SapC–DOPS Nanovesicles as Targeted Therapy for Lung Cancer

Lung cancer is the most common and the deadliest type of cancer worldwide, accounting for 1.8 million new cases and 1.6 million deaths in 2012 (1). Cigarette smoking is the single leading cause of lung cancer, followed by environmental and occupational exposure to pollutants such as radon, arsenic, asbestos, Bis(chloromethyl) ether, chromium, nickel, vinyl chloride, and polycyclic aromatic compounds (2–4). Startlingly, a direct causal relationship between atmospheric particulate matter (air pollution) and lung and other cancers was recently established. As reported by the International Agency for Research on Cancer, more than half of lung cancer–related deaths caused by air pollutants worldwide occurred in China and other East Asian countries (5). Standard treatment options for lung cancer (chemotherapy, radiation, and surgery) have several undesirable side effects that affect the quality of life of the cancer patient.

Liposomal formulations as vehicles for drug delivery are the subject of intense research. Compared with nonencapsulated, free drugs, they provide improved biocompatibility, pharmacokinetics, biodistribution, and toxicologic profile. Despite promising results in preclinical models of lung cancer (6) and many other cancer types (7), only a few nontargeted formulations (i.e., relying on passive tumor accumulation through the enhanced permeability and retention effect) have been approved for cancer treatment by regulatory agencies (8, 9). Clinical trials are under way to evaluate novel liposomal formulations of the chemotherapy drugs paclitaxel (LEP-ETU; ref. 10) and cisplatin (Lipoplatin; ref. 11) in patients with lung cancer. Liposomes can deliver large drug payloads but, as numerous clinical studies have shown, this does not always translate into enhanced therapeutic efficacy; results so far show that liposomal drug delivery offers reduced toxicity, but only modest survival benefits, compared with free drug administration (12, 13). In this study we report a novel targeted biotherapeutic agent, saposin C-dioleoylphosphatidylserine (SapC–DOPS) proteoliposomes, with selective antineoplastic activity on lung cancer cells both in vitro and in vivo.

SapC is a small glycoprotein present in all tissues that acts as biologic activator of lysosomal enzymes, such as acid β-glucosidase, acid sphingomyelinase, and acid β-galactosylceramide, which degrade fatty acid substrates resulting in the production of ceramide, a key intermediate in the synthesis of sphingolipids, and a well-known apoptosis inducer (14, 15). SapC has a strong affinity for negatively charged phospholipids such as phosphatidylserine (PS), to which it binds more avidly at acidic pH (16–18). Because the expression of PS is relatively abundant in the surface of tumor cells and tumor-associated vasculatures (19, 20), but not in normal cells, and in view of the characteristically acidic tumor microenvironment that results from glycolytic conversion of glucose into lactate, we hypothesized that SapC–DOPS nanovesicles could be an effective antitumor agent.

Previous work from our group showed that SapC–DOPS targets a variety of cancer cells in vitro and in vivo, and has anticancer effects in animal models of human solid tumors (21–26). In this study, we evaluate the tumor-targeting and cytotoxic potential of SapC–DOPS on human lung cancer cells, and assess its effectiveness in suppressing lung carcinoma growth in vivo. To this end, we first describe the pH dependence of SapC–DOPS cell binding and assess its cytotoxicity in a panel of lung cancer cells. The induction of apoptosis in cultured lung cancer cells is confirmed through DNA fragmentation analysis. Next, we use in vivo fluorescence imaging to assess SapC–DOPS tumor-selective targeting in allogeneic lung tumor mouse models. Finally, the therapeutic efficacy of SapC–DOPS against tumor growth is studied in nude mice bearing human lung cancer xenografts. Our results show that SapC–DOPS effectively targets lung cancer cells in vitro and in vivo, and possesses antitumor activity in a preclinical model of human lung cancer.

The human lung carcinoma A549 and mouse Lewis lung carcinoma (LLC) cell lines were from American Type Culture Collection (ATCC; 1998 and 2001). The human lung carcinoma cell lines SPC-A-1, H1299, NCI-H661, SK-MES-1, NCI-H460, and 801-D were obtained from the Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences from 1999 to 2010. Cells were routinely tested for mycoplasma contamination. No authentication of the cell lines was done by the authors. Human cells were cultured in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS) and 100 μg/mL streptomycin and 100 U/mL penicillin. LLC cells were grown in Dulbecco's Modified Eagle Medium supplemented with 10% FBS, 100 μg/mL streptomycin, and 100 U/mL penicillin. Culture media and FBS were purchased from Life Technologies. All cell lines were cultured at 37°C in a humidified 5% CO₂ atmosphere.

