Nanoformulation of Olaparib Amplifies PARP Inhibition and Sensitizes PTEN/TP53-Deficient Prostate Cancer to Radiation

PARP inhibitors are an emerging class of drugs that inhibit PARP-1 and PARP-2, proteins that play a critical role in base excision repair (1). When PARP function is impaired, double-stranded DNA (dsDNA) breaks accumulate with time. Cells deficient in homologous recombination cannot accurately repair these breaks, leading to genetic instability, chromosome rearrangement, and cell death (2). This synthetic lethality has proven effective for improving progression-free survival in patients with homologous repair-deficient BRCA mutations (3). PARP inhibitors have also been shown to trap PARP-1 and PARP-2 on DNA (4), thereby forming PARP–DNA complexes that are thought to be responsible for the synergism seen with PARP inhibition and alkylating agents. There is also growing evidence that PARP inhibitors may provide clinical benefit for subsets of patients proficient in homologous repair (5); however, it is not yet clear how to best select patients for treatment with PARP inhibitors.

PARP inhibitor olaparib (Lynparza, AstraZeneca Pharmaceuticals LP) has been approved by the FDA as a monotherapy for advanced ovarian cancer in patients with germline BRCA mutations (5, 6). Patient eligibility is determined using the BRACAnalysis CDx in vitro companion diagnostic (Myriad Genetics) to identify BRCA1 and BRCA2 gene variants. Olaparib has also been granted Breakthrough Therapy status by the FDA for metastatic castration-resistant prostate cancer (mCRPC) following the phase II TOPARP-A clinical trial in which men with defective DNA damage repair mechanisms responded to olaparib (7). Importantly, this trial demonstrated that mutations in other DNA damage repair genes, not just BRCA, can induce sensitivity to PARP inhibition. Olaparib monotherapy has shown efficacy across a variety of advanced tumors classified by mutation status (5, 7–10), and thus olaparib continues to be investigated in more than 40 active phase II and III clinical trials (11). Notably, 47% of these trials seek to determine whether the therapeutic efficacy of olaparib can be enhanced in combination with other small molecules, including DNA-damaging agents, cytotoxic chemotherapy, anti-angiogenic agents, and molecular-specific inhibitors.

Olaparib has been demonstrated to enhance the cytotoxicity of therapies that induce DNA damage. Oral olaparib capsules, administered in combination with paclitaxel for advanced gastric cancer, improved overall survival for patients with known DNA damage repair impairments (12). Olaparib in combination with paclitaxel and/or platinum therapy enhanced progression-free survival in advanced ovarian cancer, breast cancer, and other solid tumors (13, 14). Unfortunately, these drug combinations produced significant off-target toxicity, necessitating a reduction in olaparib dose compared with monotherapy to avoid life-threatening side effects such neutropenia, leukopenia, anemia, and thrombocytopenia (14). Similarly, dose-escalation studies of olaparib in combination gemcitabine (15), doxorubicin (16), and paclitaxel (17) found that the monotherapy daily dose (400 mg b.i.d.) could not be reached with an acceptable tolerability profile. In contrast, daily olaparib maintenance therapy after combination therapy was well tolerated (18).

Combining olaparib with radiotherapy is a promising strategy to selectively enhance DNA-damage at the primary site of drug action. Focused-beam X-ray radiation produces localized dsDNA breaks and thus would be expected to render cells more sensitive to PARP inhibition (19, 20). Olaparib has been shown to enhance the effect of radiation in both BRCA2-proficient and deficient prostate cancer cells (21). Several PARP inhibitors have been shown to have radiosensitizer activity in vivo (20, 22–25); however, these agents have not yet been demonstrated to provide radiosensitivity to radiation-resistant tumors. Clinically, olaparib has been demonstrated to have antitumor activity in a subset of prostate cancer patients with mutated DNA damage repair genes, notably BRCA2 and ATM (7), but has not been tested in combination with radiation.

