Simultaneously inactivating Src and AKT by saracatinib/capivasertib co-delivery nanoparticles to improve the efficacy of anti-Src therapy in head and neck squamous cell carcinoma

DU145, PC3, MDA-MB-231, T47D, H1299, and H1661 cells were directly purchased from ATCC and were authenticated by Human STR Profiling Cell Authentication Service. HN6, HN8, HN12, HN13, and HN17 cells were a gift from Dr. W. Andrew Yeudall [12, 13]. All cells were used for experiments before passage 10 and cultured in DMEM medium containing 10% fetal bovine serum at 37 °C in a humidified incubator supplied with 5% CO2.

Antibodies that recognize phosphorylated or total proteins, including Src, p-Src (Y416), AKT, p-AKT (S473), S6, p-S6 (S240/244), ERK1/2, p-ERK1/2 (T202/Y204), FAK, p-FAK (T397), EGFR, p-EGFR (T1068), STAT3, STAT3 (T705) were purchased from Cell Signaling Technology (Beverly, MA). β-actin antibody and D-Luciferin bioluminescent substrate were purchased from Sigma-Aldrich. Saracatinib and capivasertib were purchased from Selleckchem (Houston, TX). Plasmid HA-AKT-DN (K179 M) (Addgene, Plasmid #16243) and HA-AKT-CA (Addgene, Plasmid #16244) were a gift from Dr. Mien-Chie Hung (Addgene, Plasmid #16243) [14]. Standard cell culture, plasmid transfection, Western blotting, colony formation, CellTiter-Glo® Luminescent Cell Viability and CellTiter 96® AQueous One Solution Cell Proliferation Assay (MTS) assays were carried out as previously described [2, 3, 5].

Luciferase stable HN8 and HN12 cells (HN8-luc and HN12-luc cells) were generated by transduction of pGL4.5 vector (Promoga, Madison, WI) and selection of 150 μg/ml hygromycin (Sigma-Aldrich, St Louis, MO) for 4-5 weeks. Firefly and Renilla luciferase activity was measured by the Dual-Glo^TM Assay System (Promega).

Briefly, 1 × 10⁵ HNSCC cells were seeded into SeedEZ^TM scaffold (Lena Bioscience, Atlanta, GA) pre-coated with Poly-D-Lysine solution (Sigma-Aldrich, St. Louis, MO) [15]. After 10 days of culture, 3D cultures were incubated with free or NP-associated drugs for 12 days. At the endpoint of this experiment, cells in SeedEZ^TM scaffold supplied with complete medium were stained with Texas Red®-X phalloidin (Invitrogen), followed by fluorescence imaging (Zeiss). Cell viability in 3D cultures was quantified by an alamarBlue assay (Bio-Rad).

Briefly, HN8 cells were treated with IC90 dose of saracatinib (20 μM) and maintained in the medium containing IC50 dose of 2 μM for 5 generations. The dose was gradually increased by 1 μM every 2 or 3 weeks until the maximum tolerated dose of 5 μM was reached.

Synthesis of the peptide was carried out using the Fmoc strategy manually in a glass reaction vessel fitted with a sintered glass frit using 2-chlorotritylchloride. Coupling reactions were performed manually by using 2 equiv of N-Fmoc-protected amino acid (relative to the resin loading) activated in situ with 2 equiv of PyBOP and 4 equiv of diisopropylethylamine (DIPEA) in DMF (10 mL/g resin). The coupling efficiency was assessed by the Kaiser test. N-Fmoc protecting groups were removed by treatment with a piperidine/DMF solution (1:4) for 10 min (10 mL/g resin). The process was repeated three times and the completeness of deprotection verified by UV absorption of the piperidine washings at 301 nm. Synthetic linear peptides were recovered directly upon acid cleavage. Before cleavage, the resin was washed thoroughly with methylene chloride. The linear peptides were then released from the resin by treatments with a solution of acetic acid/trifluoroethanol/methylene chloride (1:1:8, 10 mL/mg resin, 2 × 30 min). Hexane (5-10 volumes) was added to the collected filtrates, and the crude peptides were isolated after concentration as white solids. The residue was dissolved in the minimum of methylene chloride and diethyl ether was added to precipitate peptides, followed by triturated and washed three times with diethyl ether to obtain crude materials. Peptide was further purified by preparative HPLC prior to conjugation.

