Abstract
Taxane-based chemotherapeutics are clinically available as frontline treatment regimens for cervical cancer. However, drug resistance and life-threatening toxicity impair the clinical efficacy of taxanes, so more effective and less toxic therapeutic modalities are urgently needed. Cabazitaxel has attracted increasing interest due to its potential to circumvent the drug resistance by taxanes. We previously showed that tethering docosahexaenoic acid (DHA) to cabazitaxel enabled the prodrug to self-assemble into nanoparticles in water. Despite this encouraging finding, the DHA–cabazitaxel conjugate formulation requires further optimization to enhance nanoparticle retention and tumor delivery. We here integrated this conjugate into amphiphilic poly(ethylene glycol)-block-poly(D,L-lactic acid) copolymers to assemble dCTX NPs. The nanoparticle abrogated P-glycoprotein–mediated resistance in cancer cells. In a docetaxel-resistant cervical tumor xenograft-bearing mouse model, the efficacy was augmented by the nanotherapy when compared with solution-based free drugs (i.e., docetaxel and cabazitaxel). Dose intensification of dCTX NPs markedly suppressed the tumor growth in this model. Detailed studies revealed that systemic toxicity was alleviated, and MTD of dCTX NPs was at least 3 times higher than that of free cabazitaxel in animals, which may enable dose increases for clinical studies. In conclusion, the new formulation addresses essential requirements in terms of the stability, safety, and translational capacity for initiating early-phase clinical trials.
Introduction
Cervical cancer is one of the most prevalent gynecologic malignancies. Each year, cervical cancer accounts for more than half a million new cases, and over 300,000 females die from the disease worldwide (1, 2). Persistent infection by high-risk human papillomavirus (HPV) is closely associated with the development of cervical cancer. Although effective prophylactic vaccines against the most lethal oncogenic HPV types are clinically available, only limited numbers of patients can receive the vaccine treatment (3, 4). In North America, approximately 85% of patients are diagnosed at early stages; however, in many less developed countries, most cervical cancers are diagnosed in the advanced stages because these patients are either lack of awareness or unaffordable for HPV vaccine (5–7).
Chemotherapy continues to be a mainstay treatment regimen for advanced and metastatic cervical cancer. Taxane-based chemotherapeutics [e.g., paclitaxel and docetaxel (DTX), Supplementary Fig. S1], which are the frontline treatments for cervical cancer, are clinically available alone or in combination with other agents and have a significant therapeutic benefit (8–10). Taxanes induce the apoptosis of rapidly proliferating cancer cells by binding to tubulin and stabilizing microtubules (11). Unfortunately, the clinical efficacy of taxane-based treatment regimens remains unsatisfactory because of the induction of drug resistance and dose-limiting toxicity (12). These limitations prevent cancer therapies from achieving stable and complete responses, and thus need for alternative modality.
Generally, only limited amounts (less than 1%) of small-molecule drugs reach tumors when they are systemically administered in a traditional solution-based dosage form (13). Nanoparticle-mediated drug delivery has been proposed as an alternative approach to deliver increased amounts of drugs to tumor lesions (14). Taking advantage of the unique pathologies observed in solid tumors, including defective leaky tumor vasculature and ineffective lymphatic drainage, nanoparticles with small sizes can passively accumulate in target tumors through the enhanced permeability and retention (EPR) effect (15–22). This finding has motivated many investigators to explore nanoparticle strategies used as a viable solution to achieve the magic bullet concept for cancer therapy proposed by Paul Ehrlich (23–26). Free chemotherapeutics must be stably entrapped in nanoparticles in the systemic circulation to decrease indiscriminatory distribution in healthy tissues. After accumulating in tumors, drug molecules must be released at a high efficiency (27–29). More importantly, nanoparticle approaches have been suggested to have the ability to partially overcome drug resistance by sustained drug release or by reducing the interaction of drug molecules with multidrug resistance (MDR) proteins (30, 31). Consistent with these approaches, some nanoparticle-based therapeutics have been approved for clinical use, and many others are undergoing clinical trials (32, 33). Several taxane nanoformulations, including albumin-bound paclitaxel (e.g., Abraxane) and amphiphilic block copolymer-encapsulated paclitaxel (e.g., Genexol-PM), have been approved for clinical use (34–36).
Cabazitaxel (Jevtana; Sanofi-Aventis), which is a potent mitotic inhibitor, has attracted rapidly increasing interest in preclinical studies and clinical trials due to its potential to circumvent drug resistance induced by paclitaxel and DTX. The structure of cabazitaxel provides this molecule with low affinity for the MDR1 [or P-glycoprotein (P-gp), which is encoded by the ABCB1 gene] protein, thereby avoiding drug efflux (37, 38). Despite this advantage, high systemic toxicity of cabazitaxel has been observed in phase I studies, and its MTD has been established as only 25 mg/m2 in a 1-hour infusion (i.v. injection) every 3 weeks (39). Previously, we showed that tethering a polyunsaturated fatty acid (PUFA) such as docosahexaenoic acid (DHA) to cabazitaxel enables the spontaneous self-assembly of a DHA–cabazitaxel prodrug conjugate to create organic solvent-free supramolecular nanotherapeutics (40). “PUFAylation” of cabazitaxel not only enables this conjugate to self-assemble in water without any exogenous excipients but also enhances drug tolerability in animals. Thus, this conjugate was shown to be extremely effective with low toxicity, warranting further investigation.
To further optimize the delivery matrices, and to assess the therapeutic potential in drug-resistant cervical cancer, we formulated a DHA–cabazitaxel conjugate within the clinically approved copolymer poly(ethylene glycol)-block-poly(D,L-lactic acid) (PEG-b-PLA) to assemble dCTX NPs. Interestingly, the DHA–cabazitaxel conjugate showed higher compatibility with the PEG-b-PLA matrix compared with parent cabazitaxel, and this conjugate can be stably encapsulated in PEG-b-PLA–composed nanoparticles. The optimized formulation addresses essential requirements in terms of the stability, safety, and translational capacity for future evaluation in clinical trials. We examined the cytotoxicities of this new nanotherapy in DTX-resistant HeLa human cervical cancer cells. Furthermore, in a preclinical DTX-resistant HeLa tumor xenograft-bearing mouse model, compared with solution-based free drugs (i.e., DTX and cabazitaxel), the nanoparticles showed a significantly improved antitumor efficacy. Finally, detailed studies suggested that the systemic toxicity was substantially reduced in animals, enabling increased doses for clinical studies.
