CXCR4 inhibition in tumor microenvironment facilitates anti-programmed death receptor-1 immunotherapy in sorafenib-treated hepatocellular carcinoma in mice

Cells

We used the murine HCC cell line HCA-1¹³ and human HCC cell lines JHH-7 and Hep3B (ATCC) (Supporting Information).

HCC Models

Orthotopic implantation was performed as described (Supporting Information).13 We also induced hepatocarcinogenesis and liver fibrosis in mammalian sterile 20-like 1 (Mst1^–/–Mst2^F/–) mice, as described (Supporting Information).13, 23

Imaging

The growth of intrahepatic HCCs was evaluated by high-frequency ultrasound imaging (VisualSonics, Toronto, Canada) by operators blinded to the treatment (Supporting Fig. S1). The presence of abdominal metastasis was defined as extrahepatic tumor mass infiltrating neighboring organs, significant lymph node enlargement, or peritoneal carcinomatosis. Presence and number of lung metastases were recorded for the different treatment groups as assessed macro- and microscopically in both lungs after killing the animals.

Treatment

The maximal tumor diameters were measured in two dimensions to calculate tumor volume by the formula (a/2 × b/2 × b/2) × 4/3 × Pi. Tumor dimensions were either measured by calipers at the experimental end point (after 2 weeks of treatment) or by ultrasound starting 1 week after implantation, where tumor size was typically 3 × 3 mm (corresponding to a tumor volume of ∼14 mm³). At that time, mice were randomized according to tumor size to the treatment groups to ensure similar tumor sizes at baseline (pretreatment) when treatment was started. Treatment included daily gavage with phosphate-buffered saline/vehicle (control group) or sorafenib at a dose of 50 mg/kg (in phosphate-buffered saline/1% Tween80; MGH Pharmacy) and with the CXCR4 inhibitor AMD3100 (10 mg/kg body weight, subcutaneously by osmotic minipumps; Sigma). In separate experiments, these treatments were combined with antibody against murine PD-1 (100 µg intraperitoneally every 3 days, for a total of five times).

Cell Viability Assays

Cell viability was assessed using the nucleic acid stain SYTO-60 (Invitrogen). Cells (3000 per well) were seeded into 96-well plates, allowed to adhere overnight, and exposed to a range of drug concentrations. After 72 hours, cells were fixed in 4% formaldehyde and stained with SYTO-60 and cell viability was assessed by fluorescence measurement using a SpectraMax M5 microplate reader (excitation 630 nm, emission 695 nm; Molecular Devices). The fraction of control was calculated by dividing the fluorescence obtained from the drug-treated cells by the fluorescence obtained from the control (no drug)–treated cells.

Assessment of Apoptosis by Terminal Deoxynucleotidyl Transferase–Mediated Deoxyuridine Triphosphate Nick-End Labeling Staining and Cleaved Caspase-3 Staining

Frozen sections of HCC tissues were stained using the TACS TdT Kit (R&D Systems, Minneapolis, MN) according to the manufacturer's protocol. Apoptotic cells were counted in four randomly selected visual fields for each sample. The apoptotic index was calculated as the fraction of apoptotic nuclei.

Immunohistochemistry and Immunofluorescence

For evaluation of vascular density, frozen tumor sections (7-8 µm thick) were immunostained with primary antibodies against CD31 (for endothelial cells) and counterstained with fluorescent secondary antibodies. Samples were imaged by using an Olympus confocal microscope and quantified using four or five random fields per sample. Pimonidazole (Hypoxyprobe; Hypoxyprobe, Inc.) was used as a marker of hypoxia. Hypoxyprobe (60 mg/kg body weight) was injected intraperitoneally 30 minutes before the animals were killed. Tumors were harvested and processed immediately. Hypoxia was assessed in frozen tissue sections by immunofluorescence staining of pimonidazole by anti-Hypoxyprobe-fluorescein isothiocyanate (FITC)–labeled antibody.

To evaluate the number and distribution of CD8⁺ T lymphocytes, frozen sections of HCC samples were stained for CD8⁺ cells and nuclear staining with 4′,6-diamidino-2-phenylindole. Whole tumor sections were scanned using mosaic confocal microscopy. The number of CD8⁺ T lymphocytes was recorded both in the tumor margin (200 µm) and in the tumor center (three tumor samples per group were assessed, six random regions were selected both for tumor margin and for tumor center: four groups × three mice × six peripheral × six central tumor regions). In addition, we performed immunostaining for CD8 and PD-L1/CD274 in human HCC samples (n = 16), obtained under an approved institutional review board protocol. We evaluated the number and distribution of intratumoral CD8⁺ T lymphocytes (margin versus center) in a similar manner.

