Published online before print July 21, 2008
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,1
* Disease Control and Homeostasis, Graduate School of Medical Science, and
Division of Molecular Bioregulation, Cancer Research Institute, Kanazawa University, Kanazawa, Japan; and
Department of Molecular Preventive Medicine, School of Medicine, University of Tokyo, Tokyo, Japan
1 Correspondence: Division of Molecular Bioregulation, Cancer Research Institute, Kanazawa University, 13-1 Takara-machi, Kanazawa 920-0934, Japan. E-mail: naofumim{at}kenroku.kanazawa-u.ac.jp
ABSTRACT
The first step in the generation of tumor immunity is the migration of dendritic cells (DCs) to the apoptotic tumor, which is presumed to be mediated by various chemokines. To clarify the roles of chemokines, we induced apoptosis using suicide gene therapy and investigated the immune responses following tumor apoptosis. We injected mice with a murine hepatoma cell line, BNL 1ME A.7R.1 (BNL), transfected with HSV-thymidine kinase (tk) gene and then treated the animals with ganciclovir (GCV). GCV treatment induced massive tumor cell apoptosis accompanied with intratumoral DC infiltration. Tumor-infiltrating DCs expressed chemokine receptors CCR1 and CCR5, and T cells and macrophages expressed CCL3, a ligand for CCR1 and CCR5. Moreover, tumor apoptosis increased the numbers of DCs migrating into the draining lymph nodes and eventually generated a specific cytotoxic cell population against BNL cells. Although GCV completely eradicated HSV-tk-transfected BNL cells in CCR1-, CCR5-, or CCL3-deficient mice, intratumoral and intranodal DC infiltration and the subsequent cytotoxicity generation were attenuated in these mice. When parental cells were injected again after complete eradication of primary tumors by GCV treatment, the wild-type mice completely rejected the rechallenged cells, but the deficient mice exhibited impairment in rejection. Thus, we provide definitive evidence indicating that CCR1 and CCR5 and their ligand CCL3 play a crucial role in the regulation of intratumoral DC accumulation and the subsequent establishment of tumor immunity following induction of tumor apoptosis by suicide genes.
Key Words: dendritic cells • gene therapy
INTRODUCTION
Hepatocellular carcinoma (HCC) occurs in individuals with chronic liver disease related to hepatitis B or C virus infections [1 2 3 ]. Even after the curative treatments for HCC, such as surgical resection and radiofrequency ablation, tumor recurrence often occurs because of the multicentric development of HCC in the cirrhotic liver [4 ]. Immune-based therapies, particularly those based on dendritic cells (DCs), may be theoretically effective in preventing the recurrence because of their potential capacity to search for and eradicate tumor cells irrespective of site [5 ]. However, DC-based therapy is still considered to be in its infancy, probably as a result of the lack of effective techniques for enhancing the immune response to human cancer cells including HCC, which are generally poor in immunogenicity.
Apoptotic tumor cells are generally less immunogenic than necrotic cells, but they can sometimes induce efficient antitumor immune responses depending on the type of apoptosis inducer. Indeed, some anticancer drugs can induce apoptosis of tumor cells and simultaneously enhance the immunogenicity of apoptotic cancer cells [6 7 8 ]. Ganciclovir (GCV) can activate the protease family of caspases and induce apoptosis selectively in the cells transfected with the HSV-thymidine kinase (tk) gene [9 , 10 ]. Thus, when GCV is administered systemically to tumor-bearing individuals, it induces apoptosis of HSV-tk-transfected tumor cells but not normal cells. This treatment strategy, designated as suicide gene therapy, can induce immunogenic apoptosis of the tumor cells [11 ], as evidenced by a massive intratumoral infiltration of macrophages and T cells [12 ]. Moreover, the expression of various proinflammatory cytokines is augmented at the tumor sites following GCV treatment [12 , 13 ]. Furthermore, to enhance the suicide gene therapy-induced immune responses, the simultaneous use of cytokines such as GM-CSF, IL-2, and MCP-1/CCL2 has been used with some success [14 15 16 ]. To design more effective methods of preventing tumor recurrences, it is necessary to fully understand the immune responses after tumor apoptosis induced by HSV-tk/GCV suicide gene therapy.
DCs are potent APC that play a crucial role in the establishment of adoptive immune response. Immature DCs capture and process antigens at the inflammatory sites and thereafter migrate to the draining lymph node, where they undergo phenotypical and functional maturation. At the draining lymph node, the mature DCs interact with naïve T cells and present the captured and processed antigen to T cells [17 , 18 ].
