combinations with HDAC inhibitors, many of which inhibit numerous HDAC isoforms, and therefore have major effects on chromatin.

Here we identify an epigenetic-based combination therapy for BRAF-, NRAS-, and NF1-mutant melanomas. Specifically, we show that HDAC3 is an important therapeutic target in these tumors and that the selective Class I inhibitor entinostat not only dramatically enhances the in vivo efficacy of BRAF/MEK inhibitors in BRAF-mutant malignancies with varying sensitivities to these agents, but also cooperates with MEK inhibitors in NRAS- and NF1-mutant tumors. We further demonstrate that these agents function by coordinately suppressing the transcription of homologous recombination (HR) and nonhomologous end joining (NHEJ) genes, thereby triggering excessive DNA damage in sensitive tumors. Finally, we identify a tractable biomarker that marks melanomas with broad defects in DNA repair genes and predicts efficacy. Together, these studies have identified a promising mechanism-based combinatorial strategy for treating melanomas with several RAS/RAF pathway defects and have outlined a path for clinical translation.

MITF is a lineage-specific survival gene that is amplified in a subset of melanomas and has been shown to confer resistance to MAPK pathway inhibition (9). Because HDAC inhibitors have been reported to suppress MITF expression (10), we reasoned that these agents might potentiate the therapeutic effects of BRAF and/or MEK inhibitors. A panel of melanoma cell lines harboring mutations in BRAF, NRAS, or NF1 was examined in a CellTiter-Glo–based screen. The MEK inhibitor trametinib was used to broadly suppress the MAPK pathway in all cell lines. Notably, the pan-HDAC inhibitor vorinostat dramatically potentiated the effects of trametinib in 6 of 10 lines, including BRAF-, NRAS-, and NF1-mutant cells (Fig. 1A). These effects were determined to be synergistic using the Loewe excess additivity model (Fig. 1B).

Cytotoxicity was confirmed by cell-counting assays and live-cell imaging in the presence of clinically relevant drug combinations. Combined BRAF and MEK inhibitors (dabrafenib/trametinib) were used to suppress the MAPK pathway in BRAF-mutant cells, as this represents the standard of care (2), whereas trametinib alone was used in NRAS- and NF1-mutant cell lines. In both settings, combined MAPK/HDAC suppression resulted in a dramatic loss of viable cells in just 72 hours in sensitive lines representing all three genotypes (Fig. 1C). Dabrafenib/trametinib and vorinostat effectively suppressed their respective targets in all cell lines and the combination did not further reduce ERK phosphorylation in sensitive cells nor attenuate phospho-ERK suppression in resistant cells (Fig. 1C; Supplementary Fig. S1A and S1B). These agents were not generally toxic, as they did not affect the viability of normal melanocytes, unresponsive melanoma cell lines, and a variety of other unrelated cell types (Fig. 1D; Supplementary Fig. S1A–S1D).

IncuCyte live-cell imaging was used to concomitantly measure cell death and changes in cell number over time using a fluorescent assay that detects live and dead cells. BRAF/MEK inhibitors alone killed 9% of cells, consistent with previous reports (Fig. 1E, left; ref. 11). However, although the HDAC inhibitor had little effect on its own, it dramatically potentiated cell death triggered by dabrafenib/trametinib, resulting in death of 29% of cells within just 72 hours (Fig. 1E, left). Cell death was mediated by apoptosis, as confirmed by PARP cleavage and a fluorescent caspase reporter, which similarly revealed that one-third of the cell population was undergoing apoptosis within 72 hours (Fig. 1F; Supplementary Fig. S1E). Importantly, the onset of cell death corresponded to a concomitant reduction in cell number (Fig. 1E, right). Consistent with previous observations, BRAF/MEK inhibitors alone appeared to exert delayed cytostatic effects in vitro (11); however, this apparent cytostasis was likely due to the net effects of concomitant proliferation and lower levels of cell death (Fig. 1E). In contrast, BRAF/MEK and HDAC inhibitors together synergized to cause much more cell death, thereby eliminating more resistant tumor cells (Fig. 1A–C and E).

Melanomas can be categorized as either MITF^hi or MITF^lo (12). On the basis of our original hypothesis, we expected that MITF^hi cells might be more sensitive to these agents, and if so, MITF suppression by HDAC inhibitors would correspond to sensitivity. Surprisingly, however, sensitivity was unrelated to MITF status, as both MITF^hi and MITF^lo cells responded to this combination (Supplementary Fig. S1F). In addition, MITF suppression did not correlate with sensitivity (Supplementary Fig. S1G). Taken together, these observations suggest that HDAC inhibitors broadly potentiate the therapeutic effects of BRAF/MEK pathway inhibitors in a high percentage of melanomas harboring mutations in one of several genes affecting the RAS/RAF pathway, but that these effects are unrelated to MITF status or expression changes.

Before embarking on mechanistic studies, we sought to identify the most clinically tractable agent(s). Vorinostat is a pan-HDAC inhibitor; however, more selective inhibitors have been developed (7). Importantly, the use of more selective compounds might minimize potential toxicities in humans, especially in the context of drug combinations. The effects of vorinostat were compared with mocetinostat (which inhibits Class I HDACs 1–3 and Class IV HDAC 11; ref. 13), nexturastat (a selective HDAC6 inhibitor that primarily affects acetylation of proteins other than histones; ref. 14), and entinostat (which inhibits Class I HDACs 1–3; ref. 7). Nexturastat did not kill cells when combined with MAPK pathway inhibitors, indicating that HDAC6 inhibition is not sufficient for these effects (Fig. 1G). However, mocetinostat and entinostat both cooperated with trametinib and were even more potent than vorinostat, demonstrating that the suppression of Class I HDAC proteins is sufficient for a maximal therapeutic response (Fig. 1G).

We were particularly enthusiastic about entinostat because it is the most selective agent in this panel that exerted therapeutic effects, and it has shown promising responses in phase II drug combination studies in breast cancer (15). It is also currently being evaluated in a variety of solid tumors, underscoring the potential translatability of entinostat-based combinations. Importantly, entinostat also cooperated with MAPK pathway inhibitors in BRAF-, NRAS-, and NF1-mutant cell lines (Fig. 1H–J).