Flow Cytometry

Tumor tissues were harvested and prepared for flow cytometry as previously described (Supporting Information).12 Flow-cytometric data were obtained using an LSR-II flow cytometer (Becton Dickinson) and analyzed with FACSDiva software. We used the following monoclonal anti-mouse antibodies: CD4-FITC, CD4-PE-Cy7, CD8a-FITC, CD8a-PE, CD45-PE, CD45-PE-Cy7, CD25-APC-Cy7, FoxP3-APC, Gr-1-APC, and CD11b-APC-Cy7 (BD Biosciences), as well as F4/80-FITC and F4/80-PE (eBioscience) (see Supporting Fig. S2).

Western Blotting

We used antibodies for PD-L1 (Abcam, Cambridge, MA) to measure the levels of this immune checkpoint in HCC tissues and HCA-1 cells (Supporting Information).

Quantitative Reverse-Transcriptase Polymerase Chain Reaction

Total RNA was extracted with the RNeasy Mini-Kit (Qiagen) from flow cytometry–sorted viable TAMs (7AAD^–CD45⁺F4/80⁺Gr-1^– cells). Complementary DNAs were synthesized using the TaqMan RT Kit (Applied Biosystems, Branchburg, NJ). We used specific primers for β-actin, arginase-1, interleukin-1β (IL-1β), IL-10, IL-12, tumor necrosis factor-α (TNF-α), inducible nitric oxide synthase, chemokine C-C motif ligands 17 and 22 (CCL17, CCL22), CXCR4, chemokine C-X-C motif ligand 10 (CXCL10), matrix metalloproteinase-9, and transforming growth factor-β and determined the relative level of gene expression using the Real-Time SBYR Green PCR master mix (Applied Biosystems) on a Stratagene Mx3000P qPCR System, as described.12

Protein Measurements Using a Multiplexed Enzyme-Linked Immunosorbent Assay

Frozen tumors of each group were prepared using a lysis cocktail buffer (radioimmunoprecipitation assay buffer; Boston Bioproducts, Inc., Boston, MA) including phosSTOP, cOmplete (Roche, Indianapolis, IN). Tumors were homogenized with a tissue homogenizer (Qiagen, Valencia, CA), and the suspensions were sonicated using a sonicator (Branson, Notre Dame, IN). All samples were prepared at a protein concentration of 2 µg/mL. The proinflammatory Panel 1 Mouse V-PLEX kit was used to measure the concentration of several cytokines: IL-1β, IL-2, IL-6, IL-8, IL-12, interferon-γ (IFN-γ), TNF-α, and KC-Gro (Meso Scale Discovery, Rockville, MD).

Statistical Analysis

All statistical analyses were performed by Graph Pad Prism software. The Student t test or the Mann-Whitney U test was used according to data distribution. Numbers of animals (proportions) with metastases were compared by chi-squared test. P < 0.05 was considered statistically significant.

CXCR4 Inhibition During Sorafenib Treatment Has Antivascular and Antimetastatic Effects and Delays HCC Progression

Blockade of the SDF1α/CXCR4 axis by continuous infusion of AMD3100 using osmotic pumps significantly delayed the primary growth of orthotopic HCA-1 tumors in immunocompetent C3H mice when combined with daily sorafenib treatment compared to either treatment alone (Fig. 1A). In the HCA-1 HCC model, mice develop spontaneous lung metastases that are detectable 28 days after intrahepatic tumor implantation. Despite delaying the primary tumor growth, sorafenib did not affect metastasis formation or overall survival compared to control-treated mice (Fig. 1B,C). In contrast, the addition of AMD3100 treatment significantly inhibited lung metastasis and led to an increase in overall survival (Fig. 1B,C). The tumor growth delay was seen after combination therapy was reproduced in orthotopic human HCC xenograft models using Hep3B and JHH-7 cells (Supporting Fig. S3). Of interest, tumor growth inhibition occurred in the face of minor direct effects of recombinant SDF1α or AMD3100 on HCC or endothelial cell proliferation after sorafenib treatment in vitro (Supporting Fig. S4). However, sorafenib treatment significantly reduced intratumoral microvascular density and increased hypoxia and SDF1α expression, and addition of AMD3100 to sorafenib enhanced the antivascular effects of treatment (Supporting Fig. S5). Moreover, we found that CXCR4 inhibition prevented the increase in epithelial-to-mesenchymal transition (EMT) markers in HCC cells cultured in hypoxic conditions, in a dose-dependent manner (Supporting Fig. S6). Thus, the inhibition of HCC progression induced by sorafenib and AMD3100 in these HCC models is at least in part due to tumor microenvironment–mediated effects.