Here we describe NanoOlaparib, a lipid-based nanoformulation of olaparib with demonstrated efficacy in radiation-resistant mice. Three PTEN-deficient prostate cancer cell lines were tested for their sensitivity to olaparib and NanoOlaparib. The most clinically relevant line, radiation-resistant FKO1 cells derived from a Pten/Trp53-deficient mouse model of advanced prostate cancer (26), was further investigated and found to selectively increase γ-H2AX expression in a radiation-dose dependent manner using immunocytochemistry and Western blotting. A subcutaneous model of prostate cancer, generated in mice, was used to assess the therapeutic efficacy of NanoOlaparib alone and in combination with a single dose of focused-beam X-ray radiation. To determine the potential mechanism by which NanoOlaparib works in vivo, ultra-short time-to-echo MRI (UTE-MRI) was utilized to quantify ferumoxytol accumulation in tumors following NanoOlaparib administration. The results of these studies suggest that NanoOlaparib may be a promising strategy for enhancing radiosensitivity of advanced prostate tumors with PTEN and TP53 deletions.

NanoOlaparib was synthesized using 1, 2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dioleoyl-3-tri methyl-ammonium-propane (chloride salt) (DOTAP), cholesterol, and 1,2-distearoyl-sn-glycero-3 phosphoethanolamine-N-[methoxy(polyethyleneglycol)-2000 (DSPE-PEG₂₀₀₀) purchased from Avanti Polar Lipids and olaparib (Selleck Chemicals). DPPC, DOTAP, cholesterol, DSPE-PEG₂₀₀₀, and olaparib were individually dissolved in chloroform and combined at a molar ratio of 13.6:0.43:1.29:0.717:5.75. Following overnight solvent evaporation under rotary vacuum, the lipid-drug melt was hydrated using PBS (pH 7.4, final concentration 2.5 mg/mL olaparib) at 50°C, emulsified by heating and cooling five times with agitation, and then sonicated at 25°C for 10 minutes to produce highly condensed lipid nanoparticles containing hydrophobic olaparib. Unbound drug was removed by dialysis for 4 to 6 hours. For animal studies, nanoparticles were concentrated by centrifugation at 3,200 rcf for 45 minutes and resuspension to yield a final drug concentration of 5.0 to 10.0 mg/mL. Conventional olaparib was dissolved in DMSO for use as a control.

The mean nanoparticle diameter, zeta potential, and concentration were measured by laser scattering microscopy (Zetaview, ParticleMetrix) using PBS (pH 7.4 or 6.0) as solvent. Nanoparticles were visualized using transmission electron microscopy (TEM; JEM-1010, JEOL) after being dried on 300-mesh copper-coated carbon grids (Electron Microscopy Sciences) and negatively stained with a 1.5% uranyl acetate solution (Sigma-Aldrich). Drug encapsulation and release was measured using high-performance liquid chromatography (HPLC; Agilent 1100 series). Olaparib was detected at 207 nm on a reverse phase C18 column (SUPELCO) with a mobile phase of methanol: water (64 : 36). The nanoparticle loading efficiency was determined by dissolving nanoparticles in methanol prior to HPLC. Drug release was determined by dialyzing nanoparticles against PBS and measuring the supernatant with HPLC.

Three PTEN-deficient prostate cancer cell lines were evaluated for their response to olaparib, including human PC3 (ATCC), human LNCaP (ATCC), and mouse FKO1 (Pten^pc−/−;Trp53^pc-/−, provided by Dr. Pandolfi; refs. 27, 28). All cell lines were tested and authenticated using short tandem repeat (STR) profiling. All cell lines were passaged for fewer than 6 months after receipt. Cells were grown in F12-K (PC3), RPMI (LNCaP), or DMEM media (FKO1) supplemented with 10% FBS (HyClone). The FKO1 media was further supplemented with 25 μg/mL bovine pituitary extract (Life Technologies), 5 μg/mL insulin (Sigma Aldrich), and 6 ng/mL epidermal growth factor (Life Technologies). All cells were maintained in a 5% CO₂ atmosphere at 37°C. To determine the optimal seeding densities for extended cell assays, the doubling rate of each line was measured by MTS assay for metabolic activity (Promega). The cell doubling times were found to be 36.5 hours (PC3), 64.9 hours (LNCAP), and 19.5 hours (FKO1).