Entinostat suppresses the activity of HDAC1, 2, and 3, and does not affect protein expression, as reported previously (Supplementary Fig. S1H; ref. 16). To determine whether the inhibition of a specific HDAC gene was mediating the therapeutic effects, individual HDACs were genetically suppressed using pooled siRNAs. Interestingly, only HDAC3 suppression was sufficient to kill melanomas when combined with trametinib, which was confirmed using an unrelated panel of shRNA sequences in a second sensitive cell line (Fig. 1K and L). Because entinostat is currently the most selective, clinically available agent that suppresses HDAC3 and is well tolerated even when combined with other agents, we continued our analysis using entinostat. In addition, it should be noted that although pan-HDAC inhibitors also exert activity in this setting, toxicities associated with broad HDAC inhibition have limited their clinical utility in the context of some drug combinations (17), further supporting the selection of entinostat.

Next, we investigated whether these agents could exert similar cooperative effects in vivo, using drug concentrations that mimic human exposures (18). In clinical trials, the BRAF inhibitor dabrafenib exhibits activity as a single agent in BRAF-mutant melanomas; however, when combined with trametinib, efficacy is enhanced and toxicities are reduced due to the suppression of feedback pathways (3, 4). As such, combined BRAF/MEK inhibitors are now the standard of care in BRAF-mutant melanoma. Nevertheless, a range of therapeutic responses are observed in patients. Therefore, we examined several BRAF-mutant melanoma models with differing sensitivities to these agents. The A375 xenograft model was moderately sensitive to the standard of care; average tumor regression in response to dabrafenib/trametinib was 32% (Fig. 2A). However, the inclusion of entinostat at 1 mg/kg per week, a dose that is comparable with the human dose of 5 mg once weekly being used in other clinical combination studies, dramatically enhanced efficacy (P = 0.004). Although entinostat on its own had no effect on tumor growth, when combined with dabrafenib/trametinib, tumors regressed by 70% on average (Fig. 2A).

Next, we examined the effects of entinostat in a human xenograft model that is more sensitive to dabrafenib/trametinib in vivo (Hs695T; Fig. 2B). In this study, we were also interested in examining the durability of the response and therefore continued treatment for 6 weeks. Strikingly, even in this model, entinostat substantially enhanced the therapeutic response to dabrafenib/trametinib, promoting deeper regressions (83% vs. 57%, P = 0.02), demonstrating that this combination is able to kill more residual disease. Moreover, regressions remained durable throughout the entire study.

Finally, the effects of these agents were evaluated in two different genetically engineered mouse model (GEMM) allograft models, with cooperating mutations in genes associated with resistance to BRAF inhibitors (19, 20). Importantly, the immune system is also intact in each of these models. The first model harbored mutations in Braf and Nf1 (19), as NF1 mutations have been shown to functionally confer resistance to MAPK pathway inhibitors (19, 21). As predicted, MEK/BRAF inhibition did not cause any durable regressions in this model, and instead tumors grew 18% on average; however, combined MEK/BRAF/HDAC suppression triggered tumor regression in every animal, and tumors shrunk by 43% (Fig. 2C, P = 0.03). PTEN mutations have been proposed to function by augmenting survival of cancer cells in response to MAPK pathway inhibition and are associated with smaller and less durable clinical responses in patients (22, 23). Consistent with clinical observations, the effect of dabrafenib/trametinib on Braf/Pten-mutant GEMM allografts was also relatively modest in most tumors (Fig. 2D, left), and these lesions ultimately became resistant (Fig. 2D, right). Nevertheless, the inclusion of entinostat caused tumors to shrink by more than 73%, and responses in all tumors were stable (Fig. 2D; P = 0.02). Altogether, preclinical studies in four distinct BRAF-mutant models, representing tumors with distinct genetic alterations and different sensitivities to BRAF/MEK inhibitors (sensitive, moderately sensitive, and resistant), suggest that entinostat can substantially enhance the therapeutic effects of BRAF/MEK pathway inhibitors in vivo in all of these settings. Figure 2C also demonstrates that this combination is effective in vivo in tumors that harbor NF1 mutations.

We also expanded our analysis of NRAS-mutant melanomas. Using a panel of six additional NRAS-mutant cell lines, we found that three were sensitive to combined MEK and HDAC inhibitors whereas three were not, consistent with the frequency of sensitivity to this combination overall (Fig. 2E). Importantly, these agents also cooperatively enhanced the regression of NRAS-mutant xenografts (Fig. 2F). Whereas trametinib alone exerted largely cytostatic effects (on average, tumors grew by 6%), trametinib and entinostat together induced a 42% tumor regression (P = 0.03). This observation is particularly important given that there are currently no effective treatments for NRAS-mutant tumors.

Although several genetically distinct melanoma cell lines and tumor models were sensitive to combined HDAC and MAPK pathway inhibitors, a subset was resistant to this combination (Fig. 1A). Therefore, we hypothesized that these differential responses could be exploited to identify potential biomarkers of sensitivity or resistance. Extensive genomic analysis did not reveal any recurrent mutations or copy-numberalterations that distinguished sensitive or resistant cells. We therefore performed a two-class comparison of transcriptional profiles from the sensitive (n = 6) and resistant (n = 4) cell lines shown in Fig. 1A. Eighteen genes that were differentially expressed (P < 0.001) between these two populations were identified (Fig. 3A; Supplementary Table S1); however, dramatic differences in expression were observed for only one of these genes, O⁶-methylguanine DNA methyltransferase (MGMT), which was elevated in sensitive cells by almost 9-fold (Fig. 3A and B, P = 0.0005). Western blot analysis further revealed that MGMT protein was readily detected in sensitive cell lines, whereas it was undetectable or minimally expressed in resistant cells (Fig. 3C). On the basis of these observations, two additional BRAF-mutant cell lines, predicted by the Cancer Cell Line Encyclopedia (CCLE) to have either high or low levels of MGMT mRNA, were selected for analysis. Immunoblots confirmed the expected differences in MGMT protein levels (Fig. 3D). Importantly, the MGMT-expressing cells were sensitive to dabrafenib/trametinib/entinostat, whereas the cell line that lacked MGMT expression was insensitive to this combination (Fig. 3E). Similarly, the NRAS-mutant melanomas that were sensitive to these agents expressed MGMT, whereas insensitive cells did not (Supplementary Fig. S2). Altogether, analysis of 18 different sensitive and resistant cell lines indicates that MGMT expression is a strong predictive biomarker of sensitivity to this drug combination.

