Cell Biology International ISSN 1065-6995 doi: 10.1002/cbin.10485
RESEARCH ARTICLE
Toll-like receptor 4 signaling inhibits malignant pleural effusion by altering Th1/Th17 responses Qian-Qian Xu1,2, Qiong Zhou1, Li-Li Xu2, Hua Lin1, Xiao-Juan Wang2, Wan-Li Ma1, Kan Zhai3, Zhao-Hui Tong1,3, Yunchao Su4 and Huan-Zhong Shi1,2,3* 1 Department of Respiratory and Critical Care Medicine, Union Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, China 2 Department of Respiratory and Critical Care Medicine, Beijing Chaoyang Hospital, Capital Medical University, Beijing, China 3 Center of Medical Research, Beijing Chaoyang Hospital, Capital Medical University, Beijing, China 4 Department of Pharmacology and Toxicology, Medical College of Georgia, Georgia Regents University, Augusta, GA, USA
Abstract Toll-like receptor 4 (TLR4) is involved in multiple malignancies; however, the role of TLR4 in the pathogenesis of malignant pleural effusion (MPE) remains unknown. The objectives of this study were to explore the impact of TLR4 signaling on the development of MPE in a murine model and to define the underline mechanisms by which TLR works. Development of MPE as well as proliferation and angiogenesis of pleural tumor were determined in TLR4–/– and wild type mice. Differentiation of Th1 and Th17 cells as well as their signal transductions was explored. The effects of TLR4 signaling on survival of mice bearing MPE were also investigated. Compared with wild type mice, Th1 cells were augmented, and Th17 cells were suppressed in MPE from TLR4–/– mice. The in vitro experiments showed that TLR4 deficiency promoted Th1 cell differentiation via enhancing STAT1 pathway and inhibited Th17 cell differentiation via suppressing STAT3 pathway. TLR4 deficiency promoted MPE formation and, thus, accelerated the death of mice bearing MPE, whereas intraperitoneal injection of antiIFN-g mAb or recombinant mouse IL-17 protein into TLR4–/– mice was associated with improved survival. Our data provides the first definitive evidence of a role for TLR4 signaling in protective immunity in the development of MPE. Our findings also demonstrate that TLR4 deficiency promotes MPE formation and accelerates mouse death by enhancing Th1 and suppressing Th17 response. Keywords: malignant pleural effusion; Th1 cells; Th17 cells; toll-like receptor 4
AT A GLANCE COMMENTARY
Scientific knowledge on the subject Toll-like receptor 4 (TLR4) has been well known to be one of the key immune system effectors in the host defense against microbial infection. Little has been known about the function of TLR4 in tumor immunity.
What this study adds to the field TLR4 signaling inhibited Th1 cell differentiation via suppressing STAT1 pathway and promoted Th17 cell differentiation via enhancing STAT3 pathway in a mouse model of malignant pleural effusion. TLR4 deficiency
accelerated the death of mice bearing MPE, whereas intraperitoneal injection of anti-IFN-g mAb or recombinant mouse IL-17 protein into TLR4 deficient mice was associated with improved survival.
Introduction Toll-like receptors (TLRs) are primarily involved in innate immune response to microbial pathogens through recognition of conserved pathogen-associated molecular patterns (Akira et al., 2006; Kawai T and Akira S, 2006). Pinto A et al. (2011) provided evidence that the discovery of TLRs as essential components of the immune system has introduced new subjects to the field of research on the pathogenesis of
Corresponding author: e-mail: [email protected] Qian-Qian Xu and Qiong Zhou contributed equally to the present work. Conflict of interest: The authors have declared that no conflict of interest exists.
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malignancies. Huang B et al. (2005) and Gonzalez-Reyes S et al. (2010) found that TLRs have been detected in multiple tumor cell lines and tumor tissues and have exerted a great influence on cancer immune surveillance. Furthermore, activated TLR4 signaling pathway in tumor cells may affect cancer progression, anti-apoptotic activity, and resistance to host immune responses (Andreani et al., 2007; He et al., 2007; Ikebe et al., 2009; O’Neill LA, 2008). Malignant pleural effusion (MPE), the accumulation of pleural fluid caused by cancer metastasis to the pleural space, is a frequent and clinically significant systemic manifestation of various cancers that adversely affects patient survival and quality of life (Wu et al., 2013; Ryu et al., 2014). The pathogenesis of MPE is not yet known completely. In addition to impairment of pleural fluid drainage induced by tumor metastasis, an inflammatory signaling network between tumor cells, and host vascular and immune systems has been documented to contribute to MPE development (Stathopoulos and Kalomenidis, 2012). Our previous data have revealed that several subpopulations of CD4þ T cells, including regulatory T cells (Chen et al., 2005), IL-17– producing CD4þ T cells (Th17 cells) (Ye et al., 2010), IL-22– producing CD4þ T cells (Th22 cells) (Ye et al., 2012a), and IL-9–producing CD4þ T cells (Th9 cells) (Ye et al., 2012b), play important roles as immune regulator in the pathogenesis of human MPE. It has been established that Th17 cells can exert both proand anti-tumor effects, depending on the context within the tumor microenvironment (Zou and Restifo, 2010; Ye et al., 2013). In addition, IFN-g–producing CD4þ T cells (Th1 cells) have been reported to possess some anti-tumor activity in some earlier experiments (Muranski and Restifo, 2009). Our previous data have shown that the numbers of both Th17 and Th1 cells were increased in human MPE compared with peripheral blood and that Th17 cell numbers were positively correlated with Th1 cell numbers, suggesting a collaboration between Th17 and Th1 cells in MPE (Ye et al., 2010). More recently, we have reported that the absence of Th17 cells promotes MPE formation, whereas the absence of Th1 cells inhibits MPE formation (Li et al., 2014). In the present study, we investigated the effect of TLR4 signaling on the development of MPE and determined whether TLR4 affected MPE development by regulating Th1/Th17 responses. Materials and methods
TLR4 in pleural malignancy
(20–25 g)-, and age (8–12 wk)-matched. The mice were maintained on a chow diet in a 12-h light/12-h dark environment at 25 C in Animal Care Facility of Beijing Chaoyang Hospital or in Tongji Medical College Animal Care Facility, according to institutional guidelines. All animal studies were approved by Institutional Animal Care and Utilization Committee of Beijing Chaoyang Hospital, Capital Medical University, and Tongji Medical College, Huazhong University of Science, and Technology.