Materials and Methods
Materials and compounds
DTX and cabazitaxel were purchased from Knowshine Pharmachemicals Inc. DHA was purchased from Tokyo Chemical Industry. The copolymer PEG-b-PLA was purchased from Advanced Polymer Materials Inc.
Preparation of DHA–cabazitaxel prodrug-loaded polymeric nanoparticles
The DHA–cabazitaxel conjugate was synthesized as described previously (40). The conjugate was obtained through one-step esterification with an 80% yield from cabazitaxel and DHA. DHA–cabazitaxel prodrug-loaded nanoparticles were prepared using a nanoprecipitation method. Briefly, cabazitaxel prodrugs (1 mg cabazitaxel equivalent) and PEG5k-b-PLA8k (19 mg) were dissolved in acetone (1 mL), and the solution was then added dropwise into deionized (DI) water (9 mL) while stirring. After stirring for 10 minutes, the remaining organic solvent was removed in a rotary evaporator under reduced pressure. Finally, the nanoparticle solution was concentrated with centrifugal filter devices (Amicon Ultra4, 10k molecular weight cut-off, Millipore Corp.) and washed with DI water. The particle size was measured by dynamic light scattering (DLS), and the drug concentration was determined by analytical high-performance liquid chromatography (HPLC).
Cell lines and cell culture
HeLa human cervical cancer cells and A549 human lung cancer cells were obtained from the cell bank of the Chinese Academy of Sciences (Shanghai, China). The cells were maintained in RPMI 1640 medium supplemented with 10% FBS, penicillin (100 units/mL) and streptomycin (100 μg/mL). DTX-resistant HeLa (HeLa/DTX) cells and DTX-resistant A549 (A549/DTX) cells were maintained in RPMI 1640 medium. All cells were identified by short tandem repeat DNA fingerprinting and cultured in a humidified incubator with 5% CO2 at 37°C.
Cell apoptosis assay
FITC Annexin V Apoptosis Detection Kit (556547, BD Biosciences) was used to assess the apoptotic rate of cancer cells. Briefly, 1 × 106 cells per well were seeded in 6-well plates overnight and exposed to drugs for 48 hours. Then, detached and attached cells were collected by trypsinization, washed twice with cold phosphate buffer saline (PBS, 0.01 mol/L phosphate, pH = 7.4, 135 mmol/L NaCl, 2.7 mmol/L KCl, 1.5 mmol/L KH2PO4, and 8 mmol/L K2HPO4), and resuspended in 500 μL 1 × binding buffer. Next, the cells were mixed with 5 μL FITC Annexin V and 5 μL propidium iodide (PI) and incubated for 15 minutes in the dark at room temperature. After incubation, the levels of apoptosis were analyzed with flow cytometry (BD LSRFortessa). Untreated cells suspended in PBS were used as the control.
Cell-cycle assay
Cell-cycle distribution and DNA content were determined using PI staining and flow cytometry. Cells (1 × 106 cells/well) were adhered to 6-well plates and treated with drugs for 24 hours. Next, the cells were harvested, washed with cold PBS, and fixed in 75% ethanol at −20°C overnight. Following incubation, the cells were washed with PBS twice and stained in 500 μL staining buffer containing 5 μL PI solution for 30 minutes in the dark at room temperature. Cell-cycle profiles were analyzed with flow cytometry (BD LSRFortessa). Untreated cells suspended in PBS were used as the control.
Western blot
Total cell lysates were prepared and fractionated on 10% SDS-PAGE gels and transferred to polyvinylidene difluoride membranes. The membranes were blocked with 5% milk in Tris-buffered saline with 0.1% Tween 20 (TBST) and incubated overnight at 4°C in the presence of anti-Bcl2, anti-survivin, anti-cleaved caspase 9, anti-cleaved PARP, anti-Cdc25c, anti-cyclin B1, anti-Cdc2, anti–p-Cdc2, and anti–β-actin primary antibodies (Cell Signaling Technology). After washing with TBST, the membranes were incubated with secondary antibodies for 1 hour at room temperature. Protein bands were detected using enhanced chemiluminescence (Fude Biological Technology). β-Actin was used for normalization of protein loading.
Ethics statement
The experimental protocols were approved by the Ethics Committee of the Sir Run Run Shaw Hospital, Zhejiang University School of Medicine. All of the animal studies were conducted according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
In vivo pharmacokinetic study with dCTX NPs
Sprague–Dawley rats (∼250 g, n = 6 in rats each group) were administered a single i.v. injection of free cabazitaxel (formulated in 1:1 v/v polysorbate 80/ethanol) or dCTX NPs at a dose of 15 mg/kg (cabazitaxel equivalent). Blood samples were collected at 5, 15, and 30 minutes and at 1, 2, 4, 7, 24, and 48 hours in 1.5 mL centrifugal tubes containing sodium heparin. Plasma was separated from the blood by centrifuging for 10 minutes at 3,000 × g and stored at −80°C until analysis. To analyze the drug concentrations, the plasma samples (50 μL) were added to acetonitrile (300 μL) and ultrasonicated to extract cabazitaxel and the DHA–cabazitaxel conjugate. The drug concentrations were determined by reverse-phase HPLC (RP-HPLC). The pharmacokinetic parameters were obtained by fitting the drug concentrations versus each time point to a noncompartmental pharmacokinetic model using PKSolver, an add-in program for pharmacokinetic and pharmacodynamic data analysis in Microsoft Excel (41).