Chemokines are presumed to play an essential role in the regulation of DC trafficking and DC–T cell interaction in general [19 20 21 22 ]. Circulating immature DCs express inflammatory chemokine receptors such as CCR1, CCR2, CCR5, and CCR6, and these DCs can reach the source of the inflammatory stimulus under the guidance of the ligand gradient for the expressed receptors such as CCL2, CCL3, CCL4, CCL5, CCL7, and CCL20. After capturing antigens, DCs undergo maturation, resulting in a decrease in inflammatory chemokine receptor expression and a reciprocal increase in CCR7 expression. Mature DCs expressing CCR7 migrate to T cell-rich areas of the draining lymph nodes, where the ligands for CCR7, CCL19, and/or CCL21 are expressed abundantly. However, it still remains elusive whether similar mechanisms operate in the DC migration process following massive tumor apoptosis induced by treatments such as gene therapy, chemotherapy, and radiation therapy.
Here, we demonstrate the induction of specific tumor immunity by tumor apoptosis after HSV-tk/GCV suicide gene therapy and essential roles of DCs in this process. Moreover, we provide definitive evidence to indicate that CCR1 and CCR5 and their ligand CCL3 play a key role in the regulation of intratumoral DC accumulation and the subsequent establishment of tumor immunity following induction of tumor apoptosis by HSV-tk/GCV suicide gene therapy. These observations might lay the foundation for devising novel measures to enhance antitumor immune responses to prevent tumor recurrence.
MATERIALS AND METHODS
Mice
Specific pathogen-free, 7- to 9-week-old male BALB/c mice were purchased from Charles River Japan (Yokohama, Japan) and were designated as wild-type (WT) mice. CCL3-deficient [CCL3 knockout (CCL3KO)] mice were obtained from Jackson Laboratories (Bar Harbor, ME, USA). CCR1KO mice were a gift from Dr. Philip M. Murphy [National Institute of Allergy and Infectious Diseases, National Institutes of Health (NIAID, NIH), Bethesda, MD, USA]. CCR5KO mice were generated as described previously [23
]. All mice were backcrossed to BALB/c mice for eight to 10 generations. All animal experiments were performed under specific pathogen-free conditions in accordance with the Guideline for the Care and Use of Laboratory Animals of Kanazawa University (Japan).
Tumor cell lines
A murine HCC cell line, BNL 1ME A.7R.1 (BNL), was cultured in DMEM (Sigma Chemical Co., St. Louis, MO, USA) containing 10% FBS (Gibco, Long Island, NY, USA). BNL cells were infected with the retroviral vector pG1Sv.Na harboring HSV-tk cDNA. The infected BNL cells were cultured in 10% FBS-containing DMEM in the presence of 400 µg/ml G418 (Gibco). The surviving cells were tested for sensitivity to GCV in vitro as described previously [24
]. GCV-sensitive cells were designated as BNL-tk and were used in the experiments.
Apoptosis detection assay
After culturing for 1 day with 5 µg/ml GCV, BNL-tk cells were harvested, and phosphatidyl serine levels were determined by staining the cells with propidium iodide (PI) and the Annexin V-FITC apoptosis detection kit (Calbiochem, Darmstadt, Germany) according to the manufacturer’s instructions. At least 50,000 stained cells were analyzed on a FACSCalibur system (BD Biosciences, San Diego, CA, USA) for each determination.
Tumor injection
Seven- to 9-week old male WT, CCR1KO, CCR5KO, and CCL3KO mice were inoculated s.c. into the left flank with 2 x 105 BNL-tk cells on Day 0. From Days 14 to 18 (5 consecutive days), 75 mg/kg GCV (i.p.) was administered daily (see Fig. 1C
). Tumors were removed at the indicated time intervals for immunohistochemical analysis and quantitative real-time RT-PCR. In another series of experiments, WT, CCR1KO, CCR5KO, or CCL3KO mice were inoculated with 1.5 x 105 BNL-tk on Day 0. The mice were i.p.-injected with 75 mg/kg GCV from Days 2 to 5. The animals were then rechallenged s.c. with 1.0 x 105 BNL in their right flank on Day 18, after confirming that the primary tumors were eradicated completely (see 
Fig. 5A
). Tumor sizes were evaluated twice each week using calipers, and tumor volume was calculated by the following formula: Tumor volume (mm3) = (the longest diameter) x (the shortest diameter)2/2.
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Figure 1. Apoptosis of the murine hepatoma cell line induced by GCV treatment. After 1 day of culture without (A) or with (B) 5 µg/ml GCV, BNL-tk cells were harvested and stained with the Annexin V-FITC apoptosis detection kit and PI. Then, the cells were analyzed by flow cytometry. Representative results from three individual experiments are shown here. FL1- and -2-H, Fluorescence 1- and 2-height. (C) Schematic representation of GCV treatment in vivo. Mice were s.c.-injected with 2 x 105 BNL-tk cells on Day 0. Then, GCV was i.p.-injected into mice from Days 14 to 18. Tumors were harvested on the day before GCV injection (Day 13), on Day 3 or 6 after GCV injection (Day 16 or 19) for real-time RT-PCR analysis, and on Day 19 for immunohistochemistry (IHC). (D) Apoptotic cells detected in tumor tissues with or without GCV treatment using anti-ssDNA antibody. Original magnification, x400. Original bar, 50 µm.
