Increased resistin suggests poor prognosis and promotes development of lung adenocarcinoma
Retraction in: /10.3892/or.2023.8595
Affiliations: Department of VIP Ward, Tianjin Medical University Cancer Institute and Hospital, Tianjin 300060, P.R. China, Department of Pancreatic Cancer, Tianjin Medical University Cancer Institute and Hospital, Tianjin 300060, P.R. China, Department of Public Laboratory, Tianjin Medical University Cancer Institute and Hospital, Tianjin 300060, P.R. China, Department of VIP Ward, Tianjin Medical University Cancer Institute and Hospital, Tianjin 300060, P.R. China
- Published online on: September 26, 2018 https://doi.org/10.3892/or.2018.6736
- Pages: 3392-3404
Copyright: © Zhao et al. This is an open access article distributed under the terms of Creative Commons Attribution License.
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Resistin is a cysteine-rich protein, which is mainly secreted from monocytes and macrophages in humans (1–3). It is associated with inflammation and malignant neoplasms (4–5). Blood resistin levels are demonstrated to be increased in certain cancer patients compared with healthy controls, including esophageal squamous cancer, gastric, colorectal, breast and endometrial cancer, and malignant lymphoma (6–12). Resistin is considered to be a risk factor for breast cancer (9,13) and biomarker of disease progression of esophageal squamous cancer, gastric and colorectal cancer (6–8). It is an independent prognostic factor of pancreatic ductal adenocarcinoma (5). Resistin can promote prostate cancer cell proliferation through the phosphatidylinositol 3 kinase (PI3K)/protein kinase B signaling pathway in human prostate cancer cell lines PC-3 and DU-145 (14).
However, most of the reported studies only demonstrated the association between serum or plasma resistin and malignancy. Few reports measured the level of resistin expression in cancer tissues, even though it is less well studied and controversial in lung cancer. Certain reports demonstrated a higher concentration of resistin in the blood was demonstrated in non-small cell lung cancer (NSCLC) patients compared with the controls (15–17). One of the reports assessed resistin expression in the marginal area of lung cancer tissue and non-cancer region by immunofluorescence staining in 10 cases (17). Another revealed that blood resistin levels were similar between cancer group and non-cancer group (18). Furthermore, the clinical significance and biological function remain largely unknown.
However, lung cancer is one of the most common malignancies worldwide, with higher morbidity and poorer prognosis (19–20). The most common form of lung cancer is NSCLC, which includes lung adenocarcinoma and squamous carcinoma. At present, lung adenocarcinoma replaces squamous carcinoma as the dominating type of NSCLC. The aim of the present study is to determine the resistin expression in lung adenocarcinoma tissues, clinical significance and biological function in vitro and in vivo.
Materials and methods
Patients and tissue samples
A total of 70 consecutive cases of newly diagnosed lung adenocarcinoma patients at Tianjin Medical University Cancer Institute and Hospital (Tianjin, China) from January to December 2008, with complete clinical and pathological data, were selected retrospectively in the present study and followed up for at least five years. Paired cancer and adjacent non-cancerous tissue samples, which were located more than 1 cm away from the tumor, were obtained through open surgeries. The paraffin-embedded tissue samples were stained with hematoxylin-eosin and confirmed lung as adenocarcinoma again.
The clinicopathological characteristics of the patients were recorded. The tumor staging of NSCLC was defined according to the tumor, node and metastasis system. The study was comprised of 70 cases of lung adenocarcinoma (38 male cases, 32 female cases), with an average age of 61 years old (36–77 years old). All patients received treatments (including operation, chemotherapy or radiotherapy), which conformed to the guidelines of NSCLC.