For IC₅₀ determination, exponentially growing cells were treated continuously with 0 to 100 μmol/L olaparib or NanoOlaparib for four population doublings. The final cell count was determined using an MTS assay and the IC₅₀ was calculated as the drug concentration required to inhibit growth of 50% of cells. For the short-term cell viability assay, cells in exponential growth phase were pretreated with 1 μmol/L olaparib or NanoOlaparib for 24 hours, irradiated (0, 2, or 4 Gy), replated at low density (∼300–600 cells/cm²) 4 hours after irradiation, and then cultured in complete media without drug for nine additional doubling cycles (∼6–24 days). The colonies were fixed, stained using crystal violet (BioPioneer), and dissolved in acetic acid for quantification of absorbance at 590 nm. The percent cell viability was determined relative to an untreated control. Drug uptake by subconfluent monolayers was measured using HPLC following 24-hour drug treatment and normalized for protein content using a Bradford assay. For the long-term clonogenic cell survival assay, cells in exponential growth phase were pretreated with 1 μmol/L olaparib or NanoOlaparib for 24 hours, irradiated (0–10 Gy), replated at low density (∼300–600 cells/cm²) 4 hours after irradiation, and then incubated continuously with drug for a minimum of 14 days (29, 30). Colonies stained with crystal violet were manually counted using a pen and used to estimate the surviving fraction as a function of the plating efficiency (31). To generate the radiation dose–response curves, the data were fitted to a linear quadratic model. The sensitizer enhancement ratio (SER) at 10% cell survival was calculated according to the equation:

For all radiation studies, cells received a single dose of 0–10 Gy X-rays using a 220 kVp beam delivered at 13 mA (dose rate of 5.45 Gy/min) via a 0.15 mm copper filter using a Precision Small Animal Radiation Research Platform (SARRP, XRad).

Subconfluent monolayers of FKO1 cells (5 × 10⁴ cells/cm²) were pretreated for 24 hours with 0 to 5 μmol/L olaparib, NanoOlaparib, or a vehicle control and then irradiated with 2 to 10 Gy. Cells were fixed 30 minutes after radiation treatment, permeabilized with 0.05% Triton-X100 for 10 minutes, blocked with a 1% BSA/2% goat serum solution for 10 minutes, and then labeled with anti-γ-H2AX 1:1,500 (Cell Signaling Technology), anti-Rad51 1:500 (AbCam), and/or anti-DNA polymerase θ (polθ) 1:500 for 1 hour. Following three washes, the cells were probed with Alexa Fluor 488 and 647 IgG 1:500 (Invitrogen) for 1 hour, washed, and counterstained with DAPI (Life Technologies). Images were acquired using a Zeiss LSM 710 laser-scanning confocal microscope (DAPI Ex: 350 nm, Em: 410–480 nm; Rad51 Ex: 488 nm, Em: 505–530 nm; Polθ Ex: 488 nm, Em: 505–530 nm; γH2AX Ex: 633 nm, Em: 660–710 nm) using a 40 × or 63 × objective. The mean γ-H2AX fluorescent signal intensity was analyzed using 200 nuclei per treatment group from a minimum of 10 random fields, using ImageJ v4.1. The damage enhancement ratio was calculated from the mean γ-H2AX fluorescence intensity at each radiation dose as described in ref. 32.

A subcutaneous mouse model of radiation-resistant prostate cancer was generated by a one-time subcutaneous implantation of 10⁶ (in 200 μL PBS) FKO1 cells (derived from Pten^pc−/−;Trp53^pc−/− mice) in the right flank of nude (nu/nu) mice at Northeastern University. Mice were randomized into four treatment groups (n = 6–7) when tumors reached 180 ± 20 mm³ in volume. NanoOlaparib (40 mg/kg drug) was administered twice weekly via intravenous injection for up to 12 weeks. The NanoOlaparib dose was selected based on allometric scaling of the human oral daily dose (800 mg) and then reduced by 40% to compensate for the full bioavailability of intravenously administered drug. Mice treated with radiation received a one-time X-ray dose of 10 Gy when tumors reached 250 ± 50 mm³ using a Precision Small Animal Radiation Research Platform (SARRP, XRad). Tumor size was measured twice weekly using calipers, and all animals were monitored and weighed at least twice per week. Mice were removed from the therapeutic study when tumors reached 1,500 mm³, imaged using MRI, and sacrificed immediately thereafter. Harvested tissues were fixed in 10% formalin. All animals were maintained in accordance with institutional rules and ethical guidelines for animal care.