Linear-dendritic mPEG5000-BMA4 containing four branches of amine groups, the cathepsin B (CTSB)-sensitive polymeric drug carrier, was synthesized as described previously [5]. To prepare single drug-loaded NPs, hydrophobic drugs (saracatinib or capivasertib) were loaded into NPs by the solvent evaporation method. Briefly, drug (1.0 mg) and amphiphilic polymer (10 mg) were first dissolved in anhydrous chloroform/methanol (1/1) in a 10 mL round bottom flask. The solvent mixture was evaporated under vacuum to form a thin film. PBS buffer (1 mL) was added to re-hydrate the thin film, followed by 30 min of sonication. Free drugs not associated with the NPs were removed by running the NP solutions through centrifugal filter devices (MWCO: 3.5 kDa, Microcon®). The drug-loaded formulation on the filters were recovered with PBS. To prepare co-delivery NPs (NP-com), saracatinib and capivasertib (1.95 mg, mole ratio = 1:1) were initially dissolved in methanol followed by adding amphiphilic polymer (20 mg in equivalent volume of chloroform). The mixture was transferred into a 10 mL round bottom flask, and the remaining procedure was performed similarly as preparation of single drug-loaded NPs. The amount of drugs loaded in the NPs was analyzed by HPLC (Agilent 1200 LC, Santa Clara, CA). The drug loading was calculated according to the calibration curve between the HPLC area values and concentrations of drug standard. The loading efficiency was defined as the ratio of drug loaded into NPs to the initial drug content. The size and size distribution of the drug-loaded NPs were measured by dynamic light scattering (DLS) instrument three times with an acquisition time of 30 s at room temperature.

The drug released from the single drug-loaded NPs or co-NPs was carried out in the solution with or without CTSB. Cysteine solution in Mcllvaine’s buffer (10 mm) was added in equal volume of enzyme stock solution and pre-incubated for 5 min at 37 °C. The NPs were incubated in the buffer at 37 °C for 48 h in the presence or absence of CTSB (100 nM, pH = 5.4). A drug release control study at physiological condition (without enzyme, pH 7.4) was also performed. At predetermined time points, the samples were withdrawn and analyzed by RP-HPLC (Agilent 1200 LC, Zorbax C18 column 4.6 × 150 mm) with gradient elution.

All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Augusta University. An equal number of male and female six-week-old NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice were purchased from the Jackson Laboratory (Bar Harbor, ME). An orthotopic tongue tumor model was generated as described previously [3]. Briefly, 1 × 10⁵ luciferase-containing HNSCC cells were suspended in 50 μl of PBS/Matrigel (3:1) and injected into anterior ~ 1/3 tongue of NSG mice under anesthesia. For single drug treatment, after 10 days of cell implantation, HN8- or HN12-derived tumor-bearing mice were randomized into three groups to intravenously receive vehicle (PBS), free saracatinib, or Nano-sar once every other day at 10 mg/kg body weight. For combination drug treatment, mice bearing HN12 cells were randomly assigned to one of five groups and treated intravenously once every other day with vehicle (PBS), Nano-sar, Nano-cap, the free-drug combination or Nano-com, at 10 mg/kg body weight. Mice were imaged for bioluminescent luciferase signal using a Xenogen IVIS-200 In Vivo Imaging System (PerkinElmer, Waltham, MA). When experiments were terminated, the primary tongue xenografts and major organs (including the heart, intestine, kidney, liver, lung, and spleen) from mice were excised and processed for standard histological analysis with H&E staining and immunohistochemistry (IHC).