For immunohistochemical staining, 5-µm paraffin-embedded tissue sections were heated for 1 h at 70°C, deparaffinized with a xylene soak, followed by rehydration via the addition of alcohol at decreasing concentrations (100, 95, 85 and 75%) for 5 min/step. A 96°C water-bath was used for antigen retrieval in 0.01 mol/l sodium citrate buffer (10 mM, pH 6.0) for 20 min. Next, endogenous peroxidase activity was quenched by incubation in 3% hydrogen peroxide for 15 min at room temperature. Subsequently, slides were blocked with goat serum (cat. no. ZLI-9022; OriGene Technologies, Inc., Beijing, China; 1:1) at 37°C in a wet box for 30 min and then incubated by the primary antibody (rabbit polyclonal antibody against human resistin; cat. no. BS7730; Bioworld Technology, Inc., St. Louis Park, MN, USA; 1:100) overnight at 4°C in moist chambers. Following washing with 0.01 mol/L PBS (pH 7.2) three times, slides were incubated with a biotinylated secondary rabbit anti-mouse antibody (cat. no. PV-6000; OriGene Technologies, Inc.; 1:1) for 25 min at 37°C. Following incubating in horseradish peroxidase marked streptomycin avidin working fluid at 37°C for 30 min, slides were treated with avidin biotin-peroxidase complex using 3,3′-diaminobenzidine as a chromogen, and then counterstained with hematoxylin for 30 sec at room temperature and examined by light microscopy (Olympus Corporation, Tokyo, Japan). PBS was used as a negative control instead of a primary antibody.
Evaluation of immunohistochemical staining
Existing tan or brown particles in the nucleus or cytoplasm indicated positive cells, which must conform to the following conditions: i) The cellular structure was clear; ii) the location of positive granules was accurate; iii) staining was significantly increased compared with the background.
In a 400× high power filed, randomly selected from 10 different cancer cell fields of view, the percentage of (a) positively stained cells was calculated as follows: 0–5% positive cells, score 0; 6–25% positive cells, score 1; 26–50% positive cells, score 2; 51–75% positive cells, score 3; 76–100% positive cells, score 4. Then, the (b) staining intensity was evaluated: Colorless, score 0; light yellow, score 1; deep yellow and tan, score 2; brown, score 3.
The expression of resistin was based on the product of (a) × (b): Score 0, negative (−); score 1–4, weakly positive (+); score 5–8, positive (++); score 9–12, strongly positive (+++). (−) and (+) were regarded as low expression group, (++) and (+++) were categorized as the high expression group (Fig. 1A). The result of each specimen was independently evaluated by two qualified and expert pathologists, blinded to the patients' clinical data. The few cases with discordant results were reevaluated and final scores were consensual.
The expression of resistin in lung adenocarcinoma tissues and its association with survival. (A) Immunohistochemical staining of lung adenocarcinoma tissues with a resistin antibody. The low and high expression of resistin in lung adenocarcinoma tissues are presented (magnification, ×100). High resistin expression indicated poor prognosis in lung adenocarcinoma patients. (B) PFS a curves of 70 patients with different level of resistin expression are presented. The PFS curves of age, smoking, tumor size, lymph node metastasis and clinical stage are presented. (C) OS curves of 70 patients with different level of resistin expression are presented. The OS curves of age, smoking, tumor size, lymph node metastasis and clinical stage are presented. PFS, progression-free survival; OS, overall survival; Cum, cumulative; (−) and (+), low expression group, (++) and (+++), high expression group.
Human lung adenocarcinoma cell lines A549 and H1975 were obtained from the Committee of Type Culture Collection of the Chinese Academy of Sciences (Shanghai, China). The cells were maintained in RPMI-1640 (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.) at 37°C in a humidified atmosphere of 95% air and 5% CO2. Cells were divided according to transfection of overexpression resistin plasmid for the resistin group and empty vector for the control group.
Transfection and isolation of stable transfectants
Lipofectamine™ 2000 Reagent (Invitrogen; Thermo Fisher Scientific, Inc.), endo-free maxiplasmid kit (Tiangen Biotech Co., Ltd., Beijing, China); pcDNA3.1-(+)/resistin plasmids were established by OriGene Technologies, Inc. (cat. no. RC210942) and the primers were as follows: Forward primer, 5′-CCCACCGAGAGGGATGAAAG-3′ and reverse primer, 5′-CAGTGACATGTGGTCTCGGC-3′; forward primer, 5′-CAGCTCACCATGGATGATGATATC-3′ and reverse primer, 5′-AAGCCGGCCTTGCACAT-3′ (β-actin).
A fragment of the rat resistin cDNA fragment (285 bp) was inserted at the unique EcoRI site in the anti-sense orientation as determined by sequencing. The final concentration of resistin was 100 nM. A total of 2×106 A549 and H1975 cells grown in 60 mm Petri dishes were transfected with 10 µg of the recombinant plasmid using lipofectamine, as described by the supplier (Gibco; Thermo Fisher Scientific, Inc.). A total of 24 h later, fresh RPMI-