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Figure 2. Accumulation of tumor-infiltrating CCR1-, CCR5-, or CCL3-positive cells after tumor apoptosis induced by GCV treatment. Mice were inoculated with BNL-tk cells according to the schedule shown in Figure 1C
. (A) Tumors were removed from WT mice on Day 19, and immunohistochemical analysis was performed using anti-CCR1, -CCR5, or -CCL3 antibody on tumors with or without GCV treatment. Representative results from three individual animals in each group are shown here. Original magnification, x400. Original bars, 50 µm. (B) Real-time RT-PCR was performed on total RNA extracted from the tumor of WT mice harvested on Days 13, 16, or 19. The level of chemokine mRNA was normalized to GAPDH mRNA levels. Bars, ±1 SD (n=3); *, P < 0.05, compared with the day before GCV injection.
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Figure 3. Identification of CCR1-, CCR5-, and CCL3-expressing cells in the tumor sites. Mice were inoculated with BNL-tk cells according to the schedule shown in Figure 1C
. (A–C) Tumors were removed from WT mice on Day 19 and immunostained with the indicated combinations of antibodies, as described in Materials and Methods. The digitally merged images are shown in the right panels. Representative results from three individual animals are shown. Original magnification, x400. Original bars, 50 µm.
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Figure 4. Deficiency of CCR1, CCR5, or CCL3 impaired intratumoral accumulation of DCs and T cells after tumor apoptosis induced by GCV treatment. Mice were inoculated with BNL-tk cells according to the schedule shown in Figure 1C
. (A) WT and KO mice were inoculated with 2 x 105 BNL-tk cells. The sizes of primary tumors were measured without GCV treatment and are shown here. Bars, ±1 SE. (B) The numbers of CD3-, CD4-, CD8-, F4/80-, DEC205-, or CD11c-positive cells were enumerated from the tumors harvested from mice with or without GCV. The cell density was determined in 10 randomly chosen tumor areas at 400-fold magnification. Bars, ±1 SD;*, P < 0.05, compared with WT. (C) Immunohistochemical analysis was performed using anti-CD3, anti-DEC205, or anti-CD11c antibody on tumors after GCV treatment in WT and KO mice. Representative results from three individual animals in each group are shown here. Original magnification, x400. Original bars, 50 µm.
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Figure 5. Partial failure of CCR1KO, CCR5KO, and CCL3KO mice in rejecting the rechallenged tumor. (A) Schematic representation of the treatment schedule. Mice were s.c.-injected with 1.5 x 105 BNL-tk cells on Day 0. GCV was i.p.-injected into mice from Days 2 to 5. Following complete eradication of the primary tumors, the mice were s.c.-rechallenged with 1.0 x 105 BNL cells on Day 18. LN, Lymph node. (B) Tumor sizes were measured twice per week and are shown here. Bars, ±1 SE; *, P < 0.05. WT mice, which had been injected with neither BNL-tk cells nor GCV beforehand, were injected with BNL cells and are shown here as control. (C) Mononuclear cells from the draining lymph nodes on Day 8 were incubated at a cell density of 2 x 106 cells/ml in the presence of 0.6 x 106 cells/ml BNL cells, which were irradiated at 50 Gy beforehand. After 5 days of culture, the cells were tested for in vitro cytotoxicity against BNL or CT26 cells as a control, as described in Materials and Methods. Each value represents mean ±1 SE (n=3). (D) Mononuclear cells from draining lymph nodes on Day 8 were incubated for 6 h with the lysates of BNL or CT26 cells and stained with CD8 antibody and IFN- antibody or isotype control as described in Materials and Methods. The cells were analyzed by flow cytometry, and the percentages of CD8+IFN-+ cells of CD8+ cells are indicated in the upper-right corners. Representative results from three individual animals are shown here.
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Immunohistochemical analysis
Rabbit anti-mouse CCR5 polyclonal antibodies were prepared as described previously [25
]. The removed tumor tissues were embedded in paraffin or the Sakura Tissue-Tek OCT compound (Sakura Finetek, Torrance, CA, USA) as frozen tissues. The paraffin-embedded sections were then stained with goat anti-mouse CCR1 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), rabbit anti-CCR5, goat anti-mouse CCL3 (R&D Systems, Minneapolis, MN, USA), rat anti-mouse F4/80, anti-mouse CD3 (Serotec, Oxford, UK), rabbit anti-ssDNA, or rat anti-Ki67 (Dako Cytomation, Tokyo, Japan) overnight at 4°C. Cryostat sections of the frozen tissues were fixed with 4% paraformaldehyde (PFA) in PBS and stained with rat anti-mouse DEC205 (Serotec) or hamster anti-mouse CD11c (BD Biosciences) overnight at 4°C. The sections were then incubated for 1 h at room temperature with biotinylated rabbit anti-goat IgG, biotinylated swine anti-rabbit IgG, biotinylated rabbit anti-rat IgG (Dako Cytomation), or biotinylated mouse anti-hamster IgG (BD Biosciences). The immune complexes were visualized using a catalyzed signal amplification system (Dako Cytomation) or the ELITE avidin-biotin-peroxidase and diaminobenzidine substrate kits (Vector Laboratories, Burlingame, CA, USA), except for anti-ssDNA, where a novel HRP-labeled polymer (Envision+, Dako Cytomation) was used, according to the manufacturer’s instructions. As a negative control, goat IgG (R&D Systems), rabbit IgG (Dako Cytomation), rat IgG (Cosmo Bio, Tokyo, Japan), or hamster IgG (BD Biosciences) was used instead of specific primary antibodies. The numbers of positive cells were determined in each animal in 10 randomly chosen fields at 400-fold magnification by an examiner without any prior knowledge of the experimental procedures.