In vivo biodistribution of dCTX NPs using near-infrared fluorescence imaging
Balb/c nude mice bearing HeLa/DTX tumor xenografts were used to evaluate the tissue biodistribution of dCTX NPs. We randomly divided the mice into two groups (n = 6 mice in each group) when their tumor volumes reached approximately 100 mm3. The mice were administered a single injection of dCy5.5-labeled dCTX NPs (termed dCy5.5@dCTX NPs, 1 mg/kg Cy5.5-equivalent dose and 20 mg/kg of cabazitaxel equivalent dose) via the tail vein. Free Cy5.5 (1 mg/kg) was also injected as a reference. Whole-body and ex vivo NIR fluorescence imaging were performed using a Xenogen IVIS Lumina imaging system (PerkinElmer). At time points of 24 and 48 hours post injection, the mice were sacrificed and the major organs and tumors were excised. Prior to ex vivo imaging, the tissues were washed with PBS 3 times to rule out the nanoparticle residue in the blood vessels. Finally, the tumor tissues were fixed, processed into frozen sections, and stained with 4,6-diamidino-2-phenylindole. We observed the distribution of dCy5.5@dCTX NPs under a confocal fluorescence microscope (ZEISS 880, Germany).
Antitumor activity in a preclinical DTX-resistant tumor xenograft-bearing mouse model
To evaluate the antitumor activity of the nanoparticles, 5-week-old Balb/c nude mice were subcutaneously implanted with cell suspensions containing 7 × 105 HeLa/DTX cells in the right flank. The mice bearing HeLa/DTX tumor xenografts were established approximately 10 days after implantation. When the tumor volume reached approximately 100 mm3, the mice were randomly divided into six groups (n = 6 mice in each group). The mice were i.v. injected with saline, free DTX (10 mg/kg), free cabazitaxel (5 mg/kg), and dCTX NPs (5, 10, and 15 mg/kg cabazitaxel equivalent) on days 0, 3, and 6. The tumor volume and body weight were logged until the tumors reached 2,500 mm3. The tumor volume (V) was evaluated by measuring the length (L) and width (W) and calculated using the following formula: V = (L × W2)/2, W being smaller than L. Finally, the mice were sacrificed by CO2 inhalation, and the tumor weights were obtained.
Evaluation of the drug toxicity in healthy ICR mice
The in vivo toxicity of free cabazitaxel and dCTX NPs was evaluated in healthy ICR mice (4–5 weeks old). The mice were i.v. injected with saline, free cabazitaxel, and dCTX NPs via the tail vein 3 times on days 0, 3, and 6. We obtained the body weight and peripheral blood cell counts, as well as the hepatorenal toxicity using biochemistry of mouse plasma. After the injection regimen, the major organs were excised and subjected to hematoxylin and eosin (H&E) staining to analyze the damage induced by the drugs.
Statistical analysis
All quantitative data are presented as the mean ± SD. The statistical significance between different groups was analyzed using an unpaired Student t test. A P value less than 0.05 was considered statistically significant, and a P value less than 0.01 was considered highly significant.
Results
Preparation and characterization of DHA–cabazitaxel prodrug-loaded polymeric nanoparticles
We first assessed whether the DHA–cabazitaxel conjugate can be formulated in clinically improved amphiphilic PEG-b-PLA copolymers. To test this, we followed a nanoprecipitation protocol by adding a mixture of DHA–cabazitaxel and PEG5k-b-PLA8k in acetone into DI water under stirring (Fig. 1A). Further removal of the organic solvent enabled to obtain a solution of dCTX NPs, which is expected to be systemically injectable for preclinical studies. Representative transmission electron microscope and scanning electron microscope images were shown in Fig. 1B, revealing the formation of spherical nanoparticles with a diameter of approximately 50 nm in water. Further DLS measurements suggested that the hydrodynamic diameters (DH) of dCTX NPs were approximately 73 nm (Fig. 1C). To exert the cytotoxic activity, free cabazitaxel must to be hydrolyzed from the DHA–cabazitaxel conjugate. Many forms of esterase play critical roles in the metabolism of lipids and bond cleavage of organic species. Specifically, previous studies suggest that various types of cancer cells highly express esterase, which could be used for selective conversion of ester prodrugs into therapeutically active drugs. Thus, we assessed the hydrolysis of the DHA–cabazitaxel conjugate analyzed by HPLC. Clearly, esterase activation substantially accelerated the conversion of the prodrug into active cabazitaxel, whereas the hydrolysis was negligibly observed in the absence of esterase (Fig. 1D). The results evidenced that once uptake by cancerous cells, intracellular esterase could trigger the release of active cabazitaxel. In addition, inhibition of premature hydrolysis and release of free drugs in the blood is a prerequisite to accomplish in vivo activity and to reduce side effects. We therefore evaluated the hydrolysis of the prodrug when dCTX NPs were incubated with rat whole blood or plasma (Fig. 1E). After 72-hour incubation, approximately 64% of the prodrugs remained intact. Considering that the circulation time of nanoparticles composed of PEG-b-PLA is usually less than 24 hours, we thus expect that this scaffold could constrain the drug payloads in nanoparticles during systemic circulation. Furthermore, under the current formulation, we determined the encapsulation efficiency and drug loading for dCTX NPs to be 99.8% and 4.7%, respectively.
Preparation and characterization of DHA–cabazitaxel conjugate-formulated polymeric nanoparticles (dCTX NPs). A, Chemical structure of the DHA–cabazitaxel conjugate and schematic illustration of the nanoprecipitation protocol. B, Transmission electron microscope (TEM) image of dCTX NPs. Inset: scanning electron microscope (SEM) image of nanoparticles. C, The size distribution of dCTX NPs measured by DLS analysis. D, Hydrolytic release of free cabazitaxel from the DHA–cabazitaxel conjugate. The conjugate was incubated in the mixture solution of DMSO and HEPES buffer (1:3, v/v), and the hydrolytic event was monitored by analytical HPLC. E, Release of free cabazitaxel from the DHA–cabazitaxel conjugate incubated in whole blood or in plasma. The data are presented as the mean ± SD (n = 3).