Animal imaging was performed at the Center for Translational Neuroimaging (CTNI) at Northeastern University in accordance with the Division of Laboratory Animal Medicine and Institutional Animal Care and Use Committee. MRI images were obtained at ambient temperature (∼25°C) using a Bruker Biospec 7.0T/20-cm USR horizontal magnet (Bruker) equipped with a 20-G/cm magnetic field gradient insert (ID = 12 cm, Bruker) and a 300 MHz, 30 mm mouse coil (Animal Imaging Research, LLC). T₁-weighted, T₂-weighted, and ultra-short time-to-echo (UTE) images were acquired at 150 μm resolution within a 3 cm field-of-view as previously described (33). The UTE pulse-sequence used a 200 kHz fixed trajectory with acquisition parameters TE = 13 microseconds, TR = 4 milliseconds, and θ = 20°. Mice were randomized into one of three treatment groups: untreated (no olaparib), 4 hours pretreated (NanoOlaparib, 1 × 40 mg/kg), or chronically treated (NanoOlaparib, 40 mg/kg twice weekly for ≥2 weeks). Ferumoxytol accumulation in the vasculature and tumor was monitored 0 and 24 hours respectively after a one-time intravenous bolus injection of 14 mg/kg ferumoxytol. Precontrast images were acquired immediately prior to ferumoxytol injection. To assess how NanoOlaparib treatment changes ferumoxytol accumulation, vehicle-treated mice were given one injection of NanoOlaparib (40 mg/kg) following ferumoxytol clearance and then re-injected with a new bolus of ferumoxytol (14 mg/kg) 4 hours later. T₁, T₂, and UTE signal intensity images were transformed and rendered using 3DSlicer v4.4 (34). To quantify ferumoxytol accumulation, the UTE signal intensity was measured for each voxel within the tumor volume. The fold-increase in voxel number as a function of signal intensity was determined by subtracting the precontrast signal histogram from the postcontrast histogram. The percentage of tumor volume with ferumoxytol accumulation was calculated from the fold-increase histogram by counting the total number of voxels above a threshold intensity of 750 and normalizing by the total number of tumor voxels.

Harvested tissues were embedded in paraffin, cut, and stained by the Dana-Farber/Harvard Cancer Center Research Pathology Core. To assess tissue pathology, tumor sections were stained with H&E according to manufacturer's recommendations.

Statistical significances were analyzed by Student t test or one-way analysis of variance with P ≤ 0.05 considered statistically significant. All data were expressed as mean ± SD (in vitro data) or mean ± SEM (in vivo data). Survival fraction curve was analyzed by KaleigaGraph 4.0, then fitted to the data using a linear–quadratic model. The median survival time was estimated from the Kaplan–Meier curve and a log-rank test was used pairwise to test for statistically significant differences (35).

Poorly soluble olaparib was encapsulated into lipid nanoparticles using bulk synthesis techniques. Lipid films containing cholesterol, DPPC, DOTAP, DSPE-PEG₂₀₀₀, and olaparib were hydrated with aqueous solvent and sonicated to produce monodisperse, unilamellar nanoparticles loaded with clinically relevant concentrations of inhibitor. Nanoparticles were characterized using a laser scattering video microscope to obtain nanoparticle count, size, and zeta potential information. The mean nanoparticle size was measured to be 71 ± 5 μm (PDI = 0.07) with less than 7% batch-to-batch variation at time of synthesis. The representative size distribution at synthesis and following 3 months of storage is shown in Fig. 1A. Nanoparticles had an overall surface charge of −24 ± 7 mV, indicative of effective charge shielding of the cationic lipids by PEG. Nanoparticles had a rounded or partially concave appearance under TEM when counterstained uranyl acetate (Fig. 1A, inset). Using HPLC, the average nanoparticle loading was estimated to be 5.0 mg/mL for solutions containing approximately 4 × 10⁷ nanoparticles/mL. Olaparib release at physiological pH occurred slowly (Fig. 1B), characterized by first-order drug release with a half-life of 4.3 days (R² = 0.99; Fig. 1B, inset).