When the animal experiment was terminated, blood was collected via ocular vein for determination of serum Alanine Transaminase (ALT/GPT), Aspartate Transaminase (AST/GOP) and creatinine. ALT and AST were measured by EnzyChromTM Alanine Transaminase Assay Kit and Aspartate Transaminase Assay kits (BioAssay System, Hayward, CA), respectively. Serum creatinine was measured by Creatinine Assay Kit (Cayman chemical, Ann Arbor, MI).

Sections of tongue tumors were immunostained with antibodies against phospho-Src, phospho-AKT, phospho-S6, and Ki67 as described previously [3, 5]. Negative controls included non-specific polyclonal rabbit antibody at 2 μg/ml (Abcam, Cambridge, MA). The sections were developed with the diaminobenzidine tetrahydrochloride (DAB) substrate kit (Vector Laboratories) and counterstained with hematoxylin.

For in vitro analysis, apoptosis was determined by flow cytometry using Annexin V: PE Apoptosis Detection Kit (Invitrogen, Carlsbad, CA) with 7-AAD. For in vivo analysis, apoptosis was assessed by staining for DNA fragmentation using the DeadEnd^TM Fluorometric TUNEL System (Promega, Madison, WI). All TUNEL-positive cells in each section in different treatments were counted in twenty randomly selective view fields using a fluorescence microscope.

We first evaluated the efficacy of Src inhibitor saracatinib in several types of cancer cells. In this analysis, eight cell lines derived from prostate cancer (PC3 and DU145), breast cancer (MDA-MB-231 and T47D), lung cancer (H1299 and H1611), and HNSCC (HN8 and HN12), were treated with two doses of the Src inhibitor saracatinib for 72 h. CellTiter-Glo® Luminescent Cell Viability assay showed that saracatinib displayed cytotoxicity in all cancer cell lines examined in this study (Fig. 1a). Intriguingly, two HNSCC cell lines, HN8 and HN12, exhibited a distinct response to saracatinib (Fig. 1a). HN8 cells were more sensitive to saracatinib toxicity than other cell lines (Fig. 1a). In contrast, HN12 cells did not display a dose-dependent response to saracatinib, suggesting resistance of HN12 cells to Src inhibition (Fig. 1a). The saracatinib IC50 dose varied among HNSCC cell lines, ranging from < 7 μM in HN6, HN8 and HN13 cells, to > 14 μM in HN17 and HN12 cells (Fig. 1b). Moreover, 10 μM saracatinib induced significantly more cytotoxicity in HN6, HN8 and HN13 cells than HN17 and HN12 cells (Fig. 1c). Based upon these observations, we classified HN6, HN8, and HN13 cells as saracatinib-sensitive (Sar-S) cells (IC50 < 10 μM) and HN17 and HN12 cells as Sar-R cells (IC50 > 10 μM). These differences could not be explained by variation in the amount of Src protein; however, HN6, HN8, and HN13 cells showed significantly more Src phosphorylation than HN17 and HN12 cells (Fig. 1d), correlating with sensitivity to saracatinib (Fig. 1c, d).