MPE Model A murine model of MPE by intrapleural injecting Lewis lung cancer (LLC) cells was made as described previously (Lin et al., 2014). Mice were anesthetized by isoflurane inhalation, the skin overlying the anterior and lateral chest wall was shaved and disinfected, and a 5-mm-long trans-verse skin incision was made on the right anterolateral thoracic area at the xiphoid level. Fascia and muscle were retracted, and 1.5 105 LLC cells purchased from the American Type Culture Collection ([Manassas, VA]) suspended in 50 mL phosphate buffered saline (PBS) were injected into pleural cavity through an intercostal space under direct observation. The skin incision was closed using continuous 5–0 Ethilon monofilament suture, and the animals were monitored until complete recovery.
Sample collection and processing Fourteen days after pleural injection of LLC cells, mice were euthanized by CO2 asphyxiation, and blood was drawn from the retro-orbital veins. Thereafter, the abdominal wall was opened, and the viscera were retracted to visualize the diaphragm. Pleural fluid was gently aspirated using a 3-mL syringe, and its volume was measured with a 1,000-mL pipette. The samples were made a smear slide for WrightGiemsa staining and were determined for leukocyte differential counts under light microscope. MPE and blood samples from each mouse were collected in heparin-treated tubes and were immersed in ice immediately and were then centrifuged at 300 g for 10 min. In order to obtain spleen cell suspension, spleens were removed and cut into small pieces, and then minced in PBS and filtered. The cell pellets of MPE, blood, and spleen were resuspended in PBS, and mononuclear cells were isolated by Ficoll-Hypaque gradient centrifugation (Pharmacia, Uppsala, Sweden) to determine T cell subsets within 1 h.
Animals Wild type (WT) C57BL/10 mice and TLR4–/– mice in a C57BL/10 background were purchased from the Model Animal Research Center of Nanjing University (Nanjing, China). Mice used for experiments were sex-, weight
Positron emission tomography (PET) and computed tomography (CT) scanning 1
All PET images were acquired using Trans-PET BioCaliburnTM LH (Raycan Technology Co., Ltd, Suzhou,
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China) consisting of 12 basic detector modules. Fifteenmin PET scans were begun between 60 and 70 min after retro-orbital injection of 200–300 mCi of 18-fluoro-6deoxy-glucose. In consideration of the PET system’s spatial resolution, reconstructed PET images had pixel dimensions of (0.5 mm)2 and 0.5 mm slice thickness. Furthermore, due to the OSEM reconstruction method using point spread function system response matrix, the size of the reconstructed 3D image was set to be 280 280 104 (XxYxZ) that must be smaller than or equal to the size of point spread function system response matrix. Also, CT images were acquired on a small animal CT scanner provided by the Trans-PET system with characteristics of an axial field of view of 5.0 cm, a radial field of view of 7.0 cm, and a spatial resolution of 0.5 mm. The PET and CT images were acquired with the mouse under anesthesia in a common animal holder. The coregistration of the images was implemented using the AMIDE software package (The Free Software Foundation Inc., Boston, MA).
Flow cytometry The Abs included anti–CD3, –CD4, –CD8, –IFN-g, –IL-17, –T-bet, –RORgt, –CD14, –Ly-6G, and –TLR4 mAbs, which were purchased from BD Pharmingen (San Diego, CA) or eBioscience (San Diego, CA). To determine intracellular cytokines, cells were stimulated for 4 h at 37 C with PMA (50 ng/mL, Sigma–Aldrich, St. Louis, MO) and ionomycin (1 mg/mL, Sigma–Aldrich) in the presence of Brefeldin A (10 mg/mL, Enzo Life Science). Cells were surface-stained with anti–CD3, –CD4, or –CD8 mAbs in PBS þ 2% heat-inactivated fetal bovine serum (FBS, Gibco) for 20 min at 4 C. Cells were resuspended in a fixation/permeabilization solution (Cytofix/Cytoperm; BD Pharmingen) and were incubated with anti–IFN-g or –IL-17 mAbs for 30 min at 4 C. Cells were then washed with permeabilization buffer and resuspended in PBS þ 2% FBS for flow cytometric analysis. To determine transcription factors, cells were surfacestained with anti–CD3, –CD4, or –CD8 mAbs in PBS þ 2% heat-inactivated fetal bovine serum for 20 min at 4 C. Cells were resuspended in a fixation/permeabilization solution and were incubated with anti–T-bet or –RORgt mAbs for 30 min at 4 C. Cells were then washed with permeabilization buffer and resuspended in PBS þ 2% FBS for flow cytometric analysis. To determine TLR4 expression on the surface of CD14þ monocytes/macrophages, CD4þ T cells, CD8þ T cells, and Ly-6Gþ neutrophils, cells isolated from MPE of WT mice were stained by the similar protocols except without the stimulating procedure. 1122
Flow cytometry was performed on a FACS Canto II (BD Biosciences) and the data were analyzed using BD FCSDiva Software and FCS Express 4 software (De Novo Software, Los Angeles, CA).
