Prediction of DNA Repair Inhibitor Response in Short Term Patient-Derived Ovarian Cancer Organoids

Most HGSCs exhibit functional HR repair.

The HR capacity of the organoid cultures was assessed using multiple surrogate markers. First, all organoid cultures were tested for sensitivity to the PARPi olaparib (Figure 2, Table 1), since olaparib sensitivity is a useful surrogate marker for an HR defect (7). They were also tested for sensitivity to replication fork stalling agents, including carboplatin, as well as the CHK1 inhibitor prexasertib and ATR inhibitor VE-822. These latter agents are known to induce replicative stress. A subset of organoid cultures were also tested for sensitivity to the replication stress inducing nucleoside analog gemcitabine, and to the conventional HGSC agents doxorubicin and paclitaxel. To ensure that the drug concentrations applied to the organoids could kill ovarian cancer cells, these agents were initially tested in the OVCAR8 cell line transfected with a control or BRCA1-specific siRNA, and shown to cause the expected dose-dependent cytotoxicity (Figure S4A, S4C). Sensitivity standards were established for each drug (Figure S5-S8), and the sensitivity results for all 34 lines are reported in Table 1.

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Figure 2.

Most HGSC organoids are HR proficient and lack therapeutic sensitivity to agents targeting HR defects.

A) Sensitivity dose curves of organoid cultures from one tumor site (transverse colon mesentery) from a BRCA1 mutation carrier (DF-17–39) with acquired PARPi resistance to carboplatin, olaparib, prexasertib, and VE-822. A dashed black line marks 50% untreated and a dashed grey line marks our sensitivity standard for this assay for all organoid cultures. S= sensitive and R=resistant. B) RAD51 focus formation 4 hours post 10 Gy in DF-17–39 transverse colon mesentery metastasis organoids. The top left panel is an H&E stain of the organoids, the top right panel shows cells in S phase marked by Geminin, the middle left panel marks DNA damage with γH2AX, the middle right panel shows the presence of RAD51 foci, and the bottom panels are magnified areas of the RAD51 stain. C) Sensitivity dose curves of organoid cultures generated from an omental metastasis from an untreated HGSC patient, DF-17–126, to carboplatin, olaparib, prexasertib, and VE-822. The dose curves are configured as described for panel A. D) RAD51 focus formation 4 hours post 10 Gy in DF-17–126. The top left panel is an H&E stain, the top right panel shows cells in S phase marked by Geminin, the middle left panel marks DNA damage with γH2AX, the middle right panel shows a lack of RAD51 foci, and the bottom panels are magnified areas of the Rad51 stain.

Second, the ability of organoid tumor cells to assemble RAD51 foci, either pre- or post- irradiation (IR), was tested. In the process of HR, RAD51 is loaded onto the ends of a double-strand break (DSB), allowing the resected DSB to invade the sister chromatid (6). The assembly of RAD51 foci is a surrogate marker for the ability of a cell to perform HR, up to the stage of RAD51 loading (6). This assay does not measure HR steps downstream of RAD51 loading. Organoid cultures were treated with 0 or 10 Gy, prior to assaying for RAD51 foci (23). Organoids were co-stained for γH2AX to mark DNA damage and for geminin to mark cells in S phase (Figure 2) (23). Stained slides were examined first for geminin positivity and subsequently for RAD51 foci. The detection of 1–3 cells over multiple high-power fields with RAD51 foci was scored as positive, and the corresponding tumor was scored as HR competent (Figure 2, Table 1) (23). For RAD51 positive organoids, RAD51 foci and extensive γH2AX nuclear positivity were detected both pre- and post-IR in most cases.

Organoids from two patients illustrate the utility of the HR functional assays (Figure 2). Organoids were established from four tumor sites from a BRCA1 mutation carrier whose tumor had acquired PARPi resistance (Patient DF-17–39, Figure 2A and ​and2B,2B, Table 1). All organoids were olaparib resistant, but sensitive to carboplatin, prexasertib, VE-822, and gemcitabine (Figure 2A, Table 1). Interestingly, the functional results for this patient matched the clinical response. The patient was later treated with prexasertib and exhibited stable disease at sites from which organoids had been derived. Subsequent treatment with carboplatin and gemcitabine resulted in a decrease in disease burden for several months (Table S4). The patient subsequently recurred and succumbed to her disease, suggesting acquired resistance to these agents. The organoids from all four sites exhibited RAD51 foci (Figure 2B, Table 1), indicating HR competence, consistent with olaparib resistance. Despite PARPi resistance in the organoids and in the patient’s tumor, these organoids remained carboplatin sensitive, a surrogate marker for a defect in stalled replication fork protection. These results confirmed that the HR and fork repair functions of BRCA1 are independent (24,25) and that the molecular mechanism of PARPi resistance restored HR but not stalled fork protection.