Loss of the PTEN or TP53 tumor suppressor genes is commonly observed in prostate cancer, whereas their combined loss is often observed in advanced prostate cancer. The therapeutic efficacy of olaparib and NanoOlaparib treatment was assessed in three PTEN-deficient prostate cancer cell lines with differing radiation resistance including LNCaP (AR^pos, p53^wt), PC-3 (AR^neg, p53^null), and FKO1 (AR^insensitive, p53^null; Fig. 2A). Nanoparticles were diluted to contain the same number of drug molecules as the free olaparib, with μmol/L NanoOlaparib denoting the amount of drug contained (not the concentration of nanoparticles). Cell viability was determined by the counting of cells transiently treated (24 hours) with olaparib or NanoOlaparib and then allowed to proliferate in the absence of drug for nine population doublings. Here, radiation was performed 24 hours after drug addition. Both LNCaP and PC-3 cells displayed a dose-dependent response to radiation alone, whereas the FKO1 cells were radiation-resistant. All cell lines demonstrated increased sensitivity to radiation following pretreatment with 1 μmol/L olaparib or NanoOlaparib. The relative change in sensitivity was greatest for the FKO1 cells, which demonstrated approximately 70% a decrease in cell viability with PARP inhibition compared to radiation alone. Cumulative drug uptake did not significantly differ for FKO1 cells treated 24 hours with olaparib or NanoOlaparib, as measured by solid-phase extraction followed by HPLC. The IC₅₀ for continuously treated FKO1 cells was determined to be 2.2 μmol/L (olaparib) and 3.0 μmol/L (NanoOlaparib) in the absence of radiation, demonstrating that these cells are relatively insensitive to PARP-1 inhibition as a monotherapy. Reported values for prostate cancer cell lines considered olaparib-sensitive are on the order of 0.59 μmol/L (LNCaP) and 0.79 μmol/L (PC-3), whereas cells with BRCA1/BRCA2 mutations are 0.02 to 0.2 μmol/L (29). A long-term clonogenic assay was performed to determine the fraction of FKO1 cells surviving radiation treatment alone and in combination with continuous PARP inhibition (Fig. 2B and C). Both olaparib and NanoOlaparib were found to significantly enhance the effect of radiation at doses of 6 Gy and above (P < 0.05). At 10% cell survival, the SER (SER₁₀) was measured to be 1.28 (olaparib) and 1.81 (NanoOlaparib). The relative benefit of NanoOlaparib over olaparib was observed to be greatest at 10 Gy.

FKO1 cell monolayers treated with radiation were examined for expression of phosphorylated histone H2AX (γ-H2AX), a biomarker of cellular response to dsDNA breaks (36). Compared to untreated controls, NanoOlaparib pretreatment with concentrations of 0.2 μmol/L and above produced a statistically significant increase in mean nuclear fluorescence intensity that persisted for 4 hours following irradiation (Supplementary Fig. S1A and S1B). The nuclei of cells pretreated with 1 μmol/L olaparib or NanoOlaparib showed a significant increase in both γ-H2AX foci number and foci intensity as early as 0.5 hours after irradiation (Fig. 3A). The mean nuclear fluorescence was quantified across a range of different radiation doses (Fig. 3B). Olaparib alone was found to increase nuclear γ-H2AX expression at all radiation doses; however, this damage paralleled that caused by radiation alone, suggesting that free drug increases the basal level of γ-H2AX expression in a radiation-insensitive manner. In contrast, cells treated with NanoOlaparib displayed a clear radiation dose-dependent enhancement in γ-H2AX expression. At 10 Gy, NanoOlaparib pretreatment resulted in a damage enhancement factor of 4.2 relative to vehicle-treated controls.

Rad51, a recombinase that assists in the repair of dsDNA breaks (37), and polθ, a DNA polymerase that can promote both homologous recombination (38) and microhomology-mediated end-joining (39) in response to dsDNA breaks, were also examined for changes in protein expression. Both Rad51 and polθ foci were observed in the nuclei of cells following NanoOlaparib treatment (Supplementary Fig. S2); however, neither the number of foci nor the fraction of cells expressing foci increased following combination therapy (NanoOlaparib + 6 Gy radiation). Only Rad51 expression in the nucleus was found to colocalize with γ-H2AX staining. Cytoplasmic Rad51 staining was found to increase transiently in a drug-dose dependent manner following irradiation, with peak staining observed 30 minutes after radiation (Supplementary Fig. S1A and S1C). Cytoplasmic Rad51 staining correlated linearly with increasing NanoOlaparib concentration, resulting in a 4.7-fold increase in Rad51 staining compared to non-irradiated cells treated with an equivalent dose of NanoOlaparib (Supplementary Fig. S1D). This pattern of staining has previously been shown to be a clinical hallmark of highly aggressive prostate cancer with PTEN (40) or BRCA (41) mutations, and may reflect impaired nuclear transport of Rad51 (42).