Establishment of DTX-resistant cell lines
In this study, the DTX-resistant HeLa/DTX cancer cell line was established using chemosensitive HeLa human cervical cancer cells according to a previously established protocol. Pulsed exposure of HeLa cells to DTX with stepwise increments of time was performed (42). In the adaptation stage, the parent HeLa cells were exposed to DTX at a 15 nmol/L concentration for different times ranging from 0.5 to 48 hours (i.e., 0.5, 1, 2, 4, 12, 24, and 48 hours). Each treatment was repeated in triplicate. After each treatment, the surviving cells were harvested and expanded in DTX-free medium. In the following consolidation stage, previously surviving cells were cultured in medium containing 15 nmol/L DTX for 72 hours until they expanded normally in this medium (42).
Eventually, we successfully established the DTX-resistant HeLa/DTX cells, as evidenced by the increased IC50. For instance, the IC50 value of HeLa/DTX cells after 72 hours of DTX exposure was 63.5 ± 6.0 nmol/L, which was 7.7-fold greater than that in the parent HeLa cells (Table 1). To further test the efficacy of cabazitaxel, we established DTX-resistant A549 human lung cancer cell (A549/DTX) using the same protocol. As expected, the IC50 values in A549/DTX cells increased to 205.2 ± 38.2 nmol/L after exposure to free DTX for 72 hours, indicating a 93.3-fold enhancement relative to DTX-sensitive A549 cells. To investigate the mechanism of drug resistance, we examined the expression of taxane resistance–associated P-gp in these cell lines by Western blot. A significantly increased expression level of P-gp was observed in HeLa/DTX and A549/DTX cells compared with DTX-sensitive cancer cells (Fig. 2A).
In vitro cytotoxicity of cabazitaxel dissolved in DMSO and dCTX NPs compared with DTX dissolved in DMSO after 72 hours of incubation (presented as IC50 ± SD in nmol/L)a.
The cytotoxicity of dCTX NPs compared with cabazitaxel against DTX-resistant cell lines in vitro. A, Overexpression of P-gp protein was observed in A549/DTX and HeLa cells as determined by Western blot. B, HeLa/DTX cells were treated with varying concentrations (from 3 to 24 nmol/L) of cabazitaxel and dCTX NPs (cabazitaxel equivalent concentration) for 24, 48, 72, and 96 hours. Analysis of apoptosis (C and D) and cell cycle (E and F) of dCTX NPs of HeLa/DTX and A549/DTX cells. G and H, Quantification of cell apoptosis– and cell-cycle–related proteins in A549/DTX and HeLa/DTX cells after treatment with drugs for 48 hours. The concentrations of drugs in HeLa/DTX and A549/DTX cells were 24 nmol/L for apoptosis and Western blot analysis and 12 nmol/L for cell-cycle analysis, respectively. The data are presented as the mean ± SD (n = 3); *, P < 0.05 and **, P < 0.01.
In vitro cytotoxicity of cabazitaxel prodrug nanoparticles in DTX-resistant cells
Because of the low affinity for P-gp, cabazitaxel has the potential to overcome drug resistance induced by paclitaxel and DTX. We thus assessed the activity of dCTX NPs compared with free DTX and free cabazitaxel in DTX-sensitive and the established DTX-resistant cell lines. As shown in Table 1, the IC50 values in HeLa/DTX and A549/DTX cells after 72 hours of dCTX NP treatment were 12.8 ± 3.1 and 17.7 ± 2.8 nmol/L, respectively, suggesting that dCTX NPs possessed potent activity in DTX-resistant cells. Notably, the potency of dCTX NPs was in the same range as that of free cabazitaxel (i.e., the IC50 values of cabazitaxel were 14.4 ± 2.1 and 8.5 ± 5.4 in A549/DTX and HeLa/DTX cells, respectively). Furthermore, the growth inhibition assay showed that dCTX NPs had a potent inhibitory effect in HeLa/DTX cells, and the effectiveness increased as the concentration increased when the treatment time ranged from 24 to 72 hours (Fig. 2B). Unexpectedly, compared with treatment with free cabazitaxel, treatment with dCTX NPs for 96 hours showed a statistically greater inhibitory effect. This result likely indicated the sustained release of therapeutically active cabazitaxel from the nanoparticles over time.
Efficient apoptosis induction and G2–M cell-cycle arrest by cabazitaxel prodrug nanoparticles
Similar to other taxanes, cabazitaxel exerts cytotoxic effects through mitotic arrest by stabilizing microtubules, leading to cell death. To elucidate the mechanism of action, the cell apoptosis and cell-cycle progression induced by dCTX NPs were examined. DTX treatment induced a high level of apoptosis and effectively arrested the G2–M cell cycle in both sensitive A549 and HeLa cells, but showed negligible activities in HeLa/DTX and A549/DTX cells (Fig. 2C–F; Supplementary Figs. S2 and S3). Compared with DTX, dCTX NPs increased the apoptotic rates of cells and promoted the accumulation of cells in the G2–M phase. For instance, in the HeLa/DTX cell line, the apoptotic rates for the DTX, free cabazitaxel, and dCTX NP treatments were 23.4%, 52.6%, and 49.6%, respectively. Moreover, the proportions of cells in the G2–M phase for the DTX, free cabazitaxel, and dCTX NP treatments were 11.1%, 69.1%, and 67.8%, respectively (Fig. 2E and F; Supplementary Fig. S2). Collectively, these cell-based results demonstrated that following the ligation of DHA and the nanoparticle formulation, the potency in inducing apoptosis and cell-cycle arrest by dCTX NPs compared with free cabazitaxel was not statistically decreased.
Furthermore, we conducted a detailed analysis of several regulatory proteins associated with drug-induced apoptosis and G2–M cell-cycle arrest. As shown in Fig. 2G and H, the exposure of cells to cabazitaxel and dCTX NPs led to highly efficient cleavage of PARP, caspase 9. In addition, the downregulation of the antiapoptotic protein Bcl2, surviving, and the cell-cycle–related proteins Cdc25c, Cdc2, and cyclin B1 were confirmed. Overall, our data demonstrated that cabazitaxel and dCTX NPs remained highly potent in DTX-resistant cancer cells by promoting apoptosis and G2–M phase arrest via the Bcl2/PARP and Cdc25c/Cdc2/cyclin B1 pathways.