Dioleoylphosphatidylserine (DOPS; Avanti Polar Lipids Inc.) was dried under a stream of N₂ gas, combined with recombinant SapC protein (27) in citrate/phosphate buffer (pH 5.0), and bath sonicated in PBS as described in detail previously (21). To fluorescently label SapC–DOPS nanovesicles, an aliquot of CellVue Maroon (CVM; Molecular Targeting Technologies Inc.) in ethanol was dried together with DOPS before SapC addition and bath sonication. Free CVM was eliminated from this preparation by filtration through a Sephadex G25 column (PD-10; Amersham Pharmacia Biotech). For in vivo treatment, a lyophilized formulation was used: dry DOPS was suspended in 80% tert-butanol. SapC and sucrose (10 mg/mL) were dissolved in water. DOPS suspension and SapC plus sucrose (1:0.6, vol:vol) was lyophilized in a freeze dryer (VirTis Unitop 1000L linked to a Freezemobile 25XL; The VIRTIS). The stable powder cake was resuspended in saline solution to form SapC–DOPS nanovesicles. Vesicles were monitored with a submicron particle size analyzer (Coulter Model N4 Plus). The molar ratio of SapC:DOPS was 1:7.

To evaluate the influence of pH on the binding of SapC–DOPS to tumor cells, SapC–DOPS–CVM (containing 50 μg/mL SapC) or free, equimolar CVM were dissolved in complete RPMI-1640 medium with different pH values (adjusted by addition of NaOH or HCl). A549 cells were collected by trypsinization, washed twice with PBS, and incubated in media adjusted to pH 5.5, 6.6, 7.5, or 8.5 for 2 hours at 37°C. The cells were then washed twice with PBS and analyzed by flow cytometry (FACSCanto II; BD Biosciences), using excitation at 633 nm and a 760 nm band-pass filter to detect CVM emission.

The colorimetric Cell Counting Kit-8 (CCK-8) assay (Dojindo Laboratories) was used to monitor the cytotoxic effects of SapC–DOPS. Lung carcinoma cells were plated at a density of 5,000 cells per well in 96-well plates in triplicate. After 12 hours in culture, cells were treated with SapC–DOPS at final SapC concentrations of 0 (control), 0.5, 1, 2, 4, 8, 16, 32, 64, or 128 μg/mL. After 72 hours, 10 μL CCK-8 was added to each well. Optical density (OD) values were obtained from absorbance readings at 450 nm. Viability was calculated as the percentage of viable cells compared with control using the formula: growth inhibition (%) = (ODC − ODT)/ODC × 100, where ODC and ODT are the OD values of the control and treated samples, respectively. IC₅₀ was calculated by linear interpolation using the formula: IC₅₀ = [(50 − A)/(B − A)] × (D − C) + C, where A is the first point on the curve, expressed as percent inhibition, that is less than 50%; B is the first point on the curve, expressed as percent inhibition, that is greater than or equal to 50%; C is the concentration of inhibitor that gives A% inhibition; and D is the concentration of inhibitor that gives B% inhibition (28).

Membrane phosphatidylserine (PS) levels were measured using Annexin V-FITC (Invitrogen) according to the manufacturer's instructions. In brief, cells were trypsinized, washed with PBS, and resuspended at 2,500,000 cells/mL in Annexin V binding buffer. Aliquots of cells (0.2 mL/tube) were incubated with 10 μL of Annexin V-FITC for 15 minutes at room temperature in the dark. Then, 800 μL of binding buffer was added to each tube. Samples were kept on ice and analyzed by flow cytometry within 1 hour.

Analysis of DNA fragmentation and mitochondrial membrane potential (Δψ_m) measurements were carried out in A549 cells cultured at a density of 5 × 10⁵ cells per well in 6-well plates. Twenty-four hours after plating, cultures were treated with SapC–DOPS (containing 0, 10, 20, or 40 μg/mL of SapC) for 48 hours. To identify apoptotic cells by DNA fragmentation analysis, cells were harvested, washed with PBS, and fixed in 70% ice-cold ethanol overnight. Fixed cells were incubated with 20 U/mL RNase I and 50 μg/mL propidium iodide (PI) for 30 minutes. PI signal was quantified by flow cytometry and cells in the sub-G₀/G₁ phase of the cell-cycle were identified as apoptotic.