The ability of NanoOlaparib to overcome radiation-resistance in vivo was assessed using FKO1 cells (derived from Pten^pc-/−;Trp53^pc-/− mice; ref. 26) implanted subcutaneously in nude mice. Given the large number of mice enrolled in the study and the need for regular tumor size monitoring, a subcutaneous model was selected over the spontaneous prostate cancer model. Mice received one of four treatments: untreated, 10 Gy radiation, 40 mg/kg NanoOlaparib (i.v. twice-weekly), or 40 mg/kg NanoOlaparib (i.v. twice-weekly) + 1 × 10 Gy radiation. Tumors receiving combination therapy were pretreated three times (on days 0, 3, 7) with NanoOlaparib and then irradiated on day 9, when the tumor size matched that of the radiation only controls (irradiated on day 1). For both irradiated and non-irradiated mice, NanoOlaparib treatment was continued twice weekly for the duration of the study. The individual tumor response is shown in Fig. 4A and averaged group response is shown in Fig. 4B. Irradiated FKO1 tumors demonstrated radiation-resistance in vivo, growing at the same average rate as untreated controls. NanoOlaparib-treated tumors demonstrated a slower rate tumor growth compared to mice receiving radiation and untreated controls. All tumors treated with combination therapy showed a decrease in tumor size following irradiation, and in three of six mice, the tumors continued to shrink until completely gone. The average rate of tumor growth following combination therapy showed increased variability with time as a result of three tumors growing while three were shrinking.

Enhanced survival was observed for mice treated with NanoOlaparib and combination therapy (Fig. 4C), with a median survival time of 33 days (NanoOlaparib) and 66 days (combination) compared to 22 days (controls and radiation only). In mice receiving combination therapy, the average time to reach 1,500 mm³ was found to be 48 days for those with relapsing tumors (n = 3; Fig. 4D) whereas the remaining mice showed no tumor relapse over the 90-day observation period (n = 3). No significant weight loss was observed for all treatment groups (Fig. 4E). For mice treated with NanoOlaparib, no adverse correlates of hematopoietic damage were found in the organs of mononuclear phagocyte system, including the liver, spleen, kidney, lung, and heart (Fig. 5A). Minor necrosis and inflammation appeared in the heart of only one mouse that received both radiation and NanoOlaparib. The tumors of different treatment groups showed clear cytological differences. Untreated and irradiated tumors showed densely packed, viable cells throughout the tumor volume. NanoOlaparib-treated tumors showed regional variations in tumor cell density but no necrosis. Relapsed tumors (NanoOlaparib + 10 Gy radiation) were characterized by large areas of necrosis and swollen, vacuolated cells surrounded by a dense border of cells only at the tumor margin (Fig. 5B).

Given the apparent long-term benefit of administering NanoOlaparib before and after a single dose of radiation, we sought to determine whether NanoOlaparib administration changes drug delivery to the tumor, using ferumoxytol as a surrogate nanoparticle (Fig. 6). Ferumoxytol accumulation was imaged 24 hours after nanoparticle administration using ultra-short time-to-echo (UTE) MRI, a technique that produces positive-contrast images of ferumoxytol (33). In longitudinal experiments, multiple ferumoxytol boluses were administered sufficiently apart (Fig. 6A) to allow for blood pool clearance (33). Figure 6B shows representative T₁, T₂, and UTE signal intensity images from a single longitudinal experiment, collected in a mouse treated with a lipid vehicle followed by NanoOlaparib. Here, darkening of the T₁ and T₂ images, and brightening of the UTE images, indicate the presence of ferumoxytol. A small amount of ferumoxytol accumulation was observed following vehicle pre-treatment; however, this accumulation was visibly enhanced following NanoOlaparib pretreatment. The UTE signal intensity was measured for each voxel in the tumor volume and plotted as a histogram (Fig. 6C). Ferumoxytol accumulation appeared as an extended tail that diverged from the precontrast histogram (no ferumoxytol) at a UTE signal intensity of approximately 750. To determine the relative signal enhancement with NanoOlaparib treatment, the fold-increase in voxel number was plotted as a function of signal intensity (Fig. 6D). At any given UTE signal intensity, up to a four-fold increase in the number of voxels was observed, demonstrating that NanoOlaparib-enhanced ferumoxytol accumulation is higher than would be expected for a second bolus of ferumoxytol alone.