We sought to understand the mechanism associated with differential saracatinib sensitivity in HNSCC cells. Both of the AKT and ERK1/2 signaling are critical for HNSCC survival and growth, and their cross-talk with Src has been identified, prompting us to determine the effects of saracatinib on these two pathways. Sar-S HN6 and HN8 cells and Sar-R HN12 and HN17 cells were treated with two doses of saracatinib for 24 h. In all four cell lines, saracatinib inhibited Src phosphorylation (Fig. 2a). Saracatinib inhibited phospho-ERK1/2 levels in HN6 and HN12 cells, but not in HN8 and HN17 cells, which did not correlate with the cytotoxicity studies (Fig. 2a). Interestingly, saracatinib markedly impaired the phosphorylation of AKT in Sar-S HN6 and HN8 cells, whereas Sar-R HN12 and HN17 cells showed no or only modest reduction in phospho-AKT (Fig. 2a, b), suggesting that the AKT pathway more relies on Src activation in HNSCC cells carrying high levels of phospho-Src. To address whether blocking AKT signaling increase the response of Sar-R HNSCC cells to saracatinib, HN12 and HN17 cells were transfected with dominant-negative AKT-DN (Fig. 2c). Expression of AKT-DN suppressed the survival of both HN12 and HN17 cells in the absence of saracatinib, as well as led to a more dramatic reduction in cell viability upon saracatinib treatment when compared with cells transfected with an empty vector (Fig. 2d). In contrast, expression of AKT-CA in Sar-S cells, such as HN8, induced a remarkable decrease in the sensitivity of saracatinib (Additional file 1: Figure S1). These data support the notion that AKT activation is tightly associated with the HNSCC response to saracatinib.

Capivasertib is a novel pyrrolopyrimidine-derived compound with great potential to inhibit AKT signaling [16, 17]. In line with a previous study [17], capivasertib induced an increase in AKT phosphorylation, but caused a decrease in AKT kinase activity, as evidenced by a decrease in the phosphorylation of AKT substrate GSK3β and S6 in both HN12 and HN17 cells (Fig. 2e). Saracatinib treatment alone slightly decreased the phosphorylation of AKT and S6 in HN12 cells and had no inhibitory effect on the AKT-S6 signaling in HN17 cells (Fig. 2e). Combined capivasertib with sacaratinib almost completely abrogated S6 phosphorylation (Fig. 2e), leading to a significant reduction in cell viability in both of these Sar-R cell lines compared with each single treatment (Fig. 2f). Consistently, additional capivasertib treatment failed to synergistically enhance saracatinib-induced lethality in Sar-S cell lines HN6 and HN8 (Additional file 2: Figure S2), which may be due to significant co-inhibition of AKT and Src by saracatinib itself in these cells.

It has been reported that EGFR activation can be regulated by the Src and/or AKT signaling in different types of cancer cells [18, 19]. We then determined the status of phospho-EGFR in saracatinib/capivasertib treatment. HN17 expressed lower levels of EGFR compared with HN12 cells, and phospho-EGFR was undetectable by Western blotting (Additional file 3: Figure S3). High levels of phospho-EGFR were detected in HN12 cells, which was inhibited by saracatinib (Additional file 3: Figure S3). Capivasertib increased the levels of phospho-EGFR, and its combination with saracatinib did not produce further suppression in phospho-EGFR compared with saracatinib treatment alone (Additional file 3: Figure S3). We also examined the status of other Src-related proteins, such as FAK and STAT3, in the presence of the indicated treatments. This analysis showed that both STAT3 and FAK, their activation was slightly or not affected in saracatinib treatment (Additional file 3: Figure S3). Although the phosphorylation levels of  FAK were dramatically inhibited in the drug combination in HN17 cells, there was no noticeable changes in phospho-FAK levels in HN12 cells (Additional file 3: Figure S3). These findings further suggest that AKT activation is one of the main mechanisms underpinning the response of HNSCC cells to saracatinib.

To further understand the drug action, we determined whether saracatinib, capivasertib, and their combination promoted apoptosis as cell viability was affected. Flow cytometry assays after Annexin V and 7-AAD staining revealed that apoptosis occurred in all indicated treatment (Fig. 2g, h). However, increase of apoptosis was much efficient in the drug combination compared with each drug alone (Fig. 2g and h). Moreover, the effect of co-treatment on suppression of cell proliferation was much greater compared with the single drug effect (Fig. 2i). Taken together, these observations strongly suggest that AKT blockade has the ability to increase the anticancer activity of saracatinib in HNSCC cells with low saracatinib sensitivity.