MGMT directly reverses the mutagenic DNA lesion O⁶-methylguanine, which is caused by alkylating agents (24). Accordingly, in glioblastomas, high MGMT levels are associated with resistance to alkylating chemotherapies such as temozolomide (25, 26). In a subset of glioblastomas, the MGMT promoter is epigenetically silenced by methylation, which is thought to be responsible for conferring sensitivity to temozolomide. Therefore, MGMT promoter methylation testing is routinely used in clinical practice as a predictive biomarker to guide patient management in glioblastoma. Consistent with this mechanism of regulation, we found that the MGMT promoter was differentially methylated in MGMT⁺ versus MGMT⁻ melanoma cell lines (Fig. 3F, P = 0.00011) and that treatment with the DNA-demethylating agent 5-azacitidine restored MGMT expression in MGMT⁻ cells (Fig. 3G). However, it is important to note that in this setting, MGMT expression correlates with sensitivity, rather than resistance, to BRAF/MEK/HDAC inhibitors. Regardless, this preexisting clinical test could be readily implemented to select patients for clinical trials. Importantly, two separate studies have shown that the MGMT promoter is methylated in only 21.5%–26% of human metastatic melanomas using this clinical assay (27, 28). Therefore, these observations suggest that up to 79% of patients with RAS/RAF pathway mutations could benefit from this combination.

To determine whether MGMT was a functional or passive biomarker, we genetically ablated it in sensitive cells and ectopically expressed it in resistant cells. Genetic ablation of MGMT did not make sensitive cells become resistant to these agents (Fig. 3H) and ectopic MGMT expression did not confer sensitivity (Fig. 3I). Therefore, we conclude that MGMT does not play a functional role in mediating sensitivity or resistance to this drug combination, but rather marks a distinct subset of melanomas. Regardless, the fact that a clinical test already exists makes this a clinically useful biomarker.

The strong association between MGMT expression and drug sensitivity prompted us to determine whether we could use a larger set of (primary) tumors to identify a specific genetic defect, responsible for sensitivity or resistance, which might cosegregate with high or low MGMT expression. To maximize potential differences, we compared the top 10% of tumors in The Cancer Genome Atlas (TCGA) database expressing the highest levels of MGMT mRNA (MGMT⁺) with the bottom 10% of tumors, expressing the lowest levels of MGMT mRNA (MGMT⁻). Similar to cell line studies, comparative analysis of mutations and copy-number changes did not identify any recurrent genetic alterations that were enriched in MGMT⁺ or MGMT⁻ tumors. However, these tumor cohorts exhibited distinct transcriptional profiles.

To identify potentially defective pathways and/or vulnerabilities in MGMT⁺ melanomas, we examined transcriptional signatures that were suppressed in these tumors using the Gene Ontology (GO):Biological Processes database. Interestingly, among the top 35 suppressed gene sets, numerous signatures associated with DNA repair, in particular double-strand break repair, were identified (Fig. 4A; Supplementary Table S2), suggesting that these tumors might possess inherent defects in DNA repair processes.

Concomitantly, we examined transcriptional profiles of sensitive cells exposed to MAPK/HDAC inhibitors prior to the onset of cell death. In drug-treated cells the Hallmark_DNA_Repair gene set was identified as one of the top significantly suppressed pathways in response to dabrafenib/trametinib/entinostat, suggesting that DNA repair processes were also being inhibited by this combination (Supplementary Table S3). These observations raised the intriguing possibility that preexisting defects in DNA repair, compounded by the chemical suppression of these pathways, might be mediating cell death by causing excessive DNA damage. Of note, an oxidative phosphorylation signature was also observed, but we found that reactive oxygen species were not consistently elevated by these agents, and were neither necessary nor sufficient for this therapeutic response (Supplementary Fig. S3A–S3E).

To investigate a potential role for DNA repair defects in this response, we further characterized MGMT⁺ tumors and cell lines. The transcriptional signatures identified in Fig. 4A suggested that MGMT⁺ tumors might harbor defects in HR. Using a more specific transcriptional dataset, we confirmed that MGMT⁺ tumors were, in fact, enriched for a common signature that is induced by RNAi-mediated ablation of many HR genes, termed the HRD gene signature (Fig. 4B; ref. 29). The HR-defect signature was also enriched in MGMT⁺ melanoma cell lines as compared with MGMT⁻ cells (Fig. 4C). However, to confirm that sensitive cells harbor functional defects in HR, we performed a RAD51 redistribution assay. In response to ionizing radiation, RAD51 normally accumulates in numerous distinct foci at sites of DNA damage (30). However, the appearance of these foci is prevented or reduced in cells with defects in various HR proteins (30). Indeed, whereas ionizing radiation induced a dramatic increase in RAD51 foci that colocalized with phospho-γH2AX in the two resistant cell lines, the two sensitive cell lines were substantially impaired in their ability to form RAD51 foci (Fig. 4D; Supplementary Fig. S4A). These observations demonstrate that sensitive cell lines harbor a preexisting impairment in HR.

In breast and ovarian cancers, BRCA1/2 mutations are known to underlie defects in HR (31). However, more recently, castration-resistant prostate cancers have been shown to harbor alterations in any one of five different HR genes (32). Therefore, we reanalyzed genomic data to determine whether we might have missed a similar heterogeneous mutational pattern of HR genes, but did not find this to be the case.

Because there were no recurrent genetic alterations in HR genes, the relative expression levels of all genes implicated in DNA repair were examined in MGMT⁺ versus MGMT⁻ tumors (list obtained from ref. 33). Strikingly, the majority of DNA repair genes were downregulated in MGMT⁺ tumors as compared with MGMT⁻ tumors (Fig. 4E). Genes known to be involved in HR as well as other DNA repair pathways were suppressed in MGMT⁺ tumors. Importantly, this observation held true even when known cell cycle–regulated genes were removed from the analysis, reinforcing that this is a true DNA repair effect and not a by-product of cell-cycle differences (Supplementary Fig. S4B; ref. 33). A subset of repair genes, which included MGMT and MPG, both involved in the repair of alkylated bases, along with genes that repair hydrolyzed and oxidized bases, clustered together and exhibited the inverse expression pattern as compared with the broader group of genes: These genes were expressed at higher levels in MGMT⁺ tumors and were lower in MGMT⁻ tumors (Fig. 4E). Importantly, this distinct expression pattern of DNA repair genes was not associated with prior exposure to therapy, suggesting that these genes were not selectively repressed or induced as a consequence of treatment. Therefore, there appear to be two potentially distinct populations of melanomas: DNA repair low^(MGMT+) and DNA repair intact^(MGMT−), which we hypothesize are differentially sensitive to BRAF/MEK/HDAC inhibitors. Although the mechanisms that underlie the inverse expression pattern of these gene clusters could not be ascertained, it is well known that genes with similar functions are often coregulated. We believe that this may be occurring in melanoma, especially given the related function of the coregulated genes.