To assess the reproducibility of NanoOlaparib-enhanced accumulation, the percentage of signal-enhanced voxels with ferumoxytol accumulation (defined here as a UTE signal above 750) was quantified across 11 different tumors treated NanoOlaparib, 1 × 10 Gy radiation, or no treatment (Fig. 6E). Tumors treated with a single dose of NanoOlaparib demonstrated ferumoxytol accumulation in 18.5 ± 8% of voxels, an overall 5- and 19-fold enhancement compared to irradiated tumors and untreated tumors, respectively. The overall distribution of ferumoxytol accumulation was observed to vary widely from tumor-to-tumor when rendered as maximum intensity projection images, generally appearing as one or more foci of high accumulation surrounded by large areas of diffuse accumulation (Fig. 6F).

There is a compelling need for developing an intravenous formulation of olaparib. Like chemotherapy, olaparib is administered at the maximum tolerated dose (400 mg b.i.d.) because only a small fraction of consumed drug reaches the tumor. Oral olaparib pharmacokinetics is far from ideal: the drug is metabolized rapidly into nonfunctional metabolites and thus patients must swallow up to 16 capsules per day to maintain effective drug concentrations. Up to 80% of patients can expect to experience one or more adverse events including nausea, fatigue, or diarrhea (43). In contrast, nanoformulation allows poorly soluble drugs such as olaparib to be administered directly into the bloodstream and circulate longer. This increased bioavailability is expected to enhance drug's therapeutic efficacy at clinically equivalent doses or below, opening possibilities to reduce the olaparib dose and thereby ameliorate systemic side effects.

The nanoformulation described here has several unique advantages that are not observed with the free drug. First, the nanoformulation has enhanced therapeutic efficacy in radiation-resistant cells in vitro at both high and low doses of radiation. Considering the poor bioavailability of oral olaparib, we would expect this difference to be further magnified when a bioequivalent dose is administered in vivo. Second, NanoOlaparib does not raise the basal level of γ-H2AX expression but instead enhances γ-H2AX expression in a radiation-dose dependent manner. Given that γ-H2AX expression generally correlates with dsDNA damage, this suggests NanoOlaparib may be less toxic than the free drug in tissues not receiving radiation. Third, the damage enhancement factor of NanoOlaparib was significantly higher (P < 0.05) than that of olaparib for all radiation doses above 2 Gy. In stereotactic body radiation therapy (SBRT) for prostate cancer ablation, patients can safely receive five fractions consisting of 9 to 10 Gy each (44), thus it is realistic to envision combining NanoOlaparib with high doses of radiation. For these reasons, we chose to study the therapeutic benefit of combining NanoOlaparib and radiation in cells derived from Pten^pc−/−;Trp53^pc−/− mice that have previously been reported as unresponsive to olaparib monotherapy (26).

Cancers lacking homologous recombination (HR) capacity are generally sensitive to PARP inhibition. Here we looked at polθ, a protein whose expression is related to HR. Polθ expression is frequently elevated in cancers with HR deficiency (45), likely due its secondary capacity as a promoter of microhomology-mediated end-joining (39). The restoration of normal HR function in these cells reduces polθ expression to normal levels (38), thus polθ expression is believed to inversely correlate with HR activity. In our studies, polθ expression remained constant with treatment and no colocalization of polθ was observed, suggesting that alternative nonhomologous end-joining is not a dominant mechanism of dsDNA repair in FKO1 cells. Given the normal expression of Polθ, it is unsurprising that FKO1 cells are relatively insensitive olaparib and NanoOlaparib monotherapy.

NanoOlaparib clearly displayed radiosensitizer activity in vivo. Because the half-life of peglyated lipid nanoparticles is generally several days (46), mice were pretreated three times (days 0, 3, 7) with NanoOlaparib to allow drug accumulation. Mice then received a single 10 Gy dose of radiation, selected to maximize the radiation dose-enhancement factor of NanoOlaparib while allowing rapid recovery of body weight after irradiation. Given that radiation-induced DNA damage and genomic instability requires days to weeks to result in widespread cell death (47), NanoOlaparib treatment was continued twice-weekly for the duration of the study. The single dose of radiation, when combined with NanoOlaparib pretreatment and maintenance, was sufficient to produce a complete response in half of the mice. This statistically significant response (P < 0.01) was well beyond that observed by adding the effects of each monotherapy, indicating that NanoOlaparib can act synergistically with radiation to overcome radiation-resistant tumors. NanoOlaparib alone did show statistically significant growth inhibition and enhanced overall survival. Given that previous reports of Olaparib monotherapy in Pten^pc-/−;Trp53^pc-/- mice demonstrated no tumor growth inhibition (26), we believe this difference is due to improved delivery and/or uptake of olaparib.