Double-color immunofluorescence analysis
Tumor tissues were embedded in paraffin or the OCT compound as frozen tissues. The paraffin-embedded sections were then stained with combinations of rat anti-mouse CD3 and goat anti-mouse CCL3 or anti-F4/80 and anti-CCL3 antibodies overnight at 4°C. After fixation with 4% PFA/PBS, cryostat sections were stained with the combinations rat anti-mouse CD4 (BD Biosciences) and anti-CCR1, rat anti-mouse CD8a (BD Biosciences) and anti-CCR1, anti-CD4 and anti-CCR5, anti-CD8a and anti-CCR5, rat anti-DEC205 and anti-CCR1, anti-DEC205 and anti-CCR5, PE-conjugated hamster anti-CD11c (BD Biosciences) and anti-CCR1, PE-conjugated anti-CD11c and anti-CCR5, PE-conjugated anti-CD11c and rat anti-CD11b (BD Biosciences), or PE-conjugated anti-CD11c and anti CD8a antibodies. After extensive washing, AF488 donkey anti-rat IgG (Invitrogen, Carlsbad, CA, USA) was applied as the secondary antibody to detect CD4-, CD8a-, CD3-, F4/80-, DEC205-, or CD11b-positive cells. Simultaneously, AF546 or AF488 donkey anti-goat IgG (Invitrogen) was used to detect CCR1- or CCL3-positive cells, and AF594 or AF488 donkey anti-rabbit IgG (Invitrogen) was used to detect CCR5-positive cells. The sections were observed using a confocal microscope (LSM 510 META, Zeiss, Thornwood, NY, USA). The percentage of double-positive cells was determined in each animal in five randomly chosen fields at 400-fold magnification by an examiner without any prior knowledge of the experimental procedures.
Flow cytometric analysis
Inguinal and axillary lymph nodes were removed and digested in a DNase I and collagenase solution (Sigma Chemical Co.). The resultant, single-cell preparations were stained with various combinations of FITC-labeled anti-CD4, FITC-labeled anti-CD86, PE-labeled anti-CD8, PE-labeled anti-CD11c, PE-labeled anti-CD44, and PE-labeled anti-CD62 ligand (CD62L) mAb (BD Biosciences). FITC-rat IgG, PE-hamster IgG, and PE-rat IgG were used as isotype controls (BD Biosciences). To prepare the tumor lysate, BNL or CT26 cells were suspended in PBS and subjected to four cycles of rapid freezing in liquid nitrogen and thawing at 55°C. The lysate was spun at 15,000 rpm to remove particulate cellular debris. To stain intracellular IFN-, the mononuclear cells harvested from the draining lymph nodes on Day 8 (see Fig. 5A
) were incubated with the BNL or CT26 lysates at a tumor cell:mononuclear cell ratio of 1:1 in the presence of GolgiPlug (BD Biosciences). Six hours later, surface staining was performed with APC-conjugated CD8 antibodies. Intracellular IFN- was stained after fixation and permeabilization with BD Cytofix/Cytoperm buffer with PE-conjugated IFN- antibodies or isotype control using the Mouse Intracellular Cytokine Staining starter kit (BD Biosciences). At least 100,000 stained cells were analyzed on a FACSCalibur system for each determination. The data were expressed as a proportion of positive cells (compared with cells stained with an irrelevant control antibody), and the absolute positive cell numbers were calculated after determining the total cell numbers in the lymph nodes by the following formula: Absolute positive cell numbers = total cell number in the lymph nodes x percentage of positive cells x 1/100.
Quantitative real-time RT-PCR
Total RNA was extracted from the resected tumor and lymph nodes using RNA-Bee (Tel-Test, Friendswoods, TX, USA), according to the manufacturer’s instructions. After the RNA preparations were further treated with RNase-free DNase I (Life Technologies, Gaithersburg, MD, USA) to remove residual DNA, cDNA was synthesized as described previously [26
]. Quantitative real-time PCR was performed on an Applied Biosystems StepOneTM real-time PCR system (Applied Biosystems, Foster City, CA, USA) using the comparative threshold (CT) quantification method. TaqMan® gene expression assays (Applied Biosystems) containing specific primers (Accession Numbers CCL3, Mm00441258_ml; CCL4, Mm00443111_m1; CCL5, Mm01302428_ml; CCR1, Mm00438260_s1; CCR5, Mm01216171_m1; GAPDH, Mm99999915_g1), TaqMan® minor groove binder probe (FAMTM dye-labeled), and TaqMan® fast universal PCR master mix were used with 10 ng cDNA to detect and quantify the expression levels of CCL3, CCL4, CCL5, CCR1, and CCR5. Reactions were performed for 20 s at 95°C and then for 40 cycles of 1 s at 95°C and 20 s at 60°C. GAPDH was amplified as an internal control. CT values of GAPDH were subtracted from CT values of the target genes (
CT). CT values of tumors after GCV injection were compared with CT values of tumors before GCV injection.