In contrast to patient DF-17–39, organoids generated from a patient carrying a RAD51C/FANCO germline mutation (DF-17–126), were sensitive to olaparib, suggesting an HR defect, and were also sensitive to carboplatin, prexasertib, VE-822, and gemcitabine (Figure 2C). Consistent with this olaparib sensitivity, these organoids exhibited no RAD51 focus formation, demonstrating that the tumor was indeed HR deficient (Figure 2D, Table 1).

Only two of thirty-four (6%) of the organoid cultures tested were olaparib sensitive (Figure 2, Table 1) suggesting that organoids derived from most HGSCs do not have functional HR defects, despite their genetic findings, and predicting that the corresponding tumors will be insensitive to PARP inhibition. The overall rate of carboplatin, prexasertib, VE-822 and gemcitabine sensitivity was much higher (41%, 47%, 44%, and 82%, respectively), and did not correlate with PARPi sensitivity, indicating that a functional HR defect may not be required for response to these agents. A recent study suggests that replication stress defects may be a better predictive biomarker of CHK1 inhibitor response (13). For doxorubicin and paclitaxel, the overall sensitivities were 45% and 82%, respectively and also did not correlate with an HR defect. Overall, these data suggest that most organoids derived from sporadic HGSC tumors are HR proficient and PARPi-resistant.

Correlation of tumor mutational status with organoid functional testing.

The low percentage of organoids responding to PARPi (2/23 patients, 9% of patients tested) is surprising, given the high number of patients (up to 50%) hypothesized to have HR defective tumors by genomic analysis (4,10). However, when WES data from tumor-organoid pairs were further evaluated, reasons underlying the low response rate were clarified. We queried multiple aspects of genomic HR markers including somatic mutations, copy number variation, allelic imbalance, and mutational signatures (Figure 3, Table S3, Table S5).

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Figure 3.

Mutational signatures and repair gene mutation status in tumors and organoid cultures.

A) WES data from parent tumors and organoid cultures were analyzed for numbers of SNVs and long deletions to assess for HRD mutational signatures. Deletions per megabase (Mb) compared to fraction of deletions 5-bp or longer (top panel) and fraction of C to T substitutions (C > T) and SNVs per Mb (bottom panel) for each parent tumor and organoid culture are shown for all 34 patients. The organoids and parent tumors harboring components of the HRD mutational signature have a black box enclosing them in each panel. A color code for each patient is at the top of the panels. B-G) Germline and tumor allele mutation status and repair assay status for B) RAD51C/FANCO for DF-17–126. C) BRCA2 for patient DF-18–30. D) BRCA1 for patient DF-17–107. E) BRCA1 for patient DF-17–39. F) BRCA2 for patient DF-17–115. G) BRCA2 for patient DF-18–23.

First, we assessed the parent tumor and organoid WES data for a homologous recombination defect (HRD) mutational signature (26), which is defined by somatic single nucleotide variants (SNVs), as well as insertion-deletion (indel) mutations. This signature is comprised of an elevated rate of both SNVs and deletions, as well as context-specific depletion of C>T substitutions and increased long deletions. This mutational signature was readily detected and quantified in tumors and organoids from five of the six patients with germline HR gene mutations (DF-17–39, DF-17–115, DF-17–126, DF-18–23, and DF-18–30), weakly detected in a patient with a germline variant of unknown significance in BRCA1 (DF-18–8), and not detected in sporadic patients (Figure 3A, Table S5). Components of the signature were detected in some sporadic tumors such as DF-18–7 which showed increased long deletions only, or DF-17–123 which weakly demonstrated the signature for SNVs only. Both of these cases with only parts of the HRD signature also lacked PARPi response, suggesting that residual or restored HR was present, that the signature components were only associated with impaired protection of stalled replication forks, or that the observed mutations were due to some other, non-HR, mutational mechanism. We were able to clarify these possibilities with an assay for replication fork stability (see below) (Table 1).