Several PARP inhibitors exhibit vasoactive properties, allowing for increased tumor oxygenation and/or improved drug delivery (20, 21, 48). Olaparib has been reported to transiently enhance tumor vessel perfusion in non–small cell lung cancer xenografts (20), which may explain the enhanced sensitivity of these tumors to olaparib in combination with fractionated radiation. Here, changes in nanoparticle accumulation were measured after a single dose of PARP inhibitor. We found that NanoOlaparib actively enhances nanoparticle accumulation in tumors by as much as 19-fold. Given the robust enhancement in median survival time following NanoOlaparib + radiation treatment, we speculate that the in vivo efficacy of NanoOlaparib may be potentiated by its ability to enhance its own accumulation in tumors. Whether this behavior is mediated by changes in tumor perfusion, or some other mechanism, remains to be determined.

Ferumoxytol accumulation has recently been shown to predict the localization of therapeutic nanoparticles in tumor microvasculature (49). Here, the use of ferumoxytol as a surrogate provides interesting new insights into the treatment efficacy data. We observed that ferumoxytol accumulation following NanoOlaparib treatment varied tumor by tumor, with one tumor showing significantly less accumulation. This heterogeneity may account for some of the observed differences in radiosensitization. Interestingly, the three relapsed NanoOlaparib + radiation tumors all showed a greater rate of tumor growth during the NanoOlaparib pretreatment phase (days 0–9) than those with a complete response, suggesting that these relapsed tumors were less responsive to NanoOlaparib treatment alone. Given that radiosensitization is determined by the quantity of radiosensitizer present, as well as the distribution of the radiosensitizer throughout the tumor (50), we would expect that tumors with poor NanoOlaparib uptake would display a less robust response to radiation. It may be that some FKO1 tumors are intrinsically less conducive to nanoparticle accumulation, or alternatively, less amenable to NanoOlaparib-induced enhancements in nanoparticle accumulation.

In conclusion, we have shown that NanoOlaparib renders PTEN/TP53-deficient radiation-resistant tumors sensitive to radiation. NanoOlaparib treatment is sufficient to trigger a significant tumor response, and when administered in combination with radiation, can lead to a complete response. The mechanism by which nanoformulated olaparib sensitizes tumors to radiation is complex and likely extends beyond a simple enhancement in drug accumulation, as NanoOlaparib displays several unique behaviors in vitro that are not observed using an equivalent concentration of free drug. Additionally, NanoOlaparib appears to have the capacity to enhance nanoparticle accumulation in tumors and this may help potentiate its own accumulation. The results of this study show that NanoOlaparib may be a promising strategy for enhancing the sensitivity of radiation-resistant tumors lacking BRCA mutations, such as those with PTEN and TP53 deletions.

No potential conflicts of interest were disclosed.

Conception and design: A.L. van de Ven, S. Tangutoori, C. Gharagouzloo, R. Cormack, S. Sridhar

Development of methodology: A.L. van de Ven, S. Tangutoori, J. Qiao, C. Gharagouzloo, R. Cormack, S. Sridhar

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): A.L. van de Ven, S. Tangutoori, P. Baldwin, J. Qiao, C. Gharagouzloo, N. Seitzer, J.G. Clohessy, R. Cormack, S. Sridhar

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): A.L. van de Ven, S. Tangutoori, P. Baldwin, J. Qiao, C. Gharagouzloo, N. Seitzer, R. Cormack, S. Sridhar

Writing, review, and/or revision of the manuscript: A.L. van de Ven, S. Tangutoori, C. Gharagouzloo, N. Seitzer, G.M. Makrigiorgos, R. Cormack, S. Sridhar

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S. Tangutoori, C. Gharagouzloo, P.P. Pandolfi, S. Sridhar

Study supervision: S. Sridhar

FK01 cells were kindly donated by the Pandolfi group (BIDMC/Harvard Medical School). The authors thank Benjamin Geilich for his assistance with the TEM. We thank Dana-Farber/Harvard Cancer Center for the use of the Rodent Histopathology Core.

This work was supported by the grants NSF-DGE-0965843, NIH HHS U54CA151881, and CIMIT 13-1807 (to S. Sridhar), the Mazzone Foundation (to S. Sridhar and R. Cormack), and NIH NCIR01CA082328 (to P.P. Pandolfi).

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