Cytotoxicity assay
Mononuclear cells were isolated from the draining lymph nodes at the indicated time intervals and were incubated at a cell density of 2 x 106 cells/ml in the presence of 0.6 x 106 cells/ml BNL cells, which were irradiated at 50 Gy beforehand. After 5 days of culture, the cells were tested for cytotoxicity in a lactate dehydrogenase assay using the CytoTox 96 nonradioactive cytotoxicity assay kit (Promega, Madison, WI, USA), according to the manufacturer’s instructions. Effector cells were added to target cells in triplicate at different E:T ratios. Percentage of specific lysis was calculated using the following formula: [(experimental–effector spontaneous–target spontaneous)/(target maximum–target spontaneous)] x 100%.
Adoptive transfer of DC
Draining lymph nodes were harvested on Day 8 (see Fig. 5A
) and were digested with DNase I and collagenase solution. Mononuclear cells were obtained by centrifugation over a Histopaque-1077 density gradient (Sigma Chemical Co.), and DCs were isolated by CD11c-conjugated magnetic microbeads (Miltenyi Biotec, Auburn, CA, USA). CD11c-positive DCs (2.5x105/mouse) were injected into the left flank of GCV-treated KO mice on Day 8 (see Fig. 5A
). On Day 18, DC-transferred mice were rechallenged with 1 x 105 BNL cells in their right flank, and tumor sizes were measured.
Statistical analysis
Data were analyzed statistically using one-way ANOVA followed by the Tukey-Kramer test, except for tumor progression data, which were analyzed using two-way ANOVA. Data of tumor sizes after adoptive transfer of the DC experiment were analyzed using the Mann-Whitney U test. P < 0.05 was considered statistically significant.
RESULTS
GCV treatment induces tumor cell apoptosis with intratumoral CCR1-, CCR5-, and CCL3-positive cell accumulation in WT mice
We investigated whether HSV-tk-GCV treatment can induce apoptosis in vitro in the tk-transfected murine hepatoma cell line BNL-tk. GCV treatment significantly increased the proportions of early (annexin-positive but PI-negative) and late (annexin-positive and PI-positive) apoptotic cells (Fig. 1 A and B)
. We injected GCV into WT mice i.p. after the s.c. BNL-tk tumor was formed macroscopically, according to the schedule, as shown in Figure 1C
. Microscopic analysis revealed that more than half of the tumor cells were apoptotic and that a large number of mononuclear cells had accumulated in the tumor sites on Day 19 immediately following the completion of treatment (Fig. 1D
and Supplemental Fig. 1). Thereafter, the tumor regressed macroscopically. We next investigated the chemokine receptor expression by tumor-infiltrating cells after the induction of in vivo tumor apoptosis by suicide gene therapy. Immunohistochemical analysis revealed the presence of few CCR1-, CCR5-, or CCL3-positive cells in tumors without GCV treatment (Fig. 2A
). In contrast, GCV treatment caused intratumoral infiltration of a large number of CCR1-, CCR5-, and CCL3-positive cells in WT mice, along with massive apoptosis of tumor cells (Fig. 2A
and Supplemental Fig. 2). The intratumoral mRNA expression of CCL3, CCL4, and CCL5 was markedly increased 3 days after GCV treatment, whereas that of their receptors CCR1 and CCR5 was augmented later than 3 days after GCV injection (Fig. 2B)
.