To determine whether the HDAC/MAPKi combination might be capitalizing on this potential defect and killing cells by triggering unresolvable DNA damage, we first compared phospho-H2AX (γH2AX) expression levels, a marker of double-strand DNA breaks, in sensitive and resistant cells. In sensitive cell lines, two waves of γH2AX induction were observed. γH2AX initially increased within 8 hours of treatment, increased more dramatically between 24 and 36 hours, and remained elevated (Fig. 4F). Importantly, γH2AX induction occurred prior to cell death, which begins at 40 hours (Fig. 1E; Supplementary Fig. S1E), suggesting that this event precedes cell death and is not merely a consequence of DNA damage in dying cells. In contrast, γH2AX levels remained low in resistant cells (Fig. 4G). Notably, both agents induced low levels of γH2AX as single agents; however, the drug combination was required to induce maximal DNA damage, demonstrating that these agents cooperatively induce DNA damage (Fig. 4H).

To investigate the molecular mechanism by which this combination was functioning, transcriptional profiles were examined in cells treated with vehicle, dabrafenib/trametinib, entinostat, or all three agents at 24 hours, prior to the commencement of cell death. Striking changes in the expression of DNA repair genes in response to these agents were observed. As depicted in the heat maps shown, dabrafenib/trametinib alone suppressed the expression of many DNA repair genes in sensitive cell lines, which clustered together in two groups, herein referred to as Group A genes (Fig. 4I, left and middle). Interestingly, Group A genes included several HR pathway genes such as BRCA2, BRIP1, EME1, and RBBP8 (33, 34). These four genes were suppressed between 60% and 84%. Entinostat had no effect on the expression of group A genes but in both sensitive cell lines, dabrafenib/trametinib/entinostat together not only inhibited Group A genes but also suppressed an additional large set of DNA repair genes (Group B; Fig. 4I, left and middle). Notably, the most potently suppressed genes in Group B (suppressed 65%–80%) included genes that function in the NHEJ pathway (e.g., XRCC4, XRCC5, XRCC6, PNKP, and PARP3). This striking pattern of transcriptional repression was not observed in resistant cells (Fig. 4I, right).

It should be noted that HDAC inhibitors have previously been reported to induce DNA damage in cells through a variety of direct and indirect mechanisms (35–37). Nevertheless, in this setting, we observed a potent and broad suppression of DNA repair genes that occurred only in the presence of MAPK pathway inhibitors, which was quite unexpected. Therefore, although HDAC inhibitors may contribute to DNA damage via additional mechanisms, the deep suppression of numerous DNA repair genes that occurs in response to this combination is likely to play a major role in this response. The observation that DNA repair gene networks are already impaired in sensitive cell lines further supports this model.

As noted above, Group A genes included several HR pathway genes such as BRCA2, BRIP1, EME1, and RBBP8 (Fig. 4I; refs. 33, 34). Western blot analysis confirmed that dabrafenib/trametinib potently suppressed BRIP1, BRCA2, RBBP8, and EME1 protein expression in sensitive cells (Fig. 5A). BRIP1 and BRCA2 expression was further evaluated in two sensitive and resistant cell lines. Dabrafenib/trametinib nearly depleted the expression of BRIP1 and BRCA2 proteins in sensitive cell lines, but did not do so in resistant cells (Fig. 5B). Quantitative PCR further confirmed that BRAF/MEK inhibitors suppressed the transcription of these HR genes, and did so in sensitive but not resistant cells (Fig. 5C).

To exclude the possibility that differences in expression were a secondary consequence of effects on the cell cycle, sensitive cells were treated with vehicle or dabrafenib/trametinib and cells in G₁, S, and G₂–M were separated by FACS. In both sensitive cell lines, BRIP1 and BRCA2 mRNA levels were suppressed in all phases of the cell cycle (Fig. 5D), demonstrating that the suppression of these genes is not merely due to changes in cell-cycle distribution.

Figures 4A–E demonstrate that sensitive melanomas already have broad defects in the expression of DNA repair genes that result in HR defects, as inferred by transcriptional profiles and validated by RAD51 redistribution assays. However, Fig. 5A–C demonstrate that dabrafenib/trametinib causes a nearly complete suppression of a subset of these HR genes, which would presumably enhance these defects. Indeed, we found that dabrafenib/trametinib caused a potent enrichment of the HR-defect signature in sensitive cells (Fig. 5E). To visualize the progressive suppression of this pathway, we performed single-sample gene set enrichment analysis (ssGSEA) on sensitive and resistant cell lines. Figure 5F demonstrates that genes that are suppressed in response to HR defects are lower in (untreated) sensitive cells compared with resistant cells (sample 3 vs. sample 1, P < 0.0001). Moreover, dabrafenib/trametinib causes a further potent reduction of these genes in sensitive cells (sample 4 vs. 3, P < 0.0001). Taken together, these observations suggest that BRAF/MEK inhibitors are potentiating inherent defects in the HR pathway by suppressing multiple HR pathway genes.

The RAD51 redistribution experiment confirmed that there are substantial defects in HR in untreated sensitive cells; however, the dynamic range of this assay precluded us from determining whether BRAF/MEK inhibitors could further potentiate these defects. HR impairment can also be measured by an I-Sce-I–based double-strand break repair assay; however, this requires reporter integration into a single genomic site, specifically in sensitive cells, which was unachievable after extensive effort. However, it is well established that genetic defects in HR genes, such as BRCA1 and BRCA2, confer sensitivity to PARP inhibitors, which has become a hallmark functional assay for measuring HR defects (31). Given the potent suppression of HR gene expression, in particular BRCA2, we hypothesized that BRAF/MEK inhibitors would sensitize (or further sensitize) responsive cell lines to PARP inhibitors.

Interestingly, we found that the PARP inhibitor olaparib did not exert potent cytotoxic effects as a single agent in sensitive melanomas, in contrast to BRCA1-deficient breast cancer cells (Fig. 5G). These observations suggest that the DNA repair defects present in these melanomas are less pronounced than those in BRCA1-mutant tumor cells, consistent with the observation that these cells still express HR genes (Fig. 5A). However, dabrafenib/trametinib potently sensitized melanomas to olaparib, and together dabrafenib/trametinib/olaparib killed multiple sensitive cell lines and did not kill nonresponsive cell lines (Fig. 5G). Strikingly, these observations were also recapitulated in vivo. Although the PARP inhibitor olaparib alone was unable to cause tumor regression, when it was combined with dabrafenib/trametinib tumors regressed by 47% on average (P = 0.03, Fig. 5H). Taken together, these results suggest that although baseline defects in DNA repair gene expression prime sensitive cells, MAPK pathway inhibition is required to functionally impair the HR pathway in these melanomas to the extent observed in BRCA1-mutant breast cancers.