Changes in Δψ_m were monitored using the lipophilic cationic probe 5,5′,6,6′-tetrachloro-1,1′,3,3′- tetraethyl-benzimidazolcarbocyanine iodide (JC-1). After treatment for 48 hours, the cells were trypsinized, washed with PBS, and incubated with JC-1 as per the manufacturer's instructions at 37°C in the dark for 30 minutes. Δψm was measured by flow cytometry by exciting with a 488 nm laser line and measuring emitted fluorescence with the FL1 (green) and FL2 (red) channels.

All procedures involving animals were approved by the Ethics Committee for Animal Research of Nanjing University. Murine lung tumor cells (Lewis lung cells, LLC; 5 × 10⁵ cells) were injected into 4 week-old BALB/c female nude mice subcutaneously or into FVB male mice through the tail vein. Sham animals received a PBS injection. At day 10, 200 μL SapC–DOPS–CMV, DOPS–CVM, CVM, or PBS in normal saline were intravenously injected into tumor-bearing mice. Two to 150 hours later, mice were anesthetized to effect with 2% isoflurane and imaged with an IVIS 200 imaging system (PerkinElmer).

Healthy female BALB/c nude mice (5–6 weeks old) were obtained from Vital River Lab Animal Co., Ltd., Beijing Laboratory Animal Research Center (Beijing, China), and were housed in specific pathogen-free conditions at Drum Tower Hospital Animal Center, Nanjing, China with controlled temperature (21 ± 2°C), humidity (55 ± 5%), and light (12 hour light/dark cycle). Water was available to mice ad libitum. After the mice had acclimated for 2 weeks, 2 × 10⁶ A549 cells in 0.1 mL PBS were injected subcutaneously into their dorsal right flanks. Mice were monitored for solid tumor growth for approximately 10 days, and when the tumor volume reached 100 mm³, they were randomly assigned to 5 groups (n = 6/group). The mice then received intravenous injections (0.2 mL) every other day for 5 weeks as follows: group 1 (control, normal saline), group 2 (cisplatin 4 mg/kg), group 3 (high-dose, SapC–DOPS 11.8 mg/kg), group 4 (medium-dose, SapC–DOPS 5.9 mg/kg), and group 5 (low-dose, SapC–DOPS 2.9 mg/kg). Tumor dimensions were measured every other day during treatment. After the treatment regimen was completed, the mice were sacrificed and the tumors were removed, measured, and weighed. Tumor volume (in mm³) was calculated using the formula: L × W² × 0.5. The inhibition of tumor growth was calculated as % inhibition = [1 − (average tumor weight for each treatment group/average tumor weight for the control group)] × 100%. Apoptosis in paraffin-embedded tumor samples was evaluated using a TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling) assay (Boster Bio).

A t test analysis or two-way ANOVA/Tukey test was used to determine statistical significance for experiments with two or more groups, respectively. Analyses were done with SPSS 12.0. Data are presented as the arithmetic mean ± SEM.

Acidosis, resulting from high glycolytic activity and impaired perfusion, is a prominent feature in the microenvironment of solid tumors. Because the binding of SapC to anionic phospholipids such as PS is favored by low pH, we tested whether a weakly acidic environment could promote the binding and internalization of SapC–DOPS in lung cancer cells in vitro. For these experiments, we chose A549 lung adenocarcinoma cells, which in addition to expressing high levels of surface PS (see next section), are among the best characterized and most widely used lung tumor cell lines. A549 cells were incubated with SapC–DOPS–CMV nanovesicles in culture media with different pH values (5.5, 6.6, 7.5, or 8.5), at 37°C for 2 hours. Cell-bound CVM fluorescence intensity was then assessed by flow cytometry (Ex 633 nm/Em 760 nm). The results depicted in Fig. 1A and quantified in Fig. 1B show that SapC–DOPS–CMV targeting was inversely correlated with pH, i.e., binding was higher as media pH decreased.

To evaluate the cytotoxic effects of SapC–DOPS on lung tumor cells, viability assays were performed in vitro in a panel of human lung cancer cell lines. Cells were incubated with SapC–DOPS at concentrations ranging from 0 to 128 μg/mL and 72 hours later, the effects on cell growth were measured using the CCK-8 assay. Figure 2 shows the calculated IC₅₀ values plotted as a function of cell surface membrane PS levels (low PS vs. high PS; cutoff = 52%), as assessed with annexinV-FITC staining. Similar to previous observations in glioblastoma (23) and pancreatic cancer cell lines (24), the selectivity of SapC–DOPS toward PS was evidenced by a significant, inverse correlation between IC₅₀ values and surface PS in these lung cancer cell lines.