To further determine whether activation of AKT signaling is the possible mechanism behind saracatinib resistance in HNSCC cells, we generated Sar-R cells using Sar-S HN8 cells (Fig. 3a) as described in Materials and Methods. Compared with the parental cells, increased phospho-AKT, but not phospho-ERK1/2 was found in Sar-R cells (Fig. 3b). Phospho-AKT was attenuated by AKT-DN transfection in Sar-R cells (Fig. 3c), where cell viability was significantly decreased in the tolerated doses of saracatinib seen in cells transfected with an empty vector (Fig. 3d). We then treated Sar-R cells with 5 μM capivasertib, which induced a strong suppression on phospho-S6 (Fig. 3e). Compared with saracatinib treatment alone, saracatinib combined with capivasertib dramatically decreased the viability and 3D cell growth of Sar-R cells (Fig. 3f, g). These findings suggest that inactivation of AKT has the potential to overcome saracatinib resistance in HNSCC cells.

A comparison between free and NP-associated saracatinib in HN8 and HN12 cells showed that the two preparations had similarly profound effects on phospho-Src and -AKT (Fig. 4a, h) and cytotoxicity (Fig. 4b, i), without differences in 2D colony formation (Fig. 4c, j). However, Nano-sar inhibited cell growth in the 3D culture system more efficiently than free drug (Fig. 4d, k). This comparison reveals differences in drug responses between 2D and 3D cultures.

For in vivo studies, we used a more appropriate orthotopic tongue tumor model by injection of luciferase-containing cells into the anterior portion of NSG mouse tongue. Tongue xenografts established 10 days after cell inoculation, followed by intravenous treatment with vehicle, free or NP-associated saracatinib. In HN8-derived tumor-bearing mice, treatment with either free or NP-associated saracatinib resulted in significantly smaller tumor xenografts relative to those found after vehicle treatment as measured by bioluminescence, tumor size and tumor weight (Fig. 4e, f and Additional file 4: Figure S4). In line with the in vitro 3D cell growth results, tumor growth was suppressed more significantly in mice treated with Nano-sar than those treated with free saracatinib (Fig. 4e, f and Additional file 4: Figure S4). Moreover, systemic toxicity was not seen in mice receiving various treatments, as all groups maintained similar body weight (Additional file 4: Figure S4) 1 and showed no evidence of damage to major organs (Additional file 5: Figure S5). The analysis of xenografts by IHC for phospho-Src and phospho-AKT revealed the strong staining in tumors derived from vehicle-treated mice, with a high proliferative index as measured by the proportion of Ki67-positive tumor cells (Additional file 6: Figure S6). In contrast, we found less staining of phospho-Src and -AKT, as well as lower percentages of Ki67-positive cells, in tumors derived from mice treated with either free or NP-associated saracatinib (Fig. 4g and Additional file 6: Figure S6), suggesting that Nano-sar has the great potential to induce tumor shrinkage and inactivate targeting proteins than free drug. While in HN12-derived tumor-bearing mice, the xenografts showed only marginal response to saracatinib and less than 25% of reduction in the treatment with NP-associated saracatinib on Day 21 (Fig. 4l, m and Additional file 4: Figure S4). Results from IHC analysis showed that the phosphorylation of AKT did not affected by saracatinib regardless encapsulated with NPs or not (Fig. 4n). Together, our findings support the conclusion that saracatinib cannot effectively suppress head and neck tumor growth partially due to insufficiently inhibiting AKT activation in these tumor cells.