Tumor-infiltrating DCs express CCR1 and CCR5
To determine the type of tumor-infiltrating cells expressing CCR1, CCR5, or CCL3, we performed a double-color immunofluorescence analysis. CD4- and CD8-positive T cells expressed CCR5 but not CCR1 (Fig. 3A
and Supplemental Fig. 3A; 78.2±8.9% of CD4-positive T cells and 92.6±8.2% of CD8-positive T cells expressed CCR5, and CCR1 was not detected in CD4- or CD8-positive T cells). In contrast, CD11c- and DEC205-positive cells, which infiltrated to tumor sites of WT mice after GCV treatment, expressed CCR1 and CCR5 (Fig. 3B
and Supplemental Fig. 3B; 100% of CD11c-positive DCs expressed CCR1 and CCR5, 97.5±5.6% DEC205-positive cells expressed CCR1, and 100% DEC-positive cells expressed CCR5). Moreover, tumor-infiltrating, CD11c-positive DCs exhibited a "myeloid" phenotype, as 87.8 ± 14.8% of CD11c-positive cells expressed CD11b, and none of them expressed CD8a (Supplemental Fig. 3C). Furthermore, CCL3 proteins were detected in CD3-positive T cells, F4/80-positive macrophages (Fig. 3C
; 72.0±9.1% of CD3-positive T cells and 87.0±12.0% of F4/80-positive macrophages expressed CCL3). These observations suggest that apoptosis induced by GCV treatment enhanced the expression of CCL3, CCL4, and CCL5 and then produced chemokines attracted to CD11c-positive DCs as well as CD3-positive T cells. To address this possibility, we investigated intratumoral infiltration of CD11c-positive DCs and CD3-positive T cells in CCR1KO, CCR5KO, or CCL3KO mice, which were s.c.-inoculated with 2 x 105 BNL-tk cells into WT and KO mice. The lack of CCR1, CCR5, or CCL3 had no discernible effects on the growth of primary tumors (Fig. 4A
). Then, we injected GCV i.p. into the mice as shown in Figure 1C
. Immunohistochemical analysis revealed the presence of few CD3-, F4/80-, or DEC205-positive cells in tumors without GCV treatment (Fig. 4B)
. GCV treatment induced tumor cell apoptosis in CCL3KO, CCR1KO, and CCR5KO mice to a similar extent as that in WT mice (data not shown). Moreover, GCV treatment caused intratumoral accumulation of a large number of CD3-, CD4-, and CD8-positive T cells and DEC205- and CD11c-positive DCs in WT mice (Fig. 4 B and C)
. By contrast, the increases in intratumorally accumulating DEC205- and CD11c-positive cells and to a lesser extent, CD3-, CD4-, and CD8-positive cells were attenuated in CCR1KO, CCR5KO, and CCL3KO mice (Fig. 4 B and C)
. In contrast, GCV treatment induced intratumoral infiltration of F4/80-positive macrophages (Fig. 4B)
and CD49b/DX5-positive NK cells (data not shown) in WT and KO mice to a similar extent.
Partial failure of CCR1KO, CCR5KO, and CCL3KO mice in rejecting the rechallenged tumor
Apoptosis induced by GCV treatment caused intratumoral infiltration of DCs and T cells in a CCR1- and/or CCR5-dependent manner. As intratumoral infiltration of DCs and T cells is a prerequisite for the establishment of specific tumor immunity, we examined the immune status of GCV-treated mice by rechallenging the parental BNL cell line. To completely eradicate the primary BNL-tk tumor, GCV was administered between 2 and 5 days after the tumor injection (Fig. 5A
). Primary BNL-tk tumors were eradicated completely in WT, CCR1KO, CCR5KO, and CCL3KO mice at similar rates (data not shown). When these mice were injected again with parental BNL cells, WT mice rejected them completely. In contrast, CCR1KO, CCR5KO, and CCL3KO mice failed to completely eliminate the rechallenged tumor cells, although the growth rates were retarded in these mice compared with naïve WT mice (Fig. 5B)
. A marked cytotoxicity against BNL but not in CT26 cells was observed when draining lymph node-derived mononuclear cells of GCV-treated WT mice were used as effector cells. Only a modest amount of cytotoxicity was detected when mononuclear cells in the draining lymph nodes of GCV-treated CCR1KO, CCR5KO, or CCL3KO mice were used as effector cells (Fig. 5C)
. Further, GCV-induced tumor apoptosis enhanced the mRNA expression of Th1 cytokines such as IFN-, IL-12p40, and IL-18 in the draining lymph nodes of WT mice but not of CCR1KO, CCR5KO, and CCL3KO mice (Supplemental Fig. 4). Likewise, CD8+IFN-+ cells were increased markedly in GCV-treated WT mice when lymph node-derived mononuclear cells were cocultured with BNL cell lysates, compared with tumor-bearing or tumor-free WT mice (Fig. 5D)
. Increases in CD8+IFN-+ cells were less evident in CCR1KO, CCR5KO, or CCL3KO mice treated with tumor cells and GCV compared with WT mice when lymph node-derived cells were cocultured with BNL cell lysates (Fig. 5D)
. These observations suggest that the absence of CCR1, CCR5, or CCL3 greatly impaired the apoptosis-induced establishment of specific tumor immunity.