CXCR4 Inhibition Prevents the Polarization Toward an Immunosuppressive HCC Microenvironment During Sorafenib Treatment

We next examined the effects of sorafenib treatment on tumor inflammatory cell infiltration by flow-cytometric analyses of enzymatically digested HCC tissue. While the fraction of CD45⁺ immune cells of the total number of viable cells did not change significantly, we found that sorafenib increased the numbers of F4/80⁺ TAMs and CD11b⁺Gr-1⁺ and CD45⁺CXCR4⁺ myeloid cells in both HCA-1 and JHH-7 HCC models (Fig. 2A-C, Supporting Fig. S7). Moreover, sorafenib treatment resulted in an increase in the fraction of tumor-infiltrating CD4⁺CD25⁺FoxP3⁺ Tregs in HCA-1 tumors (P < 0.05) (Fig. 2D). Addition of AMD3100 to sorafenib significantly decreased the fraction of F4/80⁺ TAMs, CD11b⁺Gr-1⁺ myeloid cells, and CD4⁺CD25⁺FoxP3⁺ Tregs in the orthotopic HCA-1 model to levels comparable to those of treatment-naive (control) HCCs (Fig. 2).

Both vascular endothelial growth factor (VEGF) and SDF1α have been reported to mediate the trafficking and retention of tumor-infiltrating myeloid (bone marrow–derived) cell populations (F4/80⁺ TAMs, Gr-1⁺ monocytic and granulocytic populations) in other tumor types and in the normal liver.24-28 However, it is currently unknown how inhibitors of these pathways affect bone marrow–derived cell activation and functional cytokine secretion. To examine this, we sorted F4/80⁺ TAMs using flow cytometry from digested HCC tissue after 14 days of treatment, extracted mRNA, and performed quantitative polymerase chain reaction analysis to measure widely accepted markers of angiogenesis and of M1 and M2 types of activation.8, 29 The TAMs from sorafenib-treated HCCs expressed more than two-fold higher levels of CXCR4 and VEGF as well as the M2-type markers CCL22 and arginase-1 compared to those from control HCCs (Supporting Table S1). On the other hand, the expression of the M1-type markers was not significantly altered in TAMs in sorafenib-treated HCCs (Supporting Table S1). AMD3100 alone or in combination with sorafenib reduced the expression of M2-type markers in TAMs without affecting that of M1-type markers (Supporting Table S1). To examine whether sorafenib and AMD3100 treatments also affect the infiltration and proliferation of T lymphocytes in HCC, we further assessed the fraction of tumor-infiltrating CD4⁺ and CD8⁺ T lymphocytes. Combination treatment with sorafenib and AMD3100 or sorafenib alone did not increase the fraction of CD4⁺ and CD8⁺ T lymphocytes (Fig. 2E,F). Thus, blockade of the SDF1α/CXCR4 axis may inhibit sorafenib-induced immunosuppression in HCC but is insufficient to enhance T-lymphocyte infiltration.

PD-L1 Is Expressed by HCC Cells and Is Increased After Sorafenib Treatment

Hypoxia has been shown to upregulate the expression of immune regulatory proteins such as PD-1/PD-L1, which inhibit cytotoxic CD8⁺ T-lymphocyte proliferation and activation. We next evaluated if treatment-induced hypoxia increases PD-L1 expression in HCA-1 tumors. Immunocompetent mice bearing orthotopic HCA-1 tumors were treated when the tumor reached a size of 14 mm³. Tumors were treated with sorafenib for 28 days, and then the mice were killed and the tumors collected. Western blot analysis showed that sorafenib treatment resulted in increased PD-L1 expression in HCA-1 tumors compared with control treated tumors (Fig. 3A). We next examined the cell types that express PD-L1 in HCC. We found that cultured HCA-1 cells expressed detectable levels of PD-L1 (Supporting Fig. S8A). By flow cytometry of digested tumor tissue, we detected, as expected, expression of PD-L1 on multiple hematopoietic cells, including on a fraction of Gr-1⁺ cells, TAMs, dendritic cells, and lymphocytes (Supporting Table S2). In addition, analysis of resected human HCC tissues showed PD-L1 expression in both cancer cells and immune cells infiltrating the tumor stroma (Supporting Fig. S8B,C). Finally, immunofluorescence analysis of HCA-1 tumors showed colocalization of PD-L1 expression with pimonidazole staining of hypoxic tissue (Supporting Fig. S8D). These data suggest that PD-L1 expression by the cancer cells and HCC stromal cells, particularly in hypoxic regions, may play a role in immunosuppression after sorafenib treatment.