Quantitative real-time PCR Total RNA was isolated from cell pellets with Trizol reagent (Invitrogen, Carlsbad, CA). cDNAs were synthesized with oligo (dT) primers using the First Strand cDNA Synthesis Kit ReverTra Ace-a-(TOYOBO, Osaka, Japan). cDNAs were used as templates in real-time PCR with the SYBR Green I Real-time PCR Master Mix (TOYOBO, Osaka, Japan). DNA was amplified under the following typical cycling conditions: denaturation at 95 C for 15 s, annealing at 58 C for 20 s, extension at 72 C for 20 s. The samples were amplified for 40 cycles. The following primers were used: for Ifng, forward primer: 50 -CATCAGCAACAACATAAGCG-30 , reverse primer: 50 -GACCTCAAACTTGGCAATACTC-30 ; for t-bet, forward primer: 50 -CAACAACCCCTTTGCGAAAG-30 , reverse primer: 50 -TCCCCCAAGCAGTTGACAGT-30 ; for Il17a, forward primer: 50 -TGTGTCTCTGATGCTGTTGC-30 , reverse primer: 50 -GGAACGGTTGAGGTAGTCTG-30 ; for RORgt, forward primer: 50 TGCAAGACTCATCGACAAGG-30 reverse primer: 50 AGGGGATTCAACATCAGTGC-30 ; for b-actin, forward primer: 50 -CTGAGAGGGAAATCGTGCGT-30 , reverse primer: 50 -CCACAGGATTCCATACCCAAGA-30 . b-actin was used as an internal control, and levels of each gene were normalized to b-actin expression using the DDCt-method, with WT mouse experimental value set to 1.
In vitro differentiation of Th1 and Th17 cells Mononuclear cells from mouse spleens were isolated by Ficoll-Hypaque gradient centrifugation. Bulk CD4þ T cells were isolated by negative selection (by depletion of CD8aþ, CD45Rþ, CD11bþ, CD25þ, CD49bþ, TCRg/dþ, and Ter119þ cells) with CD4þ cell biotin-Ab cocktail II and antibiotin microbeads. After isolation of bulk CD4þ T cells, naive CD4þCD62Lþ T cells were isolated by CD4þCD62Lþ T cell isolation kit II (Miltenyi Biotec, Aubum, CA) according to the manufacturer. The purity of naive CD4þCD62Lþ T cells was more than 90%, as measured by flow cytometry. Purified naive CD4þ T cells were cultured in RPMI-1640 medium containing 10% FBS in the presence of platebound anti–CD3 (4 mg/mL) and anti–CD28 (2 mg/mL) mAbs. Cytokines used for cell differentiation were as follow: Th1 condition, IL-2 (50 U/mL), IL-12 (5 ng/mL) and anti–IL-4 mAb (10 mg/mL); Th17 condition, IL-2 (50 U/mL), TGF-b (2 ng/ml), IL-6 (100 ng/mL), IL-1b (10 ng/mL), IL-23 (10 ng/mL), anti–IL-4 mAb (10 mg/mL),
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and anti–IFN-g mAb (1 mg/mL). In some experiments, designated concentrations of lipopolysaccharide (LPS; 0, 0.5, and 1.0 mg/mL) were added into Th1 or Th17 conditions. Cells were cultured for 5 d and then stimulated for 4 h with PMA and ionomycin in the presence of Brefeldin A before being stained for cytokines described in the Flow Cytometry section.
Determination of signal transductions of IFN-g and IL-17 Purified na€ıve CD4þ T cells were cultured in RPMI-1640 medium containing 10% FBS in the presence of plate-bound anti–CD3 (4 mg/mL) and –CD28 (2 mg/mL) mAbs in Th1 or Th17 conditions as described above for 48 h. The cells were stimulated for 30 min with IL-2 (50 U/mL) and IL-12 (5 ng/mL) for phospho (p)-signal transducer and activator of transcription (STAT) 1 and with IL-6 (100 ng/mL) and IL1b (10 ng/mL) for p-STAT3. Then, the cells were harvested, fixed with 1.5% paraformaldehyde, and were stained for p-STAT1 or p-STAT3 and subjected to flow cytometry as described above.
TLR4 in pleural malignancy
RESULTS
TLR4 expression in MPE Our previous data have shown that a large proportion of nucleated cells in MPE of WT mice were mononuclear cells, lymphocytes, and neutrophils, and a very small proportion of cells were eosinophils (Li et al., 2014), we thus first explored the expression of TLR4 on these pleural cells using flow cytometry. As shown in Table 1, CD14þ monocytes/ macrophages, CD4þ T cells, CD8þ T cells, and Ly-6Gþ neutrophils in MPE of WT mice expressed much higher level of TLR4 than their counterparts in peripheral blood or spleen (all P < 0.05), and there were no differences in TLR4 expression between blood cells and spleen cells (all P < 0.05).
Effect of TLR4 deficiency on MPE formation
The observation of survival advantage included 1 group of WT mice and 5 groups of TLR4–/– mice (each n ¼ 15). At days 1, 3, 6, 9, and 12 after intrapleural instillation of LLC cells, 2 groups of TLR4–/– mice received intraperitoneal injection of 100 mg anti-IFN-g neutralized mAb (eBioscience, clone XMG1.2) (TLR4–/– þ a-IFN-g) and 100 mg mouse isotype control mAb (eBioscience), respectively; and another 2 groups of TLR4–/– mice received intraperitoneal injection 1 mg recombinant mouse IL17A protein (R&D System, Minneapolis, MN) diluted in PBS containing 0.1% albumin (TLR4–/– þ rmIL-17) and PBS containing 0.1% albumin in a total volume of 100 mL, respectively. The mice without heartbeat and respiration were determined to be dead. The administration regimens of anti-IFN-g mAb and recombinant mouse IL-17A were based upon our preliminary experiments.