The majority of therapeutic agents, including saracatinib and capivasertib, are hydrophobic in nature and are easily incorporated into NPs via hydrophobic interaction during the NP assembly. In this study, biocompatible and amphiphilic polymers with ideal features for drug co-delivery were used to encapsulate both saracatinib and capivasertib into NPs. A short peptide of GFLG linker was incorporated to ensure that the drugs can be released upon the lysosomal enzyme, CTSB (Fig. 5a). DLS study showed that the size of Nano-sar, capivasertib-loaded NPs (Nano-cap) and the saracatinib/capivasertib co-delivery NPs (Nano-com) were 59.6, 58.3, and 83.1 nm, respectively (Fig. 5b). As expected, Nano-com exhibited enzyme-response drug release profile, as evidenced by accelerated release of both drugs (over 80% drug release in 48 h) from NPs at pH 5.4 in the presence of CTSB (Fig. 5c). In contrast, less than 20% of the drugs was released at pH 7.4 or in the absence of CTSB (Fig. 5c), indicating that drugs encapsulated into NPs can be selectively released with the cleavage by CTSB in an acidic environment. As such, Nano-com were formulated to achieve high entrapment efficiency, small size, and controlled release behavior.

We determined the in vitro effects of single and dual drug-loaded NPs in HN12 cells. Similar to the function of free drugs, Nano-sar and Nano-cap decreased the levels of phospho-Src and phospho-S6, respectively (Fig. 5d). When HN12 cells were treated with Nano-com, both Src and S6 were inactivated (Fig. 5d), leading to improved inhibitory effects on colony formation and 3D growth as compared with single nano-drug treatments (Fig. 5e, f).

These exciting in vitro data encouraged us to evaluate the therapeutic efficacy of Nano-com in mice. In in vivo study, HN12-derived tumor-bearing NSG mice received vehicle, Nano-sar, Nano-cap, free or NP-associated drug combination every other days for four times (Fig. 6a). After treatment, tumor burden was reduced in each single-arm treatment as evidenced by lower bioluminescent signal (Fig. 6b), smaller tumor size (Fig. 6c), and decreased tumor weight (Fig. 6d). Combination of saracatinib with capivasertib achieved superior anticancer effects than either treatment alone (Fig. 6b–d). Most importantly, the reduction in tumor growth was more significant in mice receiving Nano-com than those receiving free drug combinations (Fig. 6b–d). Our data show that at least in a preclinical model, the nano-based co-delivery system that incorporates saracatinib and capivasertib is a very efficient and effective method to target Sar-R HNSCC.

A rational drug interaction should increase the benefit of the administered drugs without increasing side effects. To test the hepatotoxicity and nephrotoxicity profiles of NP-associated drugs, we measured serum ALT, AST and creatinine. AST, ALT and creatinine levels were significantly elevated in mice receiving free drug combination (Additional file 7: Figure S7), suggesting that combination of saracatinib with capivasertib would induce systemic toxic. In contrast, biochemical tests revealed no liver or kidney toxicity in the NP-associated combined drug treatment group (Additional file 7: Figure S7). Moreover, H&E staining showed no morphologic changes in the major organs (Fig. 6e). These observations indicate that nano drugs we developed do not produce detectable systemic toxicities.

To further evaluate the effects of treatments on the tumor xenografts, sections were immunostained with the antibodies against phospho-Src, phospho-S6 and Ki67. Consistent with the in vitro data (Fig. 5d), Nano-sar decreased phospho-Src, but had no effect on S6 activation, while Nano-cap treatment significantly abrogated S6 phosphorylation without affecting Src activation (Fig. 7a, b). Both phospho-Src and phospho-S6 levels were significantly reduced in xenografts from mice treated with Nano-com when compared with monotherapy (Fig. 7a, b). Moreover, Nano-com suppressed Src and S6 activation in xenografts more than did the free-drug combination (Fig. 7a, b). Moreover, TUNEL assays revealed that Nano-com was more robust in the induction of tumor cell apoptosis than either single-arm treatment or non-NP based drug combination, as evidenced by the observation of the highest number of TUNEL-positive cells in the group with Nano-com treatment (Fig. 7c, d). Collectively, our study provides evidence that in a preclinical orthotopic model, co-delivery of capivasertib and saracatinib by tumor-targeting NPs is safe and highly effective.