The HRD mutational signature was strongest in patient DF-17–126, the patient with a germline mutation in RAD51C/FANCO, an ovarian cancer susceptibility gene required for HR repair (27,28). The DF-17–126 parent tumor and organoid culture had a high number of long deletions and SNVs, consistent with the HRD signature (Figure 3A, Table S5). Examination of the copy number status of the RAD51C/FANCO locus at the site of the patient’s known mutation revealed two copies and increased allelic imbalance, in favor of the mutant allele (Figure S9A), revealing copy neutral loss of heterozygosity (LOH) of the wildtype allele in both the parent tumor and organoid culture (Figure 3B). These genomic data correlated with the olaparib sensitivity and lack of RAD51 foci in the DF-17–126 organoid culture (Figure 2C and ​and2D,2D, Figure 3B, Figure S9A). Taken together, the genomic HR data combined with the functional data of the organoids strongly predict a functional HR defect in the tumor.

The HRD mutational signature was also strong in DF-18–30, a patient with a germline BRCA2 mutation who developed a carcinosarcoma (Figure 3A, Table S5). Examination of the copy number status of the BRCA2 locus indicated single copy loss and increased allelic imbalance in favor of the mutant allele (Figure S9B), thus revealing LOH of the wildtype allele (Figure 3C). Again, the genomic data correlated with the olaparib sensitivity and lack of post-IR RAD51 foci in the DF-18–30 organoid culture (Figure 3C, Table 1). This patient is currently responding to standard of care carboplatin/paclitaxel treatment with an ongoing drop in her CA125 (Table S4).

In contrast, patient DF-17–107 was a germline carrier of the p.C61G founder mutation in BRCA1. A logical prediction is that this patient’s tumor arose from LOH of the wildtype BRCA1 allele and would be olaparib sensitive. However, the tumor-derived organoids formed RAD51 foci and were olaparib resistant (Table 1), indicating that the tumor was HR proficient. Tumor-organoid pairs from this patient did not exhibit an HRD signature (Figure 3A, Table S5). Copy number interpretation of this patient’s samples was impaired by low tumor purity. Re-examination of the germline variant at the BRCA1 locus in the tumor and organoid lines revealed that only the left ovarian tumor site had mild allelic imbalance and two copies, suggesting at least clonal or subclonal copy neutral LOH in favor of the mutant allele, whereas the right ovary mass and omental metastasis did not exhibit loss of the wildtype allele (Figure S3 and Figure S9C). Taken together, these results suggest that all tumor sites likely express some wildtype BRCA1 (Figure 3D). These findings are consistent with the functional assay results and absent HRD signature (Table 1). BRCA1 mutation carriers without loss of the wildtype allele and with intact HR, based on genomic analysis, have been observed previously (29). In addition, the p.C61G mutation has been shown to be sufficient for tumorigenesis but not for platinum or PARPi sensitivity, suggesting that even in the absence of wildtype protein expression, this hypomorph may have functional DNA repair activity (30).

The PARPi-resistant BRCA1 mutation carrier (DF-17–39) profiled in Figure 2A and ​and2B,2B, exhibited the HRD signature, but the magnitude was lower than that observed for DF-17–126 (Figure 3A, Table S5). Each of the tumors and organoids from this patient demonstrated a single copy loss and increased allelic imbalance in favor of the mutant variant of BRCA1 (Figure S9D), consistent with LOH and rendering the tumor BRCA1-deficient (Figure 3E). However, at the time of organoid generation, the patient was clinically resistant to olaparib (Table S4), and the organoid cultures were olaparib-resistant and competent for RAD51 focus formation (Figure 2A and ​and2B,2B, Table 1). No genetic cause for the evolution of the PARPi resistance was discovered. Thus, while genetic analysis suggested that the tumor was HR defective, the organoid functional assays and clinical response revealed that the tumor was PARPi-resistant.