Fourteen days after pleural injection of LLC cells, pleural tumors were found to distribute on the parietal and visceral pleura of WT and TLR4–/– mice. Bloody pleural fluid could be found through diaphragm, surrounding tumor foci in the two groups (Figure 1A). MPE volume in TLR4–/– mice (0.72 0.08 mL) was much larger than that in WT mice (0.34 0.07 mL; P < 0.001) (Table 2). The numbers of nucleated cells were significantly decreased in TLR4–/– mice compared with WT mice (P < 0.001). Interestingly, the absolute numbers of total nucleated cells and each cell types, including mononuclear cells, lymphocytes, neutrophils, and eosinophils, in the whole MPE, were not different between the two groups (all P > 0.05) (Table 2). FDG-PET and CT scans were performed to evaluate the pleural tumors and MPE in the living mice. As shown in Figure 1B, increased radiotracer uptake on FDG-PET scanning was found in the areas of tumors and MPE. Compared with WT mice, the PET imaging confirmed higher 18F-FDG retention in TLR4–/– mice. The maximum standardized uptake value (SUVmax) in TLR4–/– and WT mice were 8.8 1.9 and 3.7 0.7, respectively, with statistic significance between the two groups (P ¼ 0.021) (Figure 1C).
Statistics
Table 1 Percentages of toll-like receptor 4 positive cells in a murine MPE model
Survival advantage experiment
Data were normally distributed, and all values are expressed as mean SEM. Differences between two or multiple experimental groups were compared using Student’s t-test, one-way analysis of variance (ANOVA), or two-way ANOVA, as appropriate. Survival was estimated by the Kaplan-Meier method and compared by the log-rank test. Statistical analysis was performed with SPSS version 16.0 Software (Chicago, IL), and P < 0.05 was considered statistically significant.
n Monocytes/macrophages CD4þ T cells CD8þ T cells Neutrophils
MPE
Blood
Spleen
10 15.0 3.1* 7.5 0.7* 12.3 3.4* 41.7 4.1*
10 9.4 2.3 2.4 1.2 5.3 0.5 28.9 3.7
10 10.2 1.2 1.3 0.4 3.7 1.3 32.4 4.0
MPE ¼ malignant pleural effusion. *P < 0.05 compared with blood or spleen, and comparisons were determined by two-way analysis of variance (ANOVA).
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TLR4 deficiency increased Th1 cells and decreased Th17 cells in MPE Th1 cells and Th17 cells were identified by flow cytometry (Figure 2A). In both WT and TLR4–/– mice, the numbers of Th1 (Figure 2B), and Th17 cells (Figure 2C) were significantly higher in MPE (P < 0.001 for all). More importantly, we noted that Th1 cell numbers were increased and Th17 cell numbers were decreased in MPE from TLR4–/– mice as compared with WT mice (P < 0.01 for both). The numbers of Th1 and Th17 cells in blood and spleen showed no significant differences in either TLR4–/– or WT mice (P > 0.05 for all).
Expression of IFN-g, IL-17, T-bet, and RORgt mRNA in MPE We noted that there was much higher mRNA expression of IFN-g (Figure S1A) and T-bet (Figure S1B) in MPE from TLR4–/– mice than that from WT mice (both P < 0.05). In contrast, there was much lower mRNA expression of IL-17 (Figure S1C) and RORgt (Figure S1D) in MPE from TLR4–/– mice than that from WT mice (P < 0.01 for both).
Effect of TLR4 deficiency on differentiation of Th1 and Th17 cells in vitro Figure 1 Development of malignant pleural effusions (MPE) in wild type (WT) and TLR4–/– mice. (A) Fourteen days after intrapleural injection of Lewis lung cancer cells, multiple tumor foci could be seen on the parietal and visceral pleura, and MPE was visible in the pleural space. (B) Positron emission tomography and computed tomography imaging of developing pleural tumors and MPE in WT (top panels) and TLR4–/– mice (bottom panels). (C) The maximum standardized uptake value (SUVmax) on positron emission tomography were calculated (both n ¼ 10). Data are presented as means SEM, the comparisons were determined by Student’s t-test.
Na€ıve CD4þ T cells from spleens of WT or TLR4–/– mice were purified and cultured in vitro in the presence of IL-2, IL-12, and anti–IL-4 mAb (Th1 condition); or IL-2, TGF-b, IL-6, IL-1b, IL-23, anti–IL-4, and anti–IFN-g mAbs (Th17 condition). Compared with WT na€ıve CD4þ T cells, the differentiation of Th1 cells were significantly increased (Figures 3A and 3B), and the differentiation of Th17 cells were significantly decreased (Figures 3A and 3C) from TLR4–/– na€ıve CD4þ T cells. In WT mice, addition of LPS to Th1 condition inhibited Th1 cell differentiation in a dosedependent manner (Figure 3D), while addition of LPS to Th17 condition promoted Th17 cell differentiation in a dose-dependent manner as well (Figure 3E).