Finally, to prove that defects in HR functionally contribute to the therapeutic response to combined MAPK/HDAC inhibitors, we examined the effects of RAD51 overexpression. A key step in HR is the recruitment of the RAD51 recombinase to double-strand breaks, which normally requires HR proteins such as BRCA2 (34). However, it is well known that overexpression of RAD51 can override this regulatory step in cells with various upstream HR defects (34). Therefore, we reasoned that RAD51 would be the one component that might rescue effects caused by the concomitant suppression of multiple HR proteins. Importantly, RAD51 overexpression suppressed death in response to MAPK/HDAC inhibition, confirming that defects in HR and consequently excessive DNA damage play critical functional roles in mediating the therapeutic response to these agents (Fig. 5I). It should be noted that RAD51 did not rescue the limited response to BRAF/MEK inhibitors alone, indicating that these agents mediate their canonical effects through other pathways, as would be expected.

We next sought to identify critical genes in Group B that might be responsible for the cooperativity between dabrafenib/trametinib and entinostat. Interestingly, the most potently suppressed Group B genes that could be ascribed to a specific DNA repair pathway are known to function in the NHEJ pathway (a subset listed in Fig. 4I). Importantly, it is well established that when HR is impaired, NHEJ can compensate (38). Therefore, we examined a subset of genes in the NHEJ pathway that were the most transcriptionally suppressed, specifically XRCC5 (33, 39), PNKP (33, 39), and PARP3 (33, 40), and confirmed that protein expression was also potently inhibited by dabrafenib/trametinib/entinostat (Fig. 6A). We also showed that suppression occurred at the level of transcription and occurred in different sensitive cell lines, and that these genes were not suppressed in insensitive cells (Fig. 6B). Together, these results demonstrate that combined HDAC and MAPK inhibitors potently suppress both HR and NHEJ genes.

To functionally determine whether the suppression of NHEJ pathways was required for cell death in response to this combination, we ectopically expressed LIG4. LIG4 directly mediates the NHEJ ligation step and its overexpression, analogous to RAD51 overexpression, is the one distal gene that can largely override deficiencies in (multiple) upstream components (39). Importantly, LIG4 substantially inhibited the cytotoxic effects of MAPK/HDAC suppression (Fig. 6C; P = 0.009). Taken together with the RAD51 overexpression experiment in Fig. 5I, these studies demonstrate that the impairment of both HR and NHEJ are required for cell death in response to combined MAPK/HDAC inhibition.

The converse experiment was also performed. We reasoned that if cell death was occurring because (i) MAPK pathway inhibitors suppress HR and (ii) MAPK/HDAC inhibitors together suppress NHEJ, then ablation of NHEJ genes should kill cells in the presence of MAPK pathway inhibitors. Individual NHEJ genes were suppressed by pooled siRNA sequences and cells were treated with dabrafenib/trametinib. Suppression of any one of the NHEJ genes killed cells but only in the presence of dabrafenib/trametinib (Fig. 6D; Supplementary Fig. S5A), suggesting that NHEJ pathway suppression was able to recapitulate the effects of entinostat in this context.

Taken altogether, these observations suggest that BRAF/MEK inhibitors suppress DNA repair gene function in the HR pathway, resulting in a state that functionally resembles tumors with BRCA pathway defects (model shown in Fig. 6E; ref. 38). However, when combined with entinostat, these agents now cooperatively and potently suppress a second broader set of DNA repair genes, including NHEJ pathway genes. Importantly, this broad transcriptional repression of these major DNA repair genes is lethal in tumors with a preexisting impairment in a DNA repair gene expression.

To investigate the molecular mechanism(s) by which MAPK and HDAC inhibitors might be cooperatively suppressing DNA repair genes, we first identified transcription factor binding sites that were enriched in genes suppressed by this combination, using the MSigDB Transcription factor targets database. ELK1 binding sites were the most significantly enriched in the suppressed genes and several other ETS family member binding sites were also identified (Fig. 6F). ssGSEA analysis was then used to examine the pattern of ELK1-regulated transcriptional signatures in response to single and combined agents. MAPK suppression partially inhibited these signatures, as expected, given that ERK phosphorylation contributes to the activation of ELK1 (Fig. 6G; ref. 41). HDAC inhibitors slightly suppressed ELK1 signatures; however, these signatures were maximally suppressed in the presence of combined MAPK/HDAC inhibitors (Fig. 6G). We found that ELK1 phosphorylation was inhibited in response to BRAF/MEK inhibitors, as would be predicted (Fig. 6H). However, unexpectedly, both agents individually and cooperatively suppressed total ELK1 protein expression. Microarray data indicated that this was occurring at the level of transcription, which was confirmed by qPCR (Supplementary Fig. S5B). The related gene, ELK3, showed a similar pattern of suppression (Supplementary Fig. S5B).

Twenty percent of the DNA repair genes suppressed by MAPK/HDAC inhibitors have been experimentally determined to be direct ELK1 targets (32/160 genes; ref. 42) and many more of these genes are targets of other ETS family proteins. For practical reasons, we focused on determining whether ELK was controlling the expression of any of the most prominently suppressed group A and B genes (BRIP1, BRCA2, XRCC5, and PARP3). Experimental chromatin immunoprecipitation (ChIP) data indicate that ETS family proteins directly bind sites in the promoters of all four genes, although only BRIP1, XRCC5, and PARP3 contain the ELK1 target sequence (Fig. 6I). Because ELK1 and ELK3 bind the same DNA sequences and therefore may be redundant, we investigated the consequences of genetically ablating ELK1 and/or ELK3. Notably, ELK1 ablation was sufficient to potently suppress the expression of PARP3 and BRIP1 (Fig. 6J), whereas XRCC5 expression was inhibited by the loss of both ELK1 and ELK3 (Fig. 6K). The less dramatic effect on XRCC5 expression, as compared with PARP3 and BRIP1, may be due to the presence of several ETS family binding sites in its promoter (Fig. 6I). Nevertheless, these observations demonstrate that ELK1 critically regulates the expression of these important HR and NHEJ proteins.