The BRCA2 germline mutation carrier (DF-17–115) also exhibited the HRD signature (Figure 3A, Table 1, Table S5). The parent tumor and organoid cultures showed single copy loss and increased allelic imbalance in favor of the mutant germline variant in the BRCA2 locus (Figure S9E), consistent with LOH of the wildtype allele (Figure 3F). Current treatment selection protocols assume that this would result in an HR-defective, PARPi-sensitive tumor. However, these organoids were PARPi resistant but carboplatin, prexasertib, and VE-822-sensitive, suggesting that the mechanistic defect conferred by this BRCA2 mutation was a stalled fork protection defect and not an HR defect. These results further support the need for a combined genetic and functional assay approach to targeted therapy selection.

Finally, the tumor from the neoadjuvantly-treated BRCA2 germline mutation carrier (DF-18–23) also exhibited the HRD signature (Figure 3A, Table 1, Table S5). The parent tumor and organoid cultures showed single copy loss and increased allelic imbalance in favor of the mutant germline variant in the BRCA2 locus (Figure S9F), consistent with loss of heterozygosity of the wildtype allele (Figure 3G). Current treatment selection protocols predict an HR defective, PARPi-sensitive tumor. However, these organoids were resistant to PARPi, carboplatin, prexasertib, and VE-822, although they lacked RAD51 foci. This case is particularly interesting clinically. The patient had a protracted treatment course with sporadic periods on and off chemotherapy (Table S4). After neoadjuvant chemotherapy, the patient delayed surgery for seven months, at which time her tumor markers rose and imaging revealed an expanding pelvic mass (Table S4). Organoids were established from this pelvic mass at the time of her interval debulking. Taken together, these clinical data suggest that this site was likely selected for therapy resistance through an unknown mechanism. These results further support the need for a combined genetic and functional approach to targeted therapy.

Organoids generated from the tumor from patient DF-18–8 showed a very weak HRD signature (Figure 3). These organoids were sensitive to carboplatin, prexasertib, and VE-822 but resistant to olaparib. The patient carries a germline variant of unknown significance (VUS) mutation in BRCA1 (c.4550T>C). Examination of the BRCA1 locus at the site of the patient’s known mutation revealed two copies and increased allelic imbalance, in favor of the mutant allele (Figure S9G). These results reveal copy neutral LOH of the wildtype allele in both the parent tumor and organoid culture. However, given that the HRD signature was weak, the tumor had high baseline copy number variation, and the culture was PARPi resistant, the VUS may be functional, highlighting the importance of the functional assays.

Taken together, these cases explain the PARPi insensitivity of ten of the thirty-two resistant organoids in our sample set. To understand the basis of the other twenty-two resistant organoids, we examined the WES data for somatic mutations or copy number alterations. No obvious mutations or alterations in HR pathway genes were found (Table S3), suggesting that these organoids would be expected to be resistant. Organoids with somatic mutations in BRCA pathway genes (for example, the DF-17–104 tumor harboring a RAD51C somatic mutation) were identified with a partial HRD signature (Figure 3A); however, ultimately this line was PARPi resistant, showing that a functional assay is more predictive than a single somatic mutation.

Although the low rate of PARPi sensitivity is surprising, our results are more accurate than genetic predictions alone. Clinical trials have suggested that some ‘sporadic’ HGSC cases have PARPi response (31,32), however, the stronger responses occur mostly in BRCA or other repair gene mutation carriers with monotherapy (8,33) or in combination with other therapies (34,35). Even many of these patients do not respond (31,32,36). Larger ongoing trials will determine the response rate in true sporadic patients (37). Still, the organoids in our study functionally match their genomic status, and in the cases where clinical data are available (DF-17–39 and DF-17–121), also match the clinical lack of response (Table S4).

WES data combined with the functional assays suggest that even if a patient harbors a germline mutation in an HR gene and exhibits copy number loss of the wildtype allele or an HRD mutational signature, the tumor cells may not be sensitive to therapies predicted by the mutation (38). The magnitude of the HRD signature reflects the history of a functional defect in a tumor. The most accurate assessment of current defects requires functional assays combined with the germline and somatic genomic data.

Replication fork instability correlates with sensitivity to agents targeting fork protection defects.