We further studied the cellular uptake of dCTX NPs in HeLa/DTX cells. To visualize dCTX NPs under observation of fluorescence microscope, a lipophilic fluorescent probe, DiI, was loaded into dCTX NPs to form DiI-labeled dCTX NPs (termed DiI@dCTX NPs). HeLa/DTX cells were treated with free DiI and DiI@dCTX NPs and subjected to confocal laser fluorescence microscopy (CLSM) observation for cellular uptake as well as the intracellular distribution. As shown in Supplementary Fig. S4, we found that the fluorescence signal distributed in the lysosomes as evidenced by the colocalization with the lysotracker after the treatment of DiI@dCTX NPs. By contrast, free DiI rapidly diffused into cells and randomly distributed in the cytosol and nucleus with no obvious colocalization with the lysotracker. These results indicated that dCTX NPs internalized into the cells through the endocytosis/lysosome pathway, whereas free DiI entered the cells by diffusion.
Pharmacokinetic studies of dCTX NPs
Premature release of drugs from nanocarriers during systemic circulation, which reduces drug delivery to tumor tissues and enhances systemic exposure of toxic drugs, remains a major obstacle for achieving favorable therapeutic efficacy. Through the encapsulation of the DHA–cabazitaxel prodrug in long circulating PEG-b-PLA nanoparticles, the pharmacokinetic properties of drugs can be rationally improved. To explore the role of PEG-b-PLA matrices as nanocarriers, we conducted pharmacokinetic studies by analyzing drug concentrations in the blood of Sprague–Dawley rats. Blood was obtained from the Sprague–Dawley rats at predetermined timepoints after single i.v. injection of dCTX NPs (at 15 mg/kg cabazitaxel equivalence) via the tail vein. The cabazitaxel concentrations in serum were determined by HPLC analysis. The concentration–time profiles of total cabazitaxel in plasma and related pharmacokinetic parameters are shown in Fig. 3A and B. Indeed, compared with the clinical formulation of free cabazitaxel, the PEG-b-PLA nanoparticles significantly prolonged the drug retention and circulation in rats. Free cabazitaxel exhibited rapid clearance from the blood, making this agent untraceable in our HPLC system at 1 hour after injection. The area as extrapolated from the concentration versus time curve (AUC0-t) for dCTX NPs was 984.2 ± 217.4 μg·h/mL, which was 55.9-fold greater than the plasma AUC0-t of the cabazitaxel formulation (Fig. 3B). On the basis of this substantial increase in AUC0-t, these results indicated that compared with free cabazitaxel, dCTX NPs showed extremely high drug accumulation in solid tumors.
In vivo pharmacokinetic studies of dCTX NPs. A and B, Plasma concentration–time profiles and pharmacokinetic parameters of dCTX NPs compared with free cabazitaxel following a single i.v. injection in Sprague–Dawley rats. The total amount of cabazitaxel molecules, including released cabazitaxel and intact DHA–cabazitaxel conjugate, in the plasma was extracted and analyzed by HPLC. Drugs were injected at a dose of 15 mg/kg (cabazitaxel-equivalent dose). C and D, A comparison of released free cabazitaxel and intact DHA-conjugate in the plasma of Sprague–Dawley rats after dCTX NPs were injected. In vivo plasma concentration–time profiles (C) and pharmacokinetic parameters (D) of free cabazitaxel and the respective intact DHA-conjugate after a single i.v. injection of dCTX NPs in rats. t1/2 indicates the half-life of the distribution phase; Cmax indicates maximum concentration; and AUC indicates the area under the concentration versus time curve. The data are presented as the mean ± SD (n = 6).
Furthermore, we separately quantified the concentrations of released cabazitaxel and DHA–cabazitaxel conjugate in blood samples. Unexpectedly, only a small portion (∼11%) of free cabazitaxel was hydrolyzed from dCTX NPs (Fig. 3C and D). The results suggested that the dCTX NP scaffold had excellent stability and showed negligible hydrolysis and premature release in the blood after systemic administration. Limited drug exposure in the circulation likely suggests a low systemic toxicity.
Tumor-specific nanoparticle accumulation
Next, we investigated the retention and tissue distribution of our nanotherapy by conducting near infrared (NIR) fluorescence imaging in HeLa/DTX xenograft-bearing mice. To mimic the tethering of PUFA to drugs, the NIR dye Cy5.5 was ligated to DHA, and this conjugate was coassembled with DHA–cabazitaxel in PEG-b-PLA matrices (termed dCy5.5@dCTX NPs). For comparison, free Cy5.5 dissolved in the clinical formulation of cabazitaxel was analyzed. Following a single i.v. injection, whole-body in vivo fluorescence imaging was conducted. The NIR signal decayed rapidly in mice that received free Cy5.5, but the NIR signal did not decay rapidly in the dCy5.5@dCTX NP–treated mice (Fig. 4A). Moreover, in dCy5.5@dCTX NP–treated mice, strong NIR fluorescence was observed in the tumor regions. Furthermore, we determined the quantity of Cy5.5 in the organs to indicate the distribution profiles of dCy5.5@dCTX NPs. At 24 and 48 hours post injection, we excised the major organs and tumors from the mice for ex vivo imaging and quantified the fluorescence intensities in each organ (Fig. 4C and D). As expected, free Cy5.5 showed negligible tumor accumulation, but high accumulation in the liver and kidney was observed. Noticeably, dCy5.5@dCTX NPs showed higher accumulation than free Cy5.5 in tumors.
In vivo biodistribution of dCy5.5@dCTX NPs in HeLa/DTX xenograft-bearing mice. A, In vivo whole-body NIR fluorescence imaging of mice at 24 and 48 hours after i.v. injection of free Cy5.5 dye and dCy5.5@dCTX NPs via the tail vein. The white dotted circles indicate the tumor regions. Ex vivo NIR fluorescence imaging of the major organs (B) and tumors (C) excised from the mice. D, Quantitative analysis of fluorescence intensities in the tumor region and major organs at 24 and 48 hours post administration. Representative fluorescence images (E) and quantification of fluorescence intensities (F) of tumors obtained from the mice that were i.v. injected with free Cy5.5 and dCy5.5@dCTX NPs. The fluorescence images were observed under a confocal laser scanning microscope. The scale bars represent 100 μm. The data are presented as the mean ± SD (n = 4); *, P < 0.05 and **, P < 0.01.