Table 2 Cytological characteristics in malignant pleural effusion
n Pleural fluid volume, mL Nucleated cells, 106/mL Cell differential counts, % Mononuclear cells Lymphocytes Neutrophils Eosinophils
Wild type mice
TLR4–/– mice
P-values*
15 0.34 0.07 42.8 3.4
15 0.72 0.08 21.0 4.3
RESEARCH ARTICLE
Toll-like receptor 4 signaling inhibits malignant pleural effusion by altering Th1/Th17 responses Qian-Qian Xu1,2, Qiong Zhou1, Li-Li Xu2, Hua Lin1, Xiao-Juan Wang2, Wan-Li Ma1, Kan Zhai3, Zhao-Hui Tong1,3, Yunchao Su4 and Huan-Zhong Shi1,2,3* 1 Department of Respiratory and Critical Care Medicine, Union Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, China 2 Department of Respiratory and Critical Care Medicine, Beijing Chaoyang Hospital, Capital Medical University, Beijing, China 3 Center of Medical Research, Beijing Chaoyang Hospital, Capital Medical University, Beijing, China 4 Department of Pharmacology and Toxicology, Medical College of Georgia, Georgia Regents University, Augusta, GA, USA
Abstract Toll-like receptor 4 (TLR4) is involved in multiple malignancies; however, the role of TLR4 in the pathogenesis of malignant pleural effusion (MPE) remains unknown. The objectives of this study were to explore the impact of TLR4 signaling on the development of MPE in a murine model and to define the underline mechanisms by which TLR works. Development of MPE as well as proliferation and angiogenesis of pleural tumor were determined in TLR4–/– and wild type mice. Differentiation of Th1 and Th17 cells as well as their signal transductions was explored. The effects of TLR4 signaling on survival of mice bearing MPE were also investigated. Compared with wild type mice, Th1 cells were augmented, and Th17 cells were suppressed in MPE from TLR4–/– mice. The in vitro experiments showed that TLR4 deficiency promoted Th1 cell differentiation via enhancing STAT1 pathway and inhibited Th17 cell differentiation via suppressing STAT3 pathway. TLR4 deficiency promoted MPE formation and, thus, accelerated the death of mice bearing MPE, whereas intraperitoneal injection of antiIFN-g mAb or recombinant mouse IL-17 protein into TLR4–/– mice was associated with improved survival. Our data provides the first definitive evidence of a role for TLR4 signaling in protective immunity in the development of MPE. Our findings also demonstrate that TLR4 deficiency promotes MPE formation and accelerates mouse death by enhancing Th1 and suppressing Th17 response. Keywords: malignant pleural effusion; Th1 cells; Th17 cells; toll-like receptor 4
AT A GLANCE COMMENTARY
Scientific knowledge on the subject Toll-like receptor 4 (TLR4) has been well known to be one of the key immune system effectors in the host defense against microbial infection. Little has been known about the function of TLR4 in tumor immunity.
What this study adds to the field TLR4 signaling inhibited Th1 cell differentiation via suppressing STAT1 pathway and promoted Th17 cell differentiation via enhancing STAT3 pathway in a mouse model of malignant pleural effusion. TLR4 deficiency
accelerated the death of mice bearing MPE, whereas intraperitoneal injection of anti-IFN-g mAb or recombinant mouse IL-17 protein into TLR4 deficient mice was associated with improved survival.
Introduction Toll-like receptors (TLRs) are primarily involved in innate immune response to microbial pathogens through recognition of conserved pathogen-associated molecular patterns (Akira et al., 2006; Kawai T and Akira S, 2006). Pinto A et al. (2011) provided evidence that the discovery of TLRs as essential components of the immune system has introduced new subjects to the field of research on the pathogenesis of
Corresponding author: e-mail: [email protected] Qian-Qian Xu and Qiong Zhou contributed equally to the present work. Conflict of interest: The authors have declared that no conflict of interest exists.
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malignancies. Huang B et al. (2005) and Gonzalez-Reyes S et al. (2010) found that TLRs have been detected in multiple tumor cell lines and tumor tissues and have exerted a great influence on cancer immune surveillance. Furthermore, activated TLR4 signaling pathway in tumor cells may affect cancer progression, anti-apoptotic activity, and resistance to host immune responses (Andreani et al., 2007; He et al., 2007; Ikebe et al., 2009; O’Neill LA, 2008). Malignant pleural effusion (MPE), the accumulation of pleural fluid caused by cancer metastasis to the pleural space, is a frequent and clinically significant systemic manifestation of various cancers that adversely affects patient survival and quality of life (Wu et al., 2013; Ryu et al., 2014). The pathogenesis of MPE is not yet known completely. In addition to impairment of pleural fluid drainage induced by tumor metastasis, an inflammatory signaling network between tumor cells, and host vascular and immune systems has been documented to contribute to MPE development (Stathopoulos and Kalomenidis, 2012). Our previous data have revealed that several subpopulations of CD4þ T cells, including regulatory T cells (Chen et al., 2005), IL-17– producing CD4þ T cells (Th17 cells) (Ye et al., 2010), IL-22– producing CD4þ T cells (Th22 cells) (Ye et al., 2012a), and IL-9–producing CD4þ T cells (Th9 cells) (Ye et al., 2012b), play important roles as immune regulator in the pathogenesis of human MPE. It has been established that Th17 cells can exert both proand anti-tumor effects, depending on the context within the tumor microenvironment (Zou and Restifo, 2010; Ye et al., 2013). In addition, IFN-g–producing CD4þ T cells (Th1 cells) have been reported to possess some anti-tumor activity in some earlier experiments (Muranski and Restifo, 2009). Our previous data have shown that the numbers of both Th17 and Th1 cells were increased in human MPE compared with peripheral blood and that Th17 cell numbers were positively correlated with Th1 cell numbers, suggesting a collaboration between Th17 and Th1 cells in MPE (Ye et al., 2010). More recently, we have reported that the absence of Th17 cells promotes MPE formation, whereas the absence of Th1 cells inhibits MPE formation (Li et al., 2014). In the present study, we investigated the effect of TLR4 signaling on the development of MPE and determined whether TLR4 affected MPE development by regulating Th1/Th17 responses. Materials and methods
TLR4 in pleural malignancy
(20–25 g)-, and age (8–12 wk)-matched. The mice were maintained on a chow diet in a 12-h light/12-h dark environment at 25 C in Animal Care Facility of Beijing Chaoyang Hospital or in Tongji Medical College Animal Care Facility, according to institutional guidelines. All animal studies were approved by Institutional Animal Care and Utilization Committee of Beijing Chaoyang Hospital, Capital Medical University, and Tongji Medical College, Huazhong University of Science, and Technology.