BRCA1 and BRCA2 have additional roles in the protection of stalled replication forks, discrete from their functions in HR (11,24,25,39). We next tested the organoids for their ability to protect stalled replication forks, using the DNA fiber assay (40). Organoids were sequentially pulsed for equal time periods with two nucleotide analogues, CldU and IdU, followed by exposure to the fork stalling agent hydroxyurea (HU). If the ratio of the track lengths is one, then the fork was protected during replication stalling. If the ratio of the track lengths is less than one, then the fiber containing the second analogue was degraded, indicating that the tumor cell is unable to protect its stalled forks.

Each organoid culture was analyzed by the fiber assay. The organoids were also tested for sensitivity to the fork stalling agent carboplatin, a front-line chemotherapeutic agent for HGSC (Figure 4, Table 1) (3), as well as, for sensitivity to prexasertib and VE-822, in parallel with olaparib. In some cases other chemotherapy agents, including gemcitabine, paclitaxel, and doxorubicin were examined. HU and other drug treatments were validated in the fiber assay in OVCAR8 cells (Figure S4B, S4C).

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Figure 4.

Replication fork stability correlates with carboplatin sensitivity in HGSC organoids.

A) Sensitivity dose curves (left panel) and fiber assay results (right panel) of omental metastasis organoid cultures from a sporadic HGSC patient (DF-17–116). Dose curves for carboplatin, olaparib, prexasertib, and VE-822 show sensitivity compared to the untreated control. S stands for sensitive and R stands for resistant. A dashed black line marks 50% untreated and a dashed grey line marks the sensitivity cutoff for all organoid cultures. On the right, the ratio of IdU to CldU in three biologic replicates is shown for DF-17–116 organoids treated with hydroxyurea. A black line marks a ratio of 1, and a grey line marks the average ratio for this line. At the top of the panel is a representative fiber from this line denoting an unstable fork. B) Sensitivity dose curves (left panel) and fiber assay results (right panel) of organoid cultures from a sporadic HGSC patient post-neoadjuvant chemotherapy (DF-17–132). Dose curves for carboplatin, olaparib, prexasertib, and VE-822 show sensitivity compared to the untreated control. The graphs are designed as described in A. On the right, the ratio of IdU to CldU in three biologic replicates is shown for organoids treated with hydroxyurea. A black line marks a ratio of 1, and a grey line marks the average ratio for this line. At the top of the panel is a representative fiber from this line denoting a stable fork.

As an example of a tumor with a defect in stalled fork protection, we show the organoid analysis of a sporadic HGSC patient (DF-17–116). These organoids were sensitive to all therapeutic agents tested, except for olaparib, and had an average fiber track length ratio of less than one over three biologic replicates (Figure 4A), consistent with unprotected (unstable) forks. Of note, the culture demonstrated RAD51 focus formation, consistent with the olaparib resistance. In contrast, the organoid analysis of another sporadic HGSC patient (DF-17–132), after neoadjuvant treatment, provides an example of a tumor which is competent for stalled fork protection. This organoid culture was resistant to all four agents and had a fiber track length ratio averaging one (Figure 4B), consistent with fork stability and functional fork protection. This culture was also positive for RAD51 foci.

While olaparib sensitivity correlates with the absence of RAD51 focus formation, we hypothesized that sensitivity to the other agents correlates with impaired replication fork protection (i.e. replication fork instability). Indeed, carboplatin sensitivity was associated with fork instability and carboplatin resistance was associated with fork stability in our organoid panel (Table 1). Seventeen of twenty-eight (61%) organoid cultures tested exhibited unstable forks, suggesting that this mechanistic defect is more common regardless of genomic status. Of the seventeen fork unstable lines tested for drug sensitivity, thirteen were carboplatin sensitive (76%). In contrast, ten out of eleven organoids (91%) with stable forks were carboplatin resistant.

Replication fork instability was associated with sensitivity to prexasertib, VE-822, and possibly gemcitabine. Twelve of the seventeen (71%) fork-unstable cultures were prexasertib-sensitive and nine of the eleven (82%) cultures with stable forks were prexasertib-resistant. Similarly, twelve of the seventeen (71%) fork unstable cultures were VE-822 sensitive and eight of the eleven (73%) cultures with stable forks were VE-822 resistant. Thus sensitivity to prexasertib or VE-822 may be associated with replication fork instability as determined by the DNA fiber assay. Finally, seven of the fork unstable cultures were tested for gemcitabine sensitivity, and all seven (100%) were sensitive while two of the four (50%) fork stable cultures were resistant.