To assess whether the loss in cell viability caused by SapC–DOPS was associated with the induction of apoptosis, two relevant apoptosis parameters, namely mitochondrial membrane potential (Δψm) and DNA fragmentation, were evaluated by flow cytometry in A549 cells. Δψm was assessed with the fluorescent dye JC-1. When living cells are incubated with JC-1, the dye is incorporated into the membranes of healthy, polarized mitochondria, forming aggregates that emit red (∼590 nm) fluorescence upon excitation with a 488 nm laser. During early apoptosis, the collapse of Δψm induces dissociation and release of JC-1 monomers to the cytosol, causing a fluorescence emission shift to green (∼529 nm) fluorescence. Treatment of A549 cells with SapC–DOPS (0, 10, 20, or 40 μg/mL) for 48 hours induced a dose-dependent shift from red to green fluorescence, reflecting a collapse of Δψm consistent with apoptotic cell death (Fig. 3A).

In parallel experiments, we analyzed the effects of SapC–DOPS on cell-cycle progression and apoptotic cell death by quantifying the amount of DNA-bound PI representing DNA content. As DNA is fragmented in apoptotic cells, PI fluorescence intensity is lower than in intact cells in the G₀–G₁ cycle, giving raise to the “sub G₀/G₁” phase. As shown in Fig. 3B, SapC–DOPS incubation caused a dose-dependent increase in the percentage of cells in sub G₀/G₁ phase (M1). Figure 3C shows summary data for these experiments. Taken together, the dose-dependent loss of Δψm and the DNA fragmentation observed in these experiments demonstrate that SapC–DOPS induces apoptosis in A549 lung cancer cells.

The tumor-targeting capability of SapC–DOPS was tested in mouse allogeneic tumor models generated through subcutaneous implantation or systemic injection of LLC cells into female BALB/c nude mice or male FVB mice, respectively. Ten days after tumor implantation, animals were injected into the peritoneal cavity or via tail vein with SapC–DOPS–CVM or DOPS-CVM nanovesicles, CVM alone, or PBS; in vivo imaging was performed 24 hours later. As shown in Fig. 4A (intraperitoneal injections) and Fig 4B (intravenous injections), tumor-specific fluorescence in subcutaneous xenografts was detected only in SapC–DOPS–CVM–injected mice. SapC–DOPS' tumor-targeting capability was also assessed in mice harboring allogeneic lung tumors produced by systemic injection of LLC cells. Figure 4C shows a LLC-tumor-bearing mouse and a sham (PBS-injected) mouse imaged 2 and 150 hours after intravenous injection with SapC–DOPS–CVM. As in the prior model, SapC–DOPS–CVM fluorescence localized specifically to the tumor.

The anticancer efficacy of SapC–DOPS against human lung cancer cells was evaluated in nude mice bearing subcutaneous A549 lung tumors. About 10 days after implantation (when tumors reached ∼100 mm³), animals (6 per group) were treated intravenously with SapC–DOPS, PBS, or the chemotherapy drug cisplatin, and tumor growth was monitored over time. Quantification of tumor burden revealed that SapC–DOPS treatment at all three different doses tested reduced tumor weight by approximately 50% at day 35 (Fig. 5). Cisplatin, an alkylating chemotherapy drug used here as a positive control for tumor growth inhibition (29), was also effective in reducing tumor growth.

Consistent with our observations in lung tumor cells in vitro, TUNEL staining confirmed the presence of apoptotic cells in SapC–DOPS–treated subcutaneous A549 lung tumors (Fig. 5C).

Lung cancer is the most common and the deadliest form of cancer worldwide. Because most lung tumors are diagnosed only when they reach advanced stages, standard treatments, namely surgical resection, radio- and chemotherapy, are mostly ineffective. Hence, recent efforts, including the development of immunotherapy agents, reflect the urgent need for more effective, specific therapeutic approaches (30–33). The present study shows that SapC–DOPS, a liposomal biotherapeutic agent composed of a lysosomal hydrolase coactivator (SapC) and a phospholipid (DOPS), induces apoptosis in lung tumor cells in vitro and delays the growth of lung cancer xenografts in vivo.