MPE Model A murine model of MPE by intrapleural injecting Lewis lung cancer (LLC) cells was made as described previously (Lin et al., 2014). Mice were anesthetized by isoflurane inhalation, the skin overlying the anterior and lateral chest wall was shaved and disinfected, and a 5-mm-long trans-verse skin incision was made on the right anterolateral thoracic area at the xiphoid level. Fascia and muscle were retracted, and 1.5 105 LLC cells purchased from the American Type Culture Collection ([Manassas, VA]) suspended in 50 mL phosphate buffered saline (PBS) were injected into pleural cavity through an intercostal space under direct observation. The skin incision was closed using continuous 5–0 Ethilon monofilament suture, and the animals were monitored until complete recovery.
Sample collection and processing Fourteen days after pleural injection of LLC cells, mice were euthanized by CO2 asphyxiation, and blood was drawn from the retro-orbital veins. Thereafter, the abdominal wall was opened, and the viscera were retracted to visualize the diaphragm. Pleural fluid was gently aspirated using a 3-mL syringe, and its volume was measured with a 1,000-mL pipette. The samples were made a smear slide for WrightGiemsa staining and were determined for leukocyte differential counts under light microscope. MPE and blood samples from each mouse were collected in heparin-treated tubes and were immersed in ice immediately and were then centrifuged at 300 g for 10 min. In order to obtain spleen cell suspension, spleens were removed and cut into small pieces, and then minced in PBS and filtered. The cell pellets of MPE, blood, and spleen were resuspended in PBS, and mononuclear cells were isolated by Ficoll-Hypaque gradient centrifugation (Pharmacia, Uppsala, Sweden) to determine T cell subsets within 1 h.
Animals Wild type (WT) C57BL/10 mice and TLR4–/– mice in a C57BL/10 background were purchased from the Model Animal Research Center of Nanjing University (Nanjing, China). Mice used for experiments were sex-, weight
Positron emission tomography (PET) and computed tomography (CT) scanning 1
All PET images were acquired using Trans-PET BioCaliburnTM LH (Raycan Technology Co., Ltd, Suzhou,
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China) consisting of 12 basic detector modules. Fifteenmin PET scans were begun between 60 and 70 min after retro-orbital injection of 200–300 mCi of 18-fluoro-6deoxy-glucose. In consideration of the PET system’s spatial resolution, reconstructed PET images had pixel dimensions of (0.5 mm)2 and 0.5 mm slice thickness. Furthermore, due to the OSEM reconstruction method using point spread function system response matrix, the size of the reconstructed 3D image was set to be 280 280 104 (XxYxZ) that must be smaller than or equal to the size of point spread function system response matrix. Also, CT images were acquired on a small animal CT scanner provided by the Trans-PET system with characteristics of an axial field of view of 5.0 cm, a radial field of view of 7.0 cm, and a spatial resolution of 0.5 mm. The PET and CT images were acquired with the mouse under anesthesia in a common animal holder. The coregistration of the images was implemented using the AMIDE software package (The Free Software Foundation Inc., Boston, MA).
Flow cytometry The Abs included anti–CD3, –CD4, –CD8, –IFN-g, –IL-17, –T-bet, –RORgt, –CD14, –Ly-6G, and –TLR4 mAbs, which were purchased from BD Pharmingen (San Diego, CA) or eBioscience (San Diego, CA). To determine intracellular cytokines, cells were stimulated for 4 h at 37 C with PMA (50 ng/mL, Sigma–Aldrich, St. Louis, MO) and ionomycin (1 mg/mL, Sigma–Aldrich) in the presence of Brefeldin A (10 mg/mL, Enzo Life Science). Cells were surface-stained with anti–CD3, –CD4, or –CD8 mAbs in PBS þ 2% heat-inactivated fetal bovine serum (FBS, Gibco) for 20 min at 4 C. Cells were resuspended in a fixation/permeabilization solution (Cytofix/Cytoperm; BD Pharmingen) and were incubated with anti–IFN-g or –IL-17 mAbs for 30 min at 4 C. Cells were then washed with permeabilization buffer and resuspended in PBS þ 2% FBS for flow cytometric analysis. To determine transcription factors, cells were surfacestained with anti–CD3, –CD4, or –CD8 mAbs in PBS þ 2% heat-inactivated fetal bovine serum for 20 min at 4 C. Cells were resuspended in a fixation/permeabilization solution and were incubated with anti–T-bet or –RORgt mAbs for 30 min at 4 C. Cells were then washed with permeabilization buffer and resuspended in PBS þ 2% FBS for flow cytometric analysis. To determine TLR4 expression on the surface of CD14þ monocytes/macrophages, CD4þ T cells, CD8þ T cells, and Ly-6Gþ neutrophils, cells isolated from MPE of WT mice were stained by the similar protocols except without the stimulating procedure. 1122
Flow cytometry was performed on a FACS Canto II (BD Biosciences) and the data were analyzed using BD FCSDiva Software and FCS Express 4 software (De Novo Software, Los Angeles, CA).