Apoptosis-induced migration of DCs to draining lymph nodes and intranodal T cell proliferation activation in a CCR1-, CCR5-, and/or CCL3-dependent manner
Tumor-infiltrating DCs can uptake tumor antigens at the tumor sites and migrate to the draining lymph nodes, where they mature to present antigens to T cells [17
, 18
]. Thus, we further explored the status of DCs as well as T cells in the draining lymph nodes. Following GCV treatment, tumor apoptosis increased the proportions of CD86+CD11c+ cells in the draining lymph nodes but not in distant lymph nodes in WT mice (Fig. 6A
). In contrast, GCV-induced increases in CD11c+ cell proportion were depressed in CCR1KO, CCR5KO, or CCL3KO mice (Fig. 6A)
. The levels of CD86 on CD11c+ cells were increased in GCV-treated WT mice compared with WT mice, which had been injected with neither BNL-tk cells nor GCV, although the levels of CD86 were depressed in the KO mice (mean fluorescent intensities of CD86 on CD11c+ cells: WT/BNL-tk/GCV, 114.3±8.6; WT/BNL-tk, 86.2±12.2; WT/no tumor, 86.5±2.6; CCR1KO/BNL-tk/GCV, 85.4±15.6; CCR5KO/BNL-tk/GCV, 92.3±12.6; CCL3KO/BNL-tk/GCV, 79.0±9.8). Moreover, GCV-induced tumor apoptosis significantly increased the numbers of total cells, CD4+ and CD8+ cells, in the draining lymph nodes of WT mice. GCV-induced increases in these cell populations were also attenuated in CCR1KO, CCR5KO, or CCL3KO mice (Fig. 6B)
. Lymphocytes expressing the cell proliferation marker Ki67 were increased in the paracortical areas of the draining lymph nodes of GCV-treated WT mice compared with the other groups (Fig. 6C
and Supplemental Fig. 5). Injection of BNL-tk cells marginally increased the proportion of activated CD4+ T cells, defined as CD44hiCD62LloCD4+, in the draining lymph nodes. Coinjection of GCV further augmented this increment in WT mice but not in KO mice (Fig. 6 D and E)
. These observations suggest that the absence of CCR1, CCR5, or CCL3 impaired the GCV-induced migration of DCs into the draining lymph nodes and the subsequent proliferation and activation of T cells in the draining lymph nodes.
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Figure 6. Apoptosis-induced migration of DCs to the draining lymph nodes (DLN) and intranodal T cell proliferation and activation in a CCR1-, CCR5-, and/or CCL3-dependent manner. Mice were treated according to the schedule shown in Figure 5A
. (A) Draining lymph nodes or distal lymph nodes were harvested on Day 8 as indicated in Figure 5A
. Mononuclear cells were stained with a combination of FITC-labeled anti-CD86 and PE-labeled anti-CD11c antibodies. The percentage of CD11c+CD86+ cells was determined and is indicated in the upper-right corners. Representative results from three individual animals are shown here. (B) Absolute cell numbers of each cell population in the draining lymph nodes or distal lymph nodes on Day 8 were determined as described in Materials and Methods. Error bars, ±1 SD; *, P < 0.05, compared with draining lymph nodes derived from WT mice treated with BNL-tk/GCV. (C) Draining lymph nodes harvested on Day 8 were immunostained with anti-Ki67 antibody, and percentages of Ki67+ cells in lymph nodes were determined. Error bars, ±1 SD; *, P < 0.05, compared with draining lymph nodes derived from WT mice treated with BNL-tk/GCV. (D and E) Mononuclear cells harvested on Day 8 were stained with a combination of FITC-labeled anti-CD4 and PE-labeled anti-CD44 (D) or PE-labeled anti-CD62L (E) antibodies. Histograms were gated on CD4-positive cells, and percentages of CD44hi (D) or CD62Llo (E) cells were determined. Representative results from three individual animals are shown here.
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Figure 7. Adoptive transfer of DCs harvested from GCV-treated WT mice restored the antitumor response of KO mice. After inoculation of BNL-tk tumors, CCR1KO, CCR5KO, or CCL3KO mice were treated with GCV from Days 2 to 5 as described in Figure 5A
. The DCs were then harvested from the draining lymph nodes of GCV-treated WT mice and transferred s.c. into the KO mice on Day 8 as shown in Figure 5A
. On Day 18, these mice were injected again with parental BNL cells, and tumor sizes were measured on Day 28. GCV-treated KO mice, transferred without or with DCs of naïve WT mice, were also rechallenged with parental BNL cells as control. Bars, mean; *, P < 0.05.
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Apoptosis was previously presumed to be immunologically silent or even tolerogenic [27 ]. However, recent reports have indicated that tumor cell apoptosis can induce antitumor immune responses effectively, as the immunogenicity of apoptotic tumor cells is dependent on apoptosis inducers. Indeed, gemcitabine-induced apoptosis can augment cross-priming of tumor-specific CD8+ T cells in vivo rather than cross-tolerizing [8 ]. Similarly, apoptosis induced by local radiation therapy can generate tumor antigen-specific effector cells that migrate to the tumor [28 ]. Moreover, apoptotic change caused by anthracyclin can induce the translocation of calreticulin to the apoptotic tumor cell surface, and calreticulin exposure can enhance the immunogenicity of apoptotic cancer cells [6 , 7 ]. Thus, apoptosis induced by these measures is sufficiently immunogenic to prevent tumor progression.
The combination of HSV-tk gene transfer and GCV can efficiently induce the apoptosis of the transfected tumors as observed in the present study. Here, we also observed that the treatment augmented the immune response, as evidenced by an increase in the number of tumor-infiltrating DCs. Subsequently, the number of DCs in the draining lymph nodes increased together with enhanced, specific immunity to the injected tumor. These observations suggest that tumor apoptosis induced by HSV-tk/GCV treatment is effective in generating specific tumor immunity, similar to that observed in the case of anticancer drug treatment.