In contrast to the results with carboplatin, prexasertib, VE-822, and gemcitabine, for olaparib, only two of the seventeen cultures (12%) with unstable forks were sensitive. Although all eleven (100%) of the fork-stable cultures were olaparib-resistant, these cultures were also positive for RAD51 focus formation. Taken together, these data suggest that olaparib sensitivity does not associate with fork instability. Eleven lines were tested for sensitivity to doxorubicin and twelve for paclitaxel, only seven of which had unstable forks. For doxorubicin, three of seven (43%) fork-unstable lines were sensitive, and for paclitaxel six of seven (86%) fork unstable lines were sensitive. Other determinants may govern sensitivity to these agents as well.

Replication fork stability may therefore be a predicator of response or resistance to specific DNA repair agents. Larger numbers of patient-derived organoids generated from patients treated with the drugs of interest, along with fiber assay analysis, will be required to more accurately measure the predictive power of this assay. The five tumor/organoid pairs from patients with germline HR gene mutations and an HRD signature had unstable forks; however, some tumors/organoid pairs from patients with no germline mutation and no HRD signature also had unstable forks, further suggesting diverse mechanisms of fork protection not necessarily linked to HR (Figure 3, Table 1).

Tumor samples for organoid generation were obtained from two patients who underwent laparoscopic biopsy prior to neoadjuvant chemotherapy. For the first patient, DF-17–132, at initial laparoscopic biopsy, the tumor was deemed unresectable, and after neoadjuvant treatment, the patient had only a minimal response to chemotherapy (Table S4). Organoids generated from pre- and post- neoadjuvant treatment tumor samples, exhibited RAD51 foci, stable replication forks in the case of the post-neoadjuvant samples, and resistance to the four agents tested, consistent with the carboplatin resistance of this patient in the clinic (Figure 4B, Table 1, Table S4). For a second patient who underwent laparoscopic biopsy and neoadjuvant chemotherapy prior to organoid generation, DF-18–47, the tumor was also deemed unresectable at the time of laparoscopic biopsy (Table S4). In contrast to patient DF-17–132, however, the tumor-derived organoids from the pre-treatment biopsy showed extreme sensitivity to carboplatin, prexasertib, and VE-822, but resistance to olaparib, and demonstrated unstable forks and RAD51 foci (Table 1). At the time of her interval debulking, the patient had minimal tumor tissue, insufficient for the generation of organoids. Her chemotherapy response was therefore predicted by the organoid culture results (Table S4). Taken together, these cases indicate that organoid functional and therapeutic studies may be predictive of carboplatin resistance and could be used for window-of-opportunity studies in clinical trials for predicting response to other neoadjuvant agents.

Combined prexasertib-mediated inhibition of CHK1 with carboplatin or gemcitabine promotes fork instability.

Prexasertib monotherapy is effective for some patients with sporadic HGSCs, consistent with our sporadic organoid culture results (Table 1) (13). Indeed, twelve of the seventeen fork-unstable organoids (71%) were prexasertib-sensitive, while fork stable organoids tended to be prexasertib resistant (Table 1). These results suggest that prexasertib monotherapy may not be appropriate for all patients and highlight the need for assays to understand functional defects that confer prexasertib sensitivity.

Western blot analysis of thirty-three of the organoid cultures showed that prexasertib increases DNA damage, indicated by increased expression of γH2AX, and increased replication stress, as indicated by increased phosphorylated RPA (pRPA). Prexasertib activates the ATR pathway in both fork-unstable and fork-stable lines, as shown by the increased phosphorylation of the ATR targets KAP1 (pKAP1) and CHK1 (pCHK1) (Figure 5A, Table 1) (41,42). The elevated pCHK1 level is a pharmacodynamic marker of CHK1 inhibition by prexasertib. Prexasertib primes a tumor for sensitivity to other DNA repair agents by blocking the ATR/CHK1 pathway, thereby increasing replication stress (12,41). We reasoned that other agents which amplify replication stress might synergize with prexasertib and promote fork instability and tumor cell death.

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Figure 5.

CHK1 inhibitors cause DNA damage in both sporadic and familial HGSCs and confer fork instability in the setting of carboplatin or gemcitabine treatment.