Quantitative real-time PCR Total RNA was isolated from cell pellets with Trizol reagent (Invitrogen, Carlsbad, CA). cDNAs were synthesized with oligo (dT) primers using the First Strand cDNA Synthesis Kit ReverTra Ace-a-(TOYOBO, Osaka, Japan). cDNAs were used as templates in real-time PCR with the SYBR Green I Real-time PCR Master Mix (TOYOBO, Osaka, Japan). DNA was amplified under the following typical cycling conditions: denaturation at 95 C for 15 s, annealing at 58 C for 20 s, extension at 72 C for 20 s. The samples were amplified for 40 cycles. The following primers were used: for Ifng, forward primer: 50 -CATCAGCAACAACATAAGCG-30 , reverse primer: 50 -GACCTCAAACTTGGCAATACTC-30 ; for t-bet, forward primer: 50 -CAACAACCCCTTTGCGAAAG-30 , reverse primer: 50 -TCCCCCAAGCAGTTGACAGT-30 ; for Il17a, forward primer: 50 -TGTGTCTCTGATGCTGTTGC-30 , reverse primer: 50 -GGAACGGTTGAGGTAGTCTG-30 ; for RORgt, forward primer: 50 TGCAAGACTCATCGACAAGG-30 reverse primer: 50 AGGGGATTCAACATCAGTGC-30 ; for b-actin, forward primer: 50 -CTGAGAGGGAAATCGTGCGT-30 , reverse primer: 50 -CCACAGGATTCCATACCCAAGA-30 . b-actin was used as an internal control, and levels of each gene were normalized to b-actin expression using the DDCt-method, with WT mouse experimental value set to 1.
In vitro differentiation of Th1 and Th17 cells Mononuclear cells from mouse spleens were isolated by Ficoll-Hypaque gradient centrifugation. Bulk CD4þ T cells were isolated by negative selection (by depletion of CD8aþ, CD45Rþ, CD11bþ, CD25þ, CD49bþ, TCRg/dþ, and Ter119þ cells) with CD4þ cell biotin-Ab cocktail II and antibiotin microbeads. After isolation of bulk CD4þ T cells, naive CD4þCD62Lþ T cells were isolated by CD4þCD62Lþ T cell isolation kit II (Miltenyi Biotec, Aubum, CA) according to the manufacturer. The purity of naive CD4þCD62Lþ T cells was more than 90%, as measured by flow cytometry. Purified naive CD4þ T cells were cultured in RPMI-1640 medium containing 10% FBS in the presence of platebound anti–CD3 (4 mg/mL) and anti–CD28 (2 mg/mL) mAbs. Cytokines used for cell differentiation were as follow: Th1 condition, IL-2 (50 U/mL), IL-12 (5 ng/mL) and anti–IL-4 mAb (10 mg/mL); Th17 condition, IL-2 (50 U/mL), TGF-b (2 ng/ml), IL-6 (100 ng/mL), IL-1b (10 ng/mL), IL-23 (10 ng/mL), anti–IL-4 mAb (10 mg/mL),
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and anti–IFN-g mAb (1 mg/mL). In some experiments, designated concentrations of lipopolysaccharide (LPS; 0, 0.5, and 1.0 mg/mL) were added into Th1 or Th17 conditions. Cells were cultured for 5 d and then stimulated for 4 h with PMA and ionomycin in the presence of Brefeldin A before being stained for cytokines described in the Flow Cytometry section.
Determination of signal transductions of IFN-g and IL-17 Purified na€ıve CD4þ T cells were cultured in RPMI-1640 medium containing 10% FBS in the presence of plate-bound anti–CD3 (4 mg/mL) and –CD28 (2 mg/mL) mAbs in Th1 or Th17 conditions as described above for 48 h. The cells were stimulated for 30 min with IL-2 (50 U/mL) and IL-12 (5 ng/mL) for phospho (p)-signal transducer and activator of transcription (STAT) 1 and with IL-6 (100 ng/mL) and IL1b (10 ng/mL) for p-STAT3. Then, the cells were harvested, fixed with 1.5% paraformaldehyde, and were stained for p-STAT1 or p-STAT3 and subjected to flow cytometry as described above.
TLR4 in pleural malignancy
RESULTS
TLR4 expression in MPE Our previous data have shown that a large proportion of nucleated cells in MPE of WT mice were mononuclear cells, lymphocytes, and neutrophils, and a very small proportion of cells were eosinophils (Li et al., 2014), we thus first explored the expression of TLR4 on these pleural cells using flow cytometry. As shown in Table 1, CD14þ monocytes/ macrophages, CD4þ T cells, CD8þ T cells, and Ly-6Gþ neutrophils in MPE of WT mice expressed much higher level of TLR4 than their counterparts in peripheral blood or spleen (all P < 0.05), and there were no differences in TLR4 expression between blood cells and spleen cells (all P < 0.05).
Effect of TLR4 deficiency on MPE formation
The observation of survival advantage included 1 group of WT mice and 5 groups of TLR4–/– mice (each n ¼ 15). At days 1, 3, 6, 9, and 12 after intrapleural instillation of LLC cells, 2 groups of TLR4–/– mice received intraperitoneal injection of 100 mg anti-IFN-g neutralized mAb (eBioscience, clone XMG1.2) (TLR4–/– þ a-IFN-g) and 100 mg mouse isotype control mAb (eBioscience), respectively; and another 2 groups of TLR4–/– mice received intraperitoneal injection 1 mg recombinant mouse IL17A protein (R&D System, Minneapolis, MN) diluted in PBS containing 0.1% albumin (TLR4–/– þ rmIL-17) and PBS containing 0.1% albumin in a total volume of 100 mL, respectively. The mice without heartbeat and respiration were determined to be dead. The administration regimens of anti-IFN-g mAb and recombinant mouse IL-17A were based upon our preliminary experiments.