More importantly, however, we found that RNAi-mediated suppression of ELK target genes cooperates with either MAPK or HDAC inhibitors to kill these melanomas (Fig. 6L), consistent with the observation that ELK regulates both types of genes (e.g., HR and NHEJ; Fig. 6I). The effects of ELK suppression were more pronounced when combined with MAPK inhibitors versus HDAC inhibitors, as might be expected, because it is likely that other MAPK pathway targets are important in BRAF-mutant melanomas. Growth arrest triggered by ELK1 overexpression precluded rescue experiments. However, the observation that (i) ELK activity and expression is potently suppressed in drug-treated cells, (ii) BRIP1, PARP3, and XRCC5 (and many other DNA repair genes) are direct ELK1 targets, (iii) ELK suppression substantially inhibits the expression of these genes in melanomas, and (iv) ELK ablation can partially recapitulate the effects of both drugs, all support the conclusion that ELK critically regulates this DNA repair network in response to HDAC/MAPK inhibitors. Other ETS family members may play a cooperative role in this process as well. Taken together, these findings provide a mechanistic basis for understanding the cooperativity of MAPK and HDAC inhibitors on DNA repair pathways and cell death.

Using multiple animal models of BRAF-mutant melanoma, we have shown that the Class I HDAC inhibitor entinostat dramatically enhances the efficacy of BRAF/MEK inhibitors, in both tumors that are sensitive and relatively resistant to these agents. In addition, we have uncovered a strategy for treating NRAS- and NF1-mutant tumors, for which there are currently no approved targeted therapies. Finally, we have elucidated the mechanism by which these agents function and have discovered that a clinically available biomarker can be used to select patients that are most likely to benefit from MAPK/HDAC inhibitor combinations. Together, these findings provide a promising and readily translatable strategy for improving treatments for these RAS/RAF pathway–driven melanomas.

Interestingly, in the course of this work, we also discovered a population of melanomas that exhibit unconventional defects in DNA repair. Mutations in BRCA1 or BRCA2 represent examples of classic cancer-associated DNA repair defects, and confer sensitivity to agents that stall replication forks, such as PARP inhibitors (31, 43). However, additional alterations have been proposed to phenocopy these mutations, resulting in a state sometimes referred to as “BRCAness” (discussed in ref. 38). Examples of this include mutations in other HR genes, hypermethylation of BRCA genes, amplification of genes that disrupt HR, and mutations in the transcriptional regulator CDK12. Here we observe a related but more complex paradigm. “Sensitive” melanomas exhibit measureable defects in HR genes due to the transcriptional suppression of a broad group of DNA repair genes (29). Nevertheless, these defects are not potent enough to reach a functional threshold that confers sensitivity to PARP inhibitors alone. Intriguingly, BRAF/MEK pathway inhibitors unmask these defects, by further reducing the expression of several HR/BRCA pathway genes, thereby triggering a BRCA-like state (38). Accordingly, these studies reveal a new type of “priming” defect in melanomas. Moreover, they demonstrate that in this setting, BRAF/MEK inhibitors can broadly suppress the expression of DNA repair genes, most notably in the HR/BRCA pathways, thereby creating a new therapeutic vulnerability.

Importantly, we show that combined suppression of BRAF/MEK and HDAC3 capitalizes on this vulnerability. Whereas BRAF/MEK pathway inhibitors suppress one set of DNA repair genes, most notably HR pathway genes, the addition of entinostat induces a potent suppression of a broader set of genes including essential NHEJ genes. It should be noted that these effects are not due to an additive effect of MAPK and HDAC inhibition, but rather these agents cooperate by potently suppressing a broad transcriptional network of DNA repair genes. We have shown that key nodes of this network are regulated by ELK1, which is cooperatively suppressed by combined MAPK/HDAC suppression, via effects on phosphorylation and expression. Notably, ELK1 has been shown to directly bind and regulate 20% of the DNA repair genes that are suppressed by these agents, suggesting a broader role for ELK1 in maintaining this network. Other ETS family members may also contribute to this response.

It is well established that the NHEJ pathway can compensate for defects in HR. As such, synthetic lethal interactions between HR genes and NHEJ genes have been observed (38, 44). Here we have discovered a chemical synthetic lethality between BRAF/MEK inhibitors and Class I HDAC inhibitors, which when combined coordinately, suppress these pathways. Importantly, we have experimentally demonstrated that suppression of both HR and NHEJ pathways is required for cell death.

Although the molecular basis for the priming defect in melanomas is not known, it could be related to an epigenetic event or state, or alternatively caused by specific defects in transcriptional-regulatory proteins. Fortunately, however, this defect consistently segregates with MGMT expression, which has already proven to be a tractable clinical biomarker in other settings (25, 26). In this respect, it is interesting to note that there are two clusters of DNA repair genes that show inverse expression patterns; a broad set of genes that are suppressed in MGMT⁺ tumors, and a smaller set of genes, several of which are also involved in directly repairing base modifications, that are high in MGMT⁺ tumors (and vice versa). This observation further supports the notion that these gene clusters are coregulated in melanomas. Regardless, the identification of this potent therapeutic combination, coupled with the elucidation of its mechanism of action and a tractable predictive biomarker of efficacy, will enable new mechanism-based combination trials that have the potential to improve treatments for these three major melanoma subtypes.

All cell lines were purchased from ATCC, except for MALME3M (obtained from Levi Garraway, Dana-Farber Cancer Institute, Boston, MA), Meljuso (obtained from William Hahn, Dana-Farber Cancer Institute, Boston, MA), SUM149PT (obtained from Frank McCormick, University of California, San Francisco, CA), Yugasp and Yudoso (obtained from Yale Dermatology Center, New Haven, CT), and WM3670 and WM3629 (obtained from Rockland Immunochemicals). No further authentication of these cell lines was performed. All of the cell lines were determined to be negative for Mycoplasma using the MycoAlert Mycoplasma Detection Kit (Lonza, LT07-318). Cells were used for experiments within 15 to 20 passages from thawing. Antibodies were obtained from the following sources: Cell Signaling Technologies: pERK (4370), ERK (9102), GAPDH (2118), BRIP1 (4578), XRCC5 (2753), H3K56Ac (4243), HDAC1 (5356), HDAC2 (5113), HDAC3 (3949), HDAC6 (7612), RAD51 (8875), pELK1 (9181), HA (3724), cleaved PARP (9541); Sigma Aldrich: Actin (A2066); EMD Millipore: H3K9Ac (06-942), BRCA2 (OP95), pH2AX (05-636); Santa Cruz Biotechnology: XRCC4 (365055), XRCC6 (17789), PARP3 (390771), EME1 (53275); Bethyl Laboratories: RBBP8 (A300-488A-M); Novus Biologicals: PNKP (NBP1-87257); BD Pharmingen: MGMT (557045); Abcam: ELK1 (32106), LIG4 (26039). Trametinib, dabrafenib, and olaparib were purchased from LC laboratories. Vorinostat, mocetinostat, nexturastat, and entinostat were purchased from Selleck Chemicals. 5-azacytidine, NAC, Vitamin C, and BSO were purchased from Sigma-Aldrich. Carboxy-H2DCFDA was purchased from Life Technologies (#C400).