A) Western blots examining DNA damage in one replication fork unstable and one fork stable organoid line. Lines were treated with control (con) media with no drug or media containing in the top panels the CHK1 inhibitor Prexasertib (Prex) alone, Carboplatin (Carbo) alone, or Prexasertib+Carboplatin for either 3 hours or 14 hours; or in the bottom panels with media containing no drug (con), Prexasertib alone, Gemcitabine (Gem) alone, or Prexasertib+Gemcitabine for either 3 hours or 14 hours and then harvested for western analysis. DNA damage was queried by western blot for phosphorylated KAP1 (pKAP1), phosphorylated RPA (pRPA), and γH2AX. CHK1 expression and DNA damage induced phosphorylation are also shown. Vinculin is used as a loading control. B) Biologic replicates of the fiber assay for fork stability in a line with stable forks (DF-17–134 Left Ovary). In the left panel, the first three replicates show testing of the line using standard hydroxyurea (HU) alone. Next to the HU replicates are two replicates each of the line treated with carboplatin (carbo) alone, prexasertib (prex) alone, and a combination of prexasertib and carboplatin. In the panel on the right are shown two replicates each of the line treated with control media with no drug, prexasertib (prex) alone, gemcitabine (gem) alone, or prexasertib+gemcitabine. For the prex, carbo, prex+carbo, prex, gem, and prex+gem experiments, there is no HU in the media at any step, meaning that any fork instability observed is the result of the single drug or drug combination. A black line marks the average stable ratio, and a grey line marks the average of the prexasertib+carboplatin or prexasertib+gemcitabine biologic replicates.

To test this hypothesis, we treated fork-stable or fork-unstable organoids with a combination of prexasertib plus a replication stress (RS)-inducing drug, such as carboplatin or gemcitabine. By western analysis of the RS-biomarkers pRPA and pKAP1, prexasertib alone induced significant replication stress, whereas carboplatin and gemcitabine alone induced minimal replication stress (Figure 5A). The prexasertib+carboplatin combination led to similar levels of replication stress in both fork stable and unstable lines, compared to prexasertib alone. Interestingly, the prexasertib+gemcitabine combination led to increased replication stress and ATR activation, compared to either agent alone, in both the fork-stable and unstable lines (Figure 5A).

In organoids with stable forks, such as the DF-17–134 left ovary cultures, prexasertib, carboplatin, and gemcitabine alone did not destabilize replication forks, as shown by the DNA fiber assay (Figure 5B, Table 1), however, the combination of prexasertib+carboplatin or prexasertib+gemcitabine enhanced fork instability in these organoids (Figure 5B). The enhanced destabilization of the replication fork by these drug combinations was also evident in other fork-stable organoids (DF-17–134 right ovary and DF-17–121) and in fork-unstable organoids (DF-17–116) (Table 1). In contrast, for fork unstable organoids that had acquired PARPi resistance (DF-17–39), prexasertib+gemcitabine combination induced fork instability while prexasertib+carboplatin combination did not (Table 1).

These results provide a rationale for treating HGSC with prexasertib in combination with other conventional RS-enhancing agents like carboplatin and gemcitabine. The fact that the prexasertib+carboplatin combination does not work well for all fork stable tumors highlights the importance of functional assays in deciphering targetable functional defects. Taken together, these data suggest that proper DNA damage repair drug combinations can be effective in tumors without underlying DNA repair defects. Short-term organoids are a useful model for rapidly testing these combinations.

Overcoming pitfalls of the organoid models

Results from other tumor types have raised concerns about organoid models. First, the variable time required for the generation of organoid lines in culture may lead to in vitro selection of specific tumor clones (14–16). Second, the large number of growth factors added to the organoid media may lead to epigenetic changes and clonal selection. HGSC organoids grew within 7–10 days of plating, and our experiments were performed within 1–2 passages of the organoids and within 1–2 weeks of tissue collection, mitigating the issues of selection over time. WES analysis was performed on organoid cultures during this timeframe, and the organoids did not accumulate somatic mutations. Also, early cultures did not lose any of the major driver somatic alterations in the tumors (Figure S2, Figure S3, Table S2, Table S3) indicating that the cultures maintained the heterogeneity of the parent tumors. Additionally, although ovarian organoid cultures contain a minimal stromal component and do not faithfully mimic all components of the tumor microenvironment, organoids do contain immune cells (20), and may be useful in studying the immune intereaction with the tumor in real time. Therefore, these cultures may help understand how best to utilize immuno-oncologic agents.