PD-1 Blockade Combined With CXCR4 Inhibition and Sorafenib Delays HCC Growth

We next examined whether blocking PD-1 improves the efficacy of sorafenib and CXCR4 inhibition during persistent hypoxia. To this end, we assessed the efficacy of anti-PD-1 treatment alone or in combination with sorafenib plus or minus AMD3100 (Supporting Fig. S9A). Average body weight was similar across all treatment groups, and no other signs of toxicity were observed. Wound healing (after median laparotomy) was not impaired in any of the treatment groups. As expected, the sorafenib/AMD3100 combination significantly inhibited HCA-1 tumor growth. Moreover, anti-PD-1 treatment alone showed comparable efficacy. Interestingly, combining sorafenib with anti-PD-1 treatment did not show efficacy, while the triple-combination group (sorafenib/AMD3100/anti-PD-1) showed the most pronounced delay in tumor growth versus the control group (Fig. 3B, Supporting Fig. S10A). Furthermore, the number of lung metastases was significantly reduced by anti-PD-1 treatment alone and by sorafenib plus AMD3100, and it was further inhibited by addition of anti-PD-1 treatment (Fig. 3C). Neither sorafenib alone nor anti-PD-1 antibody with sorafenib affected spontaneous lung metastasis formation.

Next, we sought to further confirm the activity of anti-PD-1 therapy in a genetically engineered (Mst-mutant) model of HCC in mice with cirrhotic livers (Supporting Fig. S9B).13 Using ultrasound imaging in this model, we found that anti-PD-1 treatment may stabilize the growth of established HCCs. Moreover, we found that the combination of anti-PD-1 antibody with sorafenib and AMD3100—but not anti-PD-1 antibody with sorafenib alone—led to regression of established tumors (Fig. 3D, Supporting Fig. S10B).

PD-1 Blockade Combined With CXCR4 Inhibition and Sorafenib Increases Tumor Cell Death In Vivo

We next evaluated whether the efficacy seen with addition of anti-PD-1 treatment to sorafenib plus AMD3100 is due to increased tumor cell death. We found that tumors in the triple-combination group had extensive areas of tumor necrosis in the HCA-1 tumors, which were not detectable to a similar extent in any of the other treatment groups (Fig. 4A). We next quantified cell apoptosis and found that the addition of anti-PD-1 to sorafenib and AMD3100 dramatically increased the number of cleaved caspase-3–positive cells in tumors compared to the other groups (Fig. 4B). Similar findings were obtained after analyzing HCC tissues from Mst-mutant mice (Fig. 4C). Thus, adding anti-PD-1 antibody to sorafenib and AMD3100 treatment significantly decreases primary tumor growth by triggering tumor necrosis and apoptosis and further reduces the incidence of lung metastases.

Addition of Anti-PD-1 Treatment to Sorafenib and AMD3100 Increases Intratumoral Penetration and Activation of CD8⁺ T Lymphocytes in HCC

We first examined the pattern of lymphocyte distribution in treatment-naive human HCCs and found significantly reduced numbers of CD8⁺ T lymphocytes penetrating into the tumor proper compared to the tumor margin (Supporting Fig. S11). We next performed a similar analysis as well as flow-cytometric analyses of lymphocytic populations in the murine HCCs. We also evaluated whether adding anti-PD-1 treatment increases or activates antitumor T lymphocytes. We first assessed the percentages of CD8⁺ T lymphocytes (cytotoxic T cells) in HCC tissues from mice in different treatment groups by flow cytometry. We found no difference in the fraction of these cells normalized either to the total number of CD45⁺ tumor-infiltrated leukocytes or to the total number of cells across treatment groups in HCA-1 tumor grafts or HCCs induced in Mst-mutant mice (Supporting Fig. S12). We then examined the distribution of the CD8⁺ T lymphocytes in tumor sections by immunofluorescence staining. Sorafenib alone did not alter CD8⁺ T-lymphocyte number or distribution, and the combination of sorafenib and AMD3100 only slightly increased the number of CD8⁺ T lymphocytes localized deeper within the primary tumors. Addition of anti-PD-1 treatment to sorafenib and AMD3100 significantly increased cell intratumoral penetration by CD8⁺ T lymphocytes in both models (Fig. 5). The tumor-infiltrating CD8⁺ T lymphocytes colocalized with apoptotic cells within the HCCs (Fig. 4B).

To test whether these tumor-infiltrating CD8⁺ T lymphocytes were activated after PD-1 blockade, we evaluated biomarkers of activation, i.e., IFN-γ, IL-2, and TNF-α expression levels. Enzyme-linked immunosorbent assay showed that addition of anti-PD-1 treatment to sorafenib and AMD3100 significantly increased the intratumoral levels of IFN-γ, IL-2, and TNF-α by 50%-130% compared to control (Fig. 6). In contrast, sorafenib or anti-PD-1 alone and the combination treatment of sorafenib plus AMD3100 or sorafenib plus anti-PD-1 did not significantly change IFN-γ, IL-2, and TNF-α expression levels (Fig. 6). Similarly, only triple-combination treatment significantly increased IFN-γ, IL-2, and TNF-α expression levels in genetically induced tumors in Mst-mutant mice (Supporting Fig. S13).