Consistent with our present observations, CCL3 and its related chemokine CCL5 were detected in macrophages infiltrating human cancer tissues [29 , 30 ]. Given their potent chemotactic activity against various types of immune cells [31 ], gene transfer of CCL3 or CCL5 induced the accumulation of immune cells including DCs, T cells, macrophages, and NK cells in the tumor sites, resulting in delayed tumor growth and prolonged survival [16 , 32 33 34 ]. Moreover, combination therapy of the HSV-tk and CCL3/CCL20 gene induces an exaggerated accumulation of DCs, CD4+ cells, CD8+ cells, NK cells, and macrophages in the tumor sites compared with HSV-tk/GCV treatment alone, and the net effects are tumor regression and prolonged survival [34 ]. However, the roles of endogenously produced CCL3 and its related chemokines in tumor apoptosis still remain to be elucidated.
In human HCC, tumor-infiltrating lymphocytes express high levels of CCR5 and CXCR3. Moreover, these lymphocytes show strong chemotactic responses to CC and CXC chemokines including CCL3, CCL4, and CXCL9 [35 ]. Additional treatments are nevertheless required to enhance the immune responses, as the CCR5- or CXCR3-positive lymphocytes are insufficient to evoke immune responses and eradicate tumor tissues. We demonstrated that suicide gene therapy-induced tumor cell apoptosis augments CCR1- and CCR5-positive cell infiltration into the hepatoma tissues. Further, these CCR1- or CCR5-positive cells are DCs and/or T cells—the cells indispensable for tumor immunity. Thus, suicide gene therapy can potentially enhance tumor immunity by attracting these immune cells to the apoptotic tumor cells.
The initial step leading to specific tumor immunity is the capture of tumor antigens by macrophages and immature DCs, both of which accumulate in tumor sites. However, in this model, CCR1KO, CCR5KO, and CCL3KO mice failed to completely eliminate the rechallenged tumor cells along with reduced intratumoral accumulation of DCs but not F4/80-positive macrophages. These observations suggest that the establishment of specific tumor immunity requires intratumoral recruitment of immature DCs but not macrophages. Activated NK cells can also induce DC maturation in lymphoid organs as well as in nonlymphoid tissues. Although NK cells express chemokine receptors such as CCR5 [36 ], the deficiency of CCR1 or CCR5 has little effects on intratumoral infiltration of NK cells. These observations preclude the crucial role of NK cells in the establishment of specific tumor immunity in this model.
Immature DCs use several chemokine receptors including CCR1, CCR2, CCR4, CCR5, CCR6, CCR8, and CXCR4 for their migration [22 ]. However, the chemokine receptor(s) regulating immature DC trafficking to tumor sites still need(s) to be determined. We previously observed that CCL3 induced mobilization of DC precursors into circulation [37 ] and detected CCL3 in tumor-infiltrating CD3+ T cells and macrophages after GCV treatment. Therefore, we investigated the roles of CCL3 and its receptors CCR1 and CCR5 in the intratumoral recruitment of DCs and the subsequent establishment of specific tumor immunity. Although CCL4 and CCL5 expression was augmented along with CCL3 expression in tumor sites, the deletion of the CCL3 gene alone markedly reduced the DC migration, intranodal T cell accumulation, and subsequent Th1 cytokine expression. Similarly, deletion of the CCL3 gene alone prevented coxsackievirus-induced myocarditis [38 ], despite enhanced, intracardiac expression of CCL3, CCL4, and CCL5 mRNA [39 ]. Thus, these three chemokines may form a positive feedback loop, and the deletion of either chemokine might reduce the expression of the others. Moreover, the lack of CCR1 or CCR5 reduces the migration of DCs to tumor sites and subsequent tumor immunity in the draining lymph nodes, such as DC and T cell accumulation and Th1 cytokine expression. As almost all CD11c- and DEC205-positive DCs express CCR1 and CCR5, DC migration may require coordinated and synergistic actions of both of these chemokine receptors.
We have provided definitive evidence regarding the essential contribution of CCL3 and its receptors to apoptosis-induced, specific tumor immunity, which exert their role by attracting DCs to tumor tissues. These observations further suggest that specific tumor immunity can be established more efficiently if some techniques such as chemokine gene transfer can augment the recruitment of immature DCs to apoptotic tumor tissues caused by chemotherapeutic agents and/or irradiation as well as suicide gene therapy.
ACKNOWLEDGEMENTS
We thank Dr. Philip M. Murphy (NIAID, NIH) for providing us with CCR1KO mice. We also thank Dr. Toshikazu Kondo (Wakayama Medical University, Wakayama, Japan) for his technical advice about double-color immunofluorescence analysis.
Received November 26, 2007; revised June 12, 2008; accepted June 30, 2008.
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