Nontargeting, HDAC1, HDAC2, HDAC3, HDAC6, XRCC4, XRCC5, XRCC6, PARP3, PNKP, BRCA2, MGMT, ELK1, and ELK3 siRNA pools were purchased from GE Healthcare/Dharmacon (D-001810-10, L-003493-00, L-003495-02, L-003496-00, L-003499-00, L-004494-00, L-010491-00, L-005084-00, L-009297-00, L-006783-00, L-003462-00, L-008856-01, L-003885-00, and L-010320-00, respectively). siRNAs were transfected into cells using RNAiMax Lipofectamine from Invitrogen. Cells were transfected approximately 24 hours before starting proliferation experiments. Control, HDAC1, HDAC2, HDAC3, and HDAC6 shRNAs were purchased from Sigma-Aldrich (SHC016, SHCLNG-NM_004964, SHCLNG-NM_008229, SHCLNG-NM_010411, and SHCLNG-NM_006044, respectively).

cDNA clones were obtained from Sino Biologicals (MGMT) and the PlasmID Repository at Harvard Medical School (LIG4 and RAD51), sequence verified and subsequently cloned into the pHAGE-FLAG-HA mammalian lentiviral expression vector.

For the initial MAPK/HDACi screen and the dose–response matrix assay, cells were plated in 96-well plates. Five replicates were done for each condition. At 24 hours, one plate of cells was frozen (−80°C) representing the time 0 plate. At this time, compounds were added to the remaining plates. After 72 hours, each of the plates was frozen. After freezing, the plates (day 0 and 3) were thawed simultaneously and cells were quantified using CellTiter-Glo (Promega) as per the manufacturer's instructions. SynergyFinder (45) was used to analyze drug combination dose–response matrix data. To determine the combination effects in excess of Loewe additivity, a Synergy Score was calculated to characterize the strength of synergistic interaction. For all other proliferation experiments, cells were seeded in triplicate into 6-well plates. Approximately 24 hours after plating, day 0 counts were taken using a hemocytometer. For inhibitor experiments, drug treatments were started at this time. Final cell counts were taken 72 hours after day 0 to determine changes in cell number versus day 0. For western blots to determine drug efficacy or efficient knockdown, lysates were collected 48 hours after the addition of inhibitors.

The live cell imager IncuCyte ZOOM (Essen Biosciences) was used for multiplexed measurements of cell proliferation alongside cell death in a single well. The mKate2 red fluorescent protein (RFP; Essen Biosciences) was transfected into the Hs695T melanoma cell line and selected for RFP-infected cells using puromycin. Successful insertion of the nuclear-restricted RFP was confirmed via direct visualization of transfected cells on the IncuCyte ZOOM fluorescent microscope. For proliferation experiments, stably transfected cells were plated in 96-well plates and allowed to settle overnight at 37°C. The green IncuCyte Cytotox or Caspase-3/7 Apoptosis reagents were then added to the tissue culture growth media to assess real-time quantification of cell death. Red and green fluorescent objects were monitored in the IncuCyte ZOOM acquiring images every 2 hours (for 72 hours) after treatment with vehicle or drugs and then quantified with the IncuCyte integrated analysis software. Relative cell death was defined as the ratio between cells with overlapping red and green cells (dead cells) and cells with red nuclei only (live cells).

Animal procedures were approved by the Center for Animal and Comparative Medicine in Harvard Medical School (Boston, MA) in accordance with the NIH Guide for the Care and Use of Laboratory Animals and the Animal Welfare Act. For cancer cell xenograft experiments, immunodeficient Nu/Nu (A375 and Hs695T) or NSG (YUDOSO) mice were inoculated subcutaneously with 3 × 10⁶ human BRAF- or NRAS-mutant melanoma cells. For allograft experiments, immunocompetent C57BL/6 mice were inoculated subcutaneously with 1.5 × 10⁵ murine Braf/Nf1-mutant melanoma cells or 4 × 10⁵ murine Braf/Pten-mutant melanoma cells. Tumor volumes were calculated by measuring length and width of the lesions and using the formula [(length) × (width)² × 0.52]. When tumors reached approximately 200–300 mm³, mice were randomly divided into different treatment groups that were administered either the MEK inhibitor trametinib (0.6 mg/kg once daily, orally), the BRAF inhibitor dabrafenib (30 mg/kg once daily, orally), the HDAC inhibitor entinostat [1 mg/kg once weekly, intraperitoneally (i.p.)], the PARP inhibitor olaparib (10 mg/kg, once daily, i.p.), or their combinations as indicated. To track changes in tumor volume, tumor size was measured at day 0 and subsequently every 3–5 days by Vernier calipers. Unpaired two-tailed t tests with unequal variance were used to compare datasets where indicated, and P values are shown.

MGMT methylation analysis was performed using the OneStep qMethyl kit (Zymo Research, D5310). Briefly, DNA from 7 different melanoma cell lines was digested with methylation-sensitive restriction enzymes in the test reaction while DNA in the reference reaction was not. After digestion, DNA from both samples was amplified using real-time PCR in the presence of fluorescent dye and quantified. Investigator was blinded to group allocation and when assessing the outcome. The following MGMT primer sequences were used: forward: 5′-GCGCTCTCTTGCTTTTCTCA-3′; reverse: 5′-CTGCAGGACCACTCGAGG-3′.

Melanoma cells transfected with control (siLuc) or siRNA pools against BRCA2 were irradiated with 10 Gy. Cells were fixed 5 hours later and analyzed by immunofluorescence for RAD51 foci and γH2AX.

Cells were treated as indicated and then stained with H2DCFDA at 10 μmol/L and analyzed by flow cytometry.