Organoid generation and culture:

Pleural effusion fluid was obtained immediately at the time of thoracentesis and was spun at 1000 RPM to create a cell pellet. The pellet was washed two to three times in Red Blood Cell lysis buffer (Biolegend Catalog number 420301), washed once in basal culture media [ (Advanced DMEM/F12 (Thermo Fisher 12634028), supplemented with 1% penicillin streptomycin, 1X Glutamax (Life Technologies Catalog number 35050061), and 1% HEPES (Life Technologies Catalog number 15630080)], and the cell pellet was allocated for WES, histology, and organoid generation. For solid tumors, the tumor was acquired immediately during the surgery. Some fresh tumor was snap frozen for later WES or fixed immediately in 10% formalin for later histologic analysis. Remaining tumor tissue was transported to the laboratory in basal culture media. The tumor was diced into approximately 2 mm sections and then crushed with the butt of a syringe. This homogenate was then poured into basal culture media containing Type II Collagenase (Life Technologies # 17101015) at a final concentration of 2.5mg/mL. The homogenate was shaken at 37°C for no longer than 30 minutes. The homogenate was then diluted 1:1 with basal culture media and filtered through a 70uM filter (Falcon Catalog number 352350). The cell suspension was then spun at 1000RPM to create a cell pellet. The pellet was washed with red blood cell lysis buffer two to three times, and then washed once with basal culture media.

For general culture, once a pellet of either liquid or solid tumor cells was obtained, the cells were mixed with growth factor reduced Matrigel (Corning Catalog number CB-40230C), with the final concentration of Matrigel at 75%, and there were approximately 10,000 or more cells/cell groups per 10uL droplet of Matrigel. The suspension was then rapidly plated into a 48-well plate with 15uL of suspension per well. Once the Matrigel was solidified, 250uL of general culture medium was added to each well. General culture medium was composed of Advanced DMEM/F12, supplemented with 1% penicillin streptomycin, 1X Glutamax, 1% HEPES, 100 ng/mL R-spondin 1 (Peprotech catalog number 120–38), 100 ng/mL Noggin (Peprotech catalog number 120–10C), 50 ng/mL EGF (Peprotech catalog number 100–15), 10 ng/mL FGF-10 (Peprotech catalog number 100–26), 10 ng/mL FGF2 (Peprotech catalog number 100–18B), 1× B27 (Life Technologies Catalog number 17504044), 10 mM Nicotinamide (Sigma Aldrich Catalog Number N0636), 1.25mM N-acetylcysteine (Sigma Aldrich catalog number A9165), 1uM Prostaglandin E2 (Catalog number R&D Systems Catalog number 2296), 10uM SB202190 (Sigma Aldrich Catalog number S7076), and 500nm A83–01 (Sigma Aldrich catalog number SML0788). Y-27632 dihydrochloride (AbMole Bioscience Catalog number M1817) was initially tested in early passage organoid cultures at a concentration of 10uM but eventually deemed unnecessary for culture maintenance. Overall tissue digestion and cell plating were similar to that described in Drost, et al 2016 (14).

We would like to thank the 23 generous patients who donated tissue for this work. This research was supported by a Stand Up To Cancer-Ovarian Cancer Research Fund Alliance-National Ovarian Cancer Coalition Ovarian Cancer Dream Team Translational Research Grant (Grant Number: SU2C-AACR-DT16–15). Stand Up To Cancer is a division of the Entertainment Industry Foundation. Research grants are administered by the American Association for Cancer Research, the Scientific Partner of SU2C. This work was also supported by grants from the U.S. National Institutes of Health (R37HL052725, P01HL048546), the U.S. Department of Defense (BC151331P1), the Leukemia and Lymphoma Society (6237–13), the Breast Cancer Research Foundation, the Richard and Susan Smith Family Foundation, and the Bridge Project, a partnership between the Koch Institute for Integrative Cancer Research at MIT and the Dana-Farber/Harvard Cancer Center (to ADD). Work on RAD51 staining was supported by a Biomarker Supplement to UM1 CA186709 (GIS and ADD). We would like to thank Eli Lilly for providing us with the Prexasertib used for the experiments in this study.