Furthermore, we examined the tumor cross-section under CLSM. The CLSM results clearly revealed that strong signals derived from Cy5.5 were distributed throughout the entire tumor tissues in the mice administered dCy5.5@dCTX NPs (Fig. 4E and F). However, the tumor tissues excised from the free Cy5.5-injected mice exhibited negligible Cy5.5 signals.
In addition to these fluorescence imaging studies, we further comparatively analyzed cabazitaxel accumulation in tumors. After a single injection of cabazitaxel and dCTX NPs, the tumor tissues were excised and the concentration of drugs was determined by analytical HPLC. Noticeably, at both time points post administration, the tumors of mice receiving dCTX NPs demonstrated statistically higher cabazitaxel concentration when compared with free cabazitaxel injected in Jevtana-mimicking formulation (Fig. 4G). Moreover, we separately quantified the amounts of released free cabazitaxel and nonhydrolytic intact DHA–cabazitaxel conjugates in tumors (Fig. 4H). We confirmed the presence of both drug forms, suggesting that the prodrug can be readily converted into the therapeutically active cabazitaxel in target tumors.
Collectively, these results were well correlated with the pharmacokinetic data, suggesting that our dCTX NP scaffold has the potential to enhance drug accumulation in tumors, with lower toxicity to normal tissues, making it a promising candidate for further investigation.
In vivo therapeutic efficacy in a preclinical DTX-resistant tumor-bearing mouse model
Encouraged by the potent in vitro cytotoxicity and superior tumor accumulation, we thus explored the therapeutic efficacy in Balb/c nude mice bearing subcutaneous DTX-resistant HeLa tumor xenografts. When the tumors grew to 100 mm3, we initiated the treatment by i.v. injecting of DTX, cabazitaxel, and dCTX NPs. Following three injections, the tumor size and body weight were obtained (Fig. 5D). Unfortunately, the DTX treatment at a dose of 10 mg/kg resulted in rapid tumor growth, which was correlated with the in vitro cytotoxicity results showing that DTX lost its activity in DTX-resistant HeLa cancer cells. As shown in Fig. 5A–C, the administration of free cabazitaxel at a 5 mg/kg dose suppressed the tumor growth but additionally caused substantial body weight loss (approximately 22%) on day 12 post administration, suggesting severe systemic toxicity in Balb/c nude mice. A comparable therapeutic effect was achieved by administering dCTX NPs at a dose of 5 mg/kg (cabazitaxel equivalence); however, the mice that were injected at this dose showed negligible body weight loss (Fig. 5D). The enhanced drug tolerability in animals inspired us to examine whether increasing the dose yields better therapeutic outcomes. Encouragingly, higher doses of dCTX NPs markedly delayed the growth of drug-resistant tumors. Specifically, treatment with dCTX NPs at 15 mg/kg was associated with the superior activity in this model. Administration of dCTX NPs continued to show durable tumor repression that persisted for at least 2 weeks after the last injection. At the end point of the therapeutic study, the mice were sacrificed, and the average weight of tumors excised from the mice that received 15 mg/kg (cabazitaxel-equivalent dose) of nanotherapy was 0.17 g, which was significantly less than that of the tumors in mice that received cabazitaxel treatment (0.48 g, cabazitaxel vs. dCTX NPs, P < 0.01). More notably, increasing the dose of the nanotherapies compared with cabazitaxel substantially alleviated toxicity in animals, as indicated by no noticeable reduction in the body weight of the mice during the treatment with dCTX NPs. As shown in Fig. 5D, administration of free cabazitaxel at 5 mg/kg led to approximately 22% body weight loss. However, dCTX NPs at 15 mg/kg (cabazitaxel equivalent) only produced a drop of approximately 11% in mouse body weight. We thus can safely conclude that the MTD of dCTX NPs was at least 3 times higher than that of free cabazitaxel in animals.
The therapeutic efficacy of dCTX NPs was evaluated in a HeLa/DTX xenograft-bearing Balb/c nude mouse model. A, Changes in the tumor volume following treatment with saline (n = 6), free DTX in its clinical formulation (10 mg/kg, n = 6), cabazitaxel (5 mg/kg, n = 6), dCTX NPs (5, 10, and 15 mg/kg cabazitaxel equivalent, n = 6) for three successive injections on days 0, 3, and 6. B and C, Tumor weights and representative images of the excised tumors from each group. D, Changes in the body weights of the mice after the treatment. The data are presented as the mean ± SD (n = 6); *, P < 0.05 and **, P < 0.01. E, Representative images from H&E staining (E), TUNEL analysis (F), and Ki67 staining (H) of the excised tumors on day 21. The scale bars represent 100 μm in length. Quantification of TUNEL-positive (G) and Ki-67–positive cells (I). Potent induction of apoptosis and inhibition of proliferation were observed in the dCTX NP–treated group, consistent with the H&E staining results.
Histologic analysis of the excised HeLa/DTX tumors further validated the antitumor activity of dCTX NPs. H&E staining (Fig. 5E) and the terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay (Fig. 5F) revealed that efficient and extensive intratumoral apoptosis in the tumor tissues was elicited by the dCTX NP treatment (10 mg/kg, cabazitaxel equivalent), consistent with the observed reduction in the tumor volume. The percentage of TUNEL-positive cells in dCTX NP–treated mouse tumors increased compared with that of other treatments (Fig. 5G). IHC staining using the proliferation marker Ki67 further confirmed the reduced intratumoral cell proliferation in the sections from the dCTX NP group (Fig. 5H), which was also verified by the quantification of Ki67-positive cells (Fig. 5I). Therefore, these histologic analyses provided the evidence that compared with the cabazitaxel treatment (5 mg/kg), dCTX NPs at 10 mg/kg (cabazitaxel equivalent) exhibited superior cytotoxic effects.