Unlike typical anticancer liposomal formulations, SapC–DOPS possesses intrinsic tumor-targeting and tumoricidal activities, which result from the combined properties of both its lipid phase and its targeting protein. In this sense, SapC–DOPS is not merely a platform for chemotherapy drug delivery, but an anticancer drug in itself. While multiple modifications are being assayed to overcome some common drawbacks of targeted liposomal formulations, such as limited cell internalization, incomplete drug release and lysosomal degradation (7), SapC–DOPS readily targets exposed PS residues on cancer cells' membranes and exploits the cell's own lysosomal machinery to induce tumor toxicity. Importantly, a key feature of liposomes, i.e., the ability to be endowed with enhanced targeting and cytotoxic capabilities by functionalization with different ligands and drugs, is also shared by SapC–DOPS. The basis of SapC–DOPS anticancer actions resides in the affinity of SapC toward PS, a membrane phospholipid which is sequestered in the inner side of the plasma membrane in healthy cells, but is aberrantly exposed on the surface of many different (nonapoptotic) tumor cells (19, 20). This peculiarity has prompted the development of a number of PS-targeting, antibody-based imaging and therapeutic probes, with recent studies using liposomal formulations showing promising results (34, 35). Upon tumor cell binding, SapC–DOPS cytotoxic activity is thought to arise from lysosomal destabilization and leakage, and/or activation of lysosomal fatty acid hydrolases and ceramide accumulation leading to caspase-mediated apoptosis (16, 21, 22, 36).

In this study, we show that SapC–DOPS exerts cytotoxic actions against a variety of human lung carcinoma cells in vitro, and that higher levels of exposed PS correlate with higher killing efficacy by SapC–DOPS (i.e., lower IC₅₀). We reported similar results in glioblastoma (23) and pancreatic cancer (24) cell lines. Conversely, SapC–DOPS' lack of toxicity toward untransformed human cells in vitro has been demonstrated in mammary (MCF-10A) cells, fibroblasts (21), and pancreatic ductal epithelial cells (24).

The binding of SapC to PS is pH dependent, with an estimated pKa of 5.3 (17, 18). The present results show that SapC–DOPS binding to A549 human lung cancer cells in media with different pHs (5.5 to 8.5) was highest at the most acidic pH value tested. Because acidosis, secondary to hypoperfusion, hypoxia and augmented glycolytic lactate production, is a common feature of the microenvironment of solid tumors, SapC–DOPS may be particularly effective to counteract hypoxia-driven malignant progression and to treat tumors that show resistance to antiangiogenic therapies (23).

The present data also indicate that treatment of A549 lung cancer cells with SapC–DOPS induces a dose-dependent decrease in Δψm paralleled by fragmentation of cellular DNA (sub G₀/G₁ peak), consistent with apoptotic cell death.

Using fluorescently labeled nanovesicles, we show that systemically injected SapC–DOPS specifically targets subcutaneously implanted mouse lung tumor allografts, as well as mouse pulmonary tumors in vivo. Finally, the therapeutic efficacy of SapC–DOPS was evidenced by its ability to significantly reduce the growth of xenografted human lung tumors. As shown by TUNEL staining, and consistent with the reported effects of SapC–DOPS on lung tumor cells in vitro, this effect can be attributed, at least in part, to apoptosis induction.

Taken together, the present results suggest that SapC–DOPS is a promising compound with potential clinical applicability for the diagnosis and treatment of lung tumors. Aberrant PS exposure is a common aspect of different cancer types, and we have shown that SapC–DOPS exerts both in vitro and in vivo antitumor actions in mouse models of neuroblastoma (21), glioblastoma (23), squamous cell carcinoma (25), pancreatic cancer (24), and metastatic brain tumors (26). Unlike chemotherapy drugs, SapC–DOPS nanovesicles are well tolerated and nontoxic in preclinical cancer models (21).

In conclusion, our in vivo and in vitro experiments suggest that SapC–DOPS nanovesicles are a promising treatment option for lung cancer, worthy of further clinical development.

X. Qi is a consultant/advisory board member for Bexion Pharmaceuticals. No potential conflicts of interest were disclosed by the other authors.

Conception and design: Y. Hou, X. Qi

Development of methodology: S. Zhao, Z. Chu, Y. Nie, X. Qi

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): S. Zhao, Z. Chu, X. Qi

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): V.M. Blanco, Y. Nie, X. Qi

Writing, review, and/or revision of the manuscript: V.M. Blanco, X. Qi

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S. Zhao, Z. Chu

Study supervision: Y. Hou, X. Qi

The authors thank Dr. S. Abu-Baker and A. Stevens for the editing help.

This work was supported in part by New Drug State Key Project grant number 009ZX09102-205 (to X. Qi), 1R01CA158372 (to X. Qi), and Research Funds/Hematology-Oncology Programmatic Support from University of Cincinnati College of Medicine (to X. Qi).

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.