Fourteen days after pleural injection of LLC cells, pleural tumors were found to distribute on the parietal and visceral pleura of WT and TLR4–/– mice. Bloody pleural fluid could be found through diaphragm, surrounding tumor foci in the two groups (Figure 1A). MPE volume in TLR4–/– mice (0.72 0.08 mL) was much larger than that in WT mice (0.34 0.07 mL; P < 0.001) (Table 2). The numbers of nucleated cells were significantly decreased in TLR4–/– mice compared with WT mice (P < 0.001). Interestingly, the absolute numbers of total nucleated cells and each cell types, including mononuclear cells, lymphocytes, neutrophils, and eosinophils, in the whole MPE, were not different between the two groups (all P > 0.05) (Table 2). FDG-PET and CT scans were performed to evaluate the pleural tumors and MPE in the living mice. As shown in Figure 1B, increased radiotracer uptake on FDG-PET scanning was found in the areas of tumors and MPE. Compared with WT mice, the PET imaging confirmed higher 18F-FDG retention in TLR4–/– mice. The maximum standardized uptake value (SUVmax) in TLR4–/– and WT mice were 8.8 1.9 and 3.7 0.7, respectively, with statistic significance between the two groups (P ¼ 0.021) (Figure 1C).
Statistics
Table 1 Percentages of toll-like receptor 4 positive cells in a murine MPE model
Survival advantage experiment
Data were normally distributed, and all values are expressed as mean SEM. Differences between two or multiple experimental groups were compared using Student’s t-test, one-way analysis of variance (ANOVA), or two-way ANOVA, as appropriate. Survival was estimated by the Kaplan-Meier method and compared by the log-rank test. Statistical analysis was performed with SPSS version 16.0 Software (Chicago, IL), and P < 0.05 was considered statistically significant.
n Monocytes/macrophages CD4þ T cells CD8þ T cells Neutrophils
MPE
Blood
Spleen
10 15.0 3.1* 7.5 0.7* 12.3 3.4* 41.7 4.1*
10 9.4 2.3 2.4 1.2 5.3 0.5 28.9 3.7
10 10.2 1.2 1.3 0.4 3.7 1.3 32.4 4.0
MPE ¼ malignant pleural effusion. *P < 0.05 compared with blood or spleen, and comparisons were determined by two-way analysis of variance (ANOVA).
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TLR4 deficiency increased Th1 cells and decreased Th17 cells in MPE Th1 cells and Th17 cells were identified by flow cytometry (Figure 2A). In both WT and TLR4–/– mice, the numbers of Th1 (Figure 2B), and Th17 cells (Figure 2C) were significantly higher in MPE (P < 0.001 for all). More importantly, we noted that Th1 cell numbers were increased and Th17 cell numbers were decreased in MPE from TLR4–/– mice as compared with WT mice (P < 0.01 for both). The numbers of Th1 and Th17 cells in blood and spleen showed no significant differences in either TLR4–/– or WT mice (P > 0.05 for all).
Expression of IFN-g, IL-17, T-bet, and RORgt mRNA in MPE We noted that there was much higher mRNA expression of IFN-g (Figure S1A) and T-bet (Figure S1B) in MPE from TLR4–/– mice than that from WT mice (both P < 0.05). In contrast, there was much lower mRNA expression of IL-17 (Figure S1C) and RORgt (Figure S1D) in MPE from TLR4–/– mice than that from WT mice (P < 0.01 for both).
Effect of TLR4 deficiency on differentiation of Th1 and Th17 cells in vitro Figure 1 Development of malignant pleural effusions (MPE) in wild type (WT) and TLR4–/– mice. (A) Fourteen days after intrapleural injection of Lewis lung cancer cells, multiple tumor foci could be seen on the parietal and visceral pleura, and MPE was visible in the pleural space. (B) Positron emission tomography and computed tomography imaging of developing pleural tumors and MPE in WT (top panels) and TLR4–/– mice (bottom panels). (C) The maximum standardized uptake value (SUVmax) on positron emission tomography were calculated (both n ¼ 10). Data are presented as means SEM, the comparisons were determined by Student’s t-test.
Na€ıve CD4þ T cells from spleens of WT or TLR4–/– mice were purified and cultured in vitro in the presence of IL-2, IL-12, and anti–IL-4 mAb (Th1 condition); or IL-2, TGF-b, IL-6, IL-1b, IL-23, anti–IL-4, and anti–IFN-g mAbs (Th17 condition). Compared with WT na€ıve CD4þ T cells, the differentiation of Th1 cells were significantly increased (Figures 3A and 3B), and the differentiation of Th17 cells were significantly decreased (Figures 3A and 3C) from TLR4–/– na€ıve CD4þ T cells. In WT mice, addition of LPS to Th1 condition inhibited Th1 cell differentiation in a dosedependent manner (Figure 3D), while addition of LPS to Th17 condition promoted Th17 cell differentiation in a dose-dependent manner as well (Figure 3E).
Table 2 Cytological characteristics in malignant pleural effusion
n Pleural fluid volume, mL Nucleated cells, 106/mL Cell differential counts, % Mononuclear cells Lymphocytes Neutrophils Eosinophils
Wild type mice
TLR4–/– mice
P-values*
15 0.34 0.07 42.8 3.4
15 0.72 0.08 21.0 4.3