In vivo safety of dCTX NPs
Finally, the safety profiles of the nanoparticle formulations were carefully examined in healthy mice. Clinically, cabazitaxel causes high systemic toxicity in patients with cancer. Neutropenia is one of the most severe side effects, which prevents increasing the dose of cabazitaxel. We anticipate that the nanoparticle approach presented in this study could partially alleviate the in vivo toxicity of cabazitaxel when used in vivo. To validate this assumption, we performed a toxicity study in healthy ICR mice (n = 10 mice in each group) using dCTX NPs compared with the clinical formulation of cabazitaxel. Figure 6A–C and Supplementary Table S1 show variations in the hematologic parameters in mice that received i.v. injections of dCTX NPs (10 and 20 mg/kg, cabazitaxel equivalent) compared with those that received the same dosage of cabazitaxel. After the injection of cabazitaxel, the mice spontaneously exhibited a substantial reduction in the white blood cell (WBC) counts (∼70%), neutrophil leukocyte (NE) counts (∼95%), and lymphocyte (LY) counts (∼50%) on day 6 compared with these blood counts on day 0. As a result of dCTX NP administration, myelosuppression and a reduction in blood cells were observed, but the differences between day 6 and day 0 were not statistically significant. In addition, after stopping the dosage, these hematologic parameters recovered to the normal range.
Systemic toxicity of the nanotherapy compared with the clinical formulation of cabazitaxel in ICR mice. A–C, WBC, NE, and LY counts in mice on days 0, 6, 9, and 15. The mice were administrated with dCTX NPs (10 and 20 mg/kg, cabazitaxel equivalent dose) 3 successive times via the tail vein. Saline and cabazitaxel were also injected as controls. Analysis of AST (D), ALT (E), total bilirubin (TBIL; F), ALP (G), creatinine (CR; H), and BUN (I) in the ICR mice on day 9 after three injections. The data are presented as the mean ± SD (n = 6); *, P < 0.05; **, P < 0.01; and ***, P < 0.001.
Furthermore, we analyzed a series of serological markers to indicate the presence of lesions in the liver and kidneys induced by the drugs. As shown in Fig. 6D–I, increased expression levels of aspartate aminotransferase (AST), alanine aminotransferase (ALT), alkaline phosphatase (ALP), and blood urea nitrogen (BUN) were observed in the cabazitaxel-treated mice, indicating the acute hepatorenal toxicity. Notably, dCTX NPs show these types of changes for these indicators.
The in vivo safety of dCTX NPs was further supported by the stable body weight of the ICR mice after receiving the treatment. Administration of dCTX NPs at 10 mg/kg (cabazitaxel equivalence) exhibited stable body weight gain during the observation period. However, after injecting mice with cabazitaxel at a 10 mg/kg dose, a significant body weight loss (∼16%) was observed (Fig. 7A). Finally, to assess drug toxicity, we performed histologic analyses of organs excised from the mice. As shown in Fig. 7B, the absence of signs of necrosis and cell death in all organs indicated that the mice can tolerate dCTX NPs therapy at high doses. Unfortunately, renal damage was observed in the cabazitaxel-treated mice. Collectively, these results indicated that we successfully converted toxic cabazitaxel into a safe nanotherapy that is extremely well tolerated in animals by encapsulation of cabazitaxel prodrug in nanoparticles.
Toxicity evaluation of the nanotherapy compared with the clinical formulation of cabazitaxel in healthy ICR mice. A, Changes in the body weights were observed for 18 days after the three administrations of the drugs. The mice were i.v. injected with cabazitaxel and dCTX NPs (5 and 10 mg/kg, cabazitaxel equivalent dose). The data are presented as the mean ± SD (n = 6). B, Representative images of H&E staining of the major organs.
Discussion
Microtubules are components of the cytoskeleton that are involved in many crucial cellular interphase functions, including maintenance of cell shape, intracellular transport, mitosis, motility, and cell signal transmission. They are composed of α-tubulin and β-tubulin heterodimers in dynamic equilibrium with tubulin polymers in cells. During cell division, the principal function of microtubules is the formation of the mitotic spindle (43). Therefore, the dynamic equilibrium is an attractive drug target. Taxane drugs bind to tubulin, thereby stabilizing microtubules and promoting tubulin polymerization (11). Eventually, these agents arrest quickly proliferating cells at the G2–M phase and lead to efficient tumor cell apoptosis. Currently, FDA-approved taxane drugs that target the taxane-binding site include paclitaxel, DTX, and cabazitaxel. However, the clinical efficacy of taxanes (e.g., paclitaxel and DTX) is typically limited by the development of drug resistance. Repeated drug exposure causes cancer cells to evolve to overexpress the ATP-dependent drug efflux pump P-gp (44, 45). Hence, overexpression of P-gp by tumor cells is responsible for constitutive and acquired resistance to taxanes. Accordingly, many efforts have been dedicated to developing new taxane derivatives that are not strong substrates for P-gp.
In addressing the issue of taxane resistance, cabazitaxel has emerged as a second-generation taxane. Compared with paclitaxel and DTX, cabazitaxel has been suggested to have a low affinity for P-gp drug efflux, circumventing the resistance of current taxanes (37, 46). Many previous reports have suggested that cabazitaxel has high potency in a broad spectrum of taxane-resistant cancer cells according to in vitro cell-based assays (37, 47). Clinically, cabazitaxel has been approved by the FDA for treating the patients with metastatic castration-resistant prostate cancer (mCRPC; ref. 38). Compared with previous standard chemotherapeutic treatments, chemotherapy with DTX treatment provides a survival benefit of approximately 2 to 3 months. Unfortunately, patients with mCRPC who had previously been treated with DTX become taxane resistance (48). Clinical data have shown that the patients receiving cabazitaxel have significantly longer overall survival than those receiving other therapeutic modalities (49). Despite these favorable clinical outcomes, the success of using cabazitaxel has been limited in clinical practice due to the high toxicity of the drug. In phase I studies, the MTD of cabazitaxel was established as only 25 mg/m2 in a 1-hour infusion every 3 weeks (39). This MTD is substantially lower than that of paclitaxel and DTX (175 and 60–100 mg/m2 for paclitaxel and DTX administration every 3 weeks, respectively; refs. 50, 51). However, similar to other taxanes, cabazitaxel is not soluble in aqueous solutions. To make cabazitaxel applicable for i.v. injections, surfactants and/or organic solvents are required. Currently, only one formulation of cabazitaxel in polysorbate 80 and ethanol (under the trademark Jevtana) has been approved for the clinical use. Although polysorbate 80 has relatively low toxicity, the use of this excipient inevitably causes severe hypersensitivity reactions (52, 53).