Live cells were separated in different phases of the cell cycle for subsequent gene expression analysis. Cells treated with vehicle or drugs were incubated with 10 μg/mL Hoechst 33342 for 1 hour at 37°C in the dark. After trypsinization, cells were resuspended at a concentration of 1 × 10⁷ cells per mL for sorting. Cells were sorted on the basis of the amount of DNA by defining two regions for sorting: one for G₁ and the other for G₂–M. The separated cells were collected in RNAse-free conditions and further processed for RNA extraction.

RNA was isolated from Hs695T, SKMEL2, and A2058 cells 24 hours after treatment with indicated drugs. Prior to RNA extraction, ERCC spike-in control mix was added to TRIzol and RNA isolation was performed following the manufacturer's protocol. RNA clean-up was performed using the Qiagen RNeasy kit (74201). The Molecular Biology Core Facility at Dana-Farber Cancer Institute (Boston, MA) hybridized RNA to the Affymetrix Human 2.0 ST array chip. Analyses used BRB-ArrayTools developed by R. Simon and the BRB-ArrayTools Development Team. Thresholds were set at P = 0.001. Microarray data can be accessed from the Gene Expression Omnibus (GEO) database (accession number: GSE125565). GSEA and ssGSEA were performed using the Broad Institute interface (http://software.broadinstitute.org/gsea/index.jsp). The signature of HR-associated genes (PENG_HRD_SIGNATURE) was developed from genome-wide transcriptome profiling of HR DNA Repair (29) and added to the KEGG signature file from MSIGDB for subsequent analysis (http://software.broadinstitute.org/gsea/msigdb/genesets.jsp?collection=CP:KEGG). Tumor sample transcriptional profiles were obtained from the TCGA SKCM provisional dataset (http://firebrowse.org/?cohort=SKCM) and stratified on the basis of median-centered z-scores of MGMT mRNA expression. The top 10% of MGMT expressors comprised the “MGMT⁺” cohort, whereas the bottom 10% of MGMT expressors comprised the “MGMT⁻” cohort used in GSEA. Melanoma cell line transcriptional profiles were obtained from CCLE (https://portals.broadinstitute.org/ccle/home) and stratified on the basis of gene-centric robust multiarray average–normalized MGMT mRNA expression. The top 10% of MGMT expressors comprised the “MGMT⁺” cohort, whereas the bottom 10% of MGMT expressors comprised the “MGMT⁻” cohort used in GSEA analyses. Experimental ChIP data were obtained from the ChIP-X database (http://amp.pharm.mssm.edu/Harmonizome/).

For quantitative measurements, graphs represent mean values ± SD. Where indicated, the data are presented as log₂ fold (left axis) and percent change (right axis) over initial measurements. Unpaired two-tailed t tests with unequal variance were used to compare datasets where indicated and P values are shown. Data were graphed and analyzed using GraphPad Prism v.6.

All proliferation studies in Figs. 1, 3, 4, 5 and Supplementary Figs. S1 and S3 were performed ≥ 3 times. Synergy studies were performed in five cell lines at different times with qualitatively similar findings. The large preclinical animal studies were performed one time; however, qualitatively similar results were obtained in 5 different animal models and responses mimicked effects observed numerous times in vitro. The γH2AX experiment (Fig. 4D) is shown in three cell lines and has been repeated in a separate set of cell lines. The microarray experiment was performed once (3 technical replicates) in three different cell lines and key targets were confirmed by qPCR and western blot analysis (n > 3) as shown throughout Figs. 5 and 6.

L.A. Garraway is a senior vice president, Oncology Research, at Eli Lilly and Company, was previously affiliated with Howard Hughes Medical Institute, was an equity consultant at Foundation Medicine, reports receiving commercial research grants from Novartis and Astellas, and has ownership interest (including stock, patents, etc.) in Tango Therapeutics. K.T. Flaherty reports receiving commercial research grants from Novartis and Sanofi, reports receiving other commercial research support from Novartis, Genentech, Array BioPharma, and Incyte, has ownership interest (including stock, patents, etc.) in Clovis Oncology, Loxo Oncology, Fog Pharma, Tvardi, Checkmate Pharmaceuticals, Strata Oncology, X4 Pharmaceuticals, Apricity, Vivid Biosciences, PIC Therapeutics, Fount Therapeutics, Shattuck Labs, and Oncoceutics, and is a consultant/advisory board member for Loxo Oncology, Strata Oncology, Aeglea, Array BioPharma, Shattuck Labs, Arch Oncology, Tolero Pharmaceuticals, Apricity, Oncoceutics, Fog Pharma, Neon Therapeutics, Tvardi, Vivid Biosciences, Novartis, Genentech, Bristol-Myers Squibb, Merck, Takeda, Verastem, Checkmate Pharmaceuticals, Boston Biomedical, Pierre Fabre, Cell Medica, PIC Therapeutics, Debiopharm, X4 Pharmaceuticals, Sanofi, Amgen, Asana, Adaptimmune, and Fount Therapeutics. S.J. Elledge is a consultant/advisory board member for CRISPR Therapeutics, Homology Medicines, Maze Therapeutics, X-CHEM, MPM Capital, and Tenaya Therapeutics; and is a founder of Miramus and Maze Therapeutics. K. Cichowski is a consultant/advisory board member for Genentech. No potential conflicts of interest were disclosed by the other authors.

Conception and design: O. Maertens, S.J. Elledge, K. Cichowski

Development of methodology: O. Maertens, T. De Raedt

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): O. Maertens, C.E. Emerson, A.G. Gavin, T.C. Wong, T. De Raedt, L.A. Garraway, K.T. Flaherty, S. Pathania

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): O. Maertens, R. Kuzmickas, H.E. Manchester, C.E. Emerson, A.G. Gavin, T.C. Wong, T. De Raedt, K.T. Flaherty, S. Pathania, S.J. Elledge, K. Cichowski

Writing, review, and/or revision of the manuscript: O. Maertens, R. Kuzmickas, C. Bowman-Colin, E. Hatchi, K.T. Flaherty, S.J. Elledge, K. Cichowski

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): H.E. Manchester, C.E. Emerson, C.J. Guild

Study supervision: L.A. Garraway, S.J. Elledge, K. Cichowski

This work was supported by grants from the NCI (R01CA111754, to K. Cichowski), the Ludwig Center at Harvard (to K. Cichowski and S.J. Elledge), and the NF Research Initiative at Boston Children's Hospital made possible by an anonymous gift (to O. Maertens).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.