To mitigate these obstacles, we previously tethered the DHA motif to this potent agent through an ester linkage to create a DHA–cabazitaxel conjugate (40). Unexpectedly, despite the substantial hydrophobicity of the overall molecule, this prodrug is capable of self-assembling into nanoparticles without the use of excipients. After uptake by cells where a high concentration of esterases is presented, the prodrug undergoes cleavage of the ester bond, releasing therapeutically active cabazitaxel and a DHA moiety. The latter composite is abundant in the humans and plays an essential role in biological functions, obviating concerns over modification-associated side effects. Furthermore, our previous results demonstrated the effectiveness and low toxicity of this conjugate in animal models, making it a promising new drug candidate for further preclinical evaluation in taxane-resistant cancer. Previous studies have suggested that the formulation matrices have a substantial impact on the in vivo performance of nanoparticle therapeutics. Hence, with the aim of optimizing the delivery matrices and assessing the efficacy in taxane-resistant tumors, a clinically approved copolymer matrix (i.e., PEG-b-PLA) with high stability in the blood and a low critical micelle concentration was used to formulate the DHA–cabazitaxel prodrug.
In DTX-resistant HeLa and A549 cells, dCTX NPs potently induced the cell apoptosis, arrested the cell cycle at the G2–M phase, and showed potency that was comparable with that of free cabazitaxel. Interestingly, compared with free cabazitaxel, treatment with dCTX NPs for 96 hours led to a statistically higher inhibitory activity (Fig. 2A). In general, prodrug constructs require additional cleavage and release steps to yield their active drug forms, decreasing the potency of prodrug formulations. In this study, we did not observe delayed in vitro potency. This lack of a finding could be attributed to the rapid release of active cabazitaxel under esterase-rich intracellular conditions because cabazitaxel is conjugated to DHA by a hydrolytic ester bond.
Furthermore, we explored the potential of using dCTX NPs in a preclinical mouse model bearing DTX-resistant cervical cancer xenografts. Administration of DTX at 10 mg/kg showed limited antitumor efficacy, indicating the maintenance of drug resistance when the tumor cells were grown in mice (Fig. 5). Interestingly, compared with cabazitaxel, dCTX NPs slightly improved the efficacy (not statistically significant) but markedly decreased drug toxicity in nude mice. Notably, we safely increased the dosage of dCTX NPs to 10 and 15 mg/kg, which enhanced the durability of tumor reduction. Finally, we administered dCTX NPs to healthy ICR mice via the tail vein to examine the tolerability of the nanoparticles. Compared with cabazitaxel (formulated in 1:1 v/v polysorbate 80/ethanol), dCTX NPs were proven to have a better safety profile, as indicated by a stable body weight and negligible changes in hematologic parameters (Figs. 6 and 7). In addition, the absence of liver and kidney dysfunction and the absence of signs of necrosis and cell death in the major organs (H&E staining) were confirmed in the nanoparticle-treated mice, suggesting the marked safety of the nanoparticles. These results were well correlated with the in vivo pharmacokinetic analysis, strongly supporting the hypothesis that individual conjugates are stably encapsulated within the hydrophobic core of the nanoparticles. Moreover, the long-circulation characteristics of PEG-b-PLA nanoparticles could extend the drug persistence in the blood, thus making the nanoparticles preferentially accumulate in tumor lesions via the EPR effect. Detailed pharmacokinetic studies showed that the nanoparticle reservoir can impair the premature release of free cabazitaxel during circulation, supporting the markedly alleviated drug toxicity in animals. Nonetheless, alleviation of the toxicity of the anticancer drug cabazitaxel facilitated the dose increases and thereby improved the efficacy.
In conclusion, the data presented in this study provide compelling evidence that the integration of DHA–cabazitaxel into nanoparticles reduces relevant side effects by limiting the systemic exposure of free toxic drugs while enhancing the therapeutic efficacy in a preclinical mouse model bearing a taxane-resistant cervical malignancy. The overall nanosystems (i.e., the formulation consisting of polymer matrices and encapsulated prodrug entities) are composed of FDA-approved materials without introducing additional uncertified molecules. Following further investigation, we aim to initiate early-phase clinical trials to assess the safety and efficacy of this nanotherapy in patients with taxane-resistant cancer.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Authors' Contributions
Conception and design: B. Xie, W. Han, H. Wang
Development of methodology: B. Xie, J. Wan, X. Chen, H. Wang
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): B. Xie, J. Wan, X. Chen
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): B. Xie, J. Wan, X. Chen, W. Han, H. Wang
Writing, review, and/or revision of the manuscript: B. Xie, J. Wan, W. Han, H. Wang
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): B. Xie, J. Wan, X. Chen
Study supervision: W. Han, H. Wang
Acknowledgments
This work was supported by grants from the National Natural Science Foundation of China (grants 81773193 and 81571799 to H. Wang, and 81972745 and 81572361 to W. Han), the Zhejiang Province Preeminence Youth Fund (grant LR19H160002 to H. Wang), and the Ten Thousand Plan Youth Talent Support Program of Zhejiang Province (grant ZJWR0108009 to W. Han).
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.
Footnotes
Note: Supplementary data for this article are available at Molecular Cancer Therapeutics Online (http://mct.aacrjournals.org/).
Mol Cancer Ther 2020;19:822–34
- Received June 20, 2019.
- Revision received November 7, 2019.
- Accepted December 13, 2019.
- Published first December 17, 2019.
- ©2019 American Association for Cancer Research.
References
Volume 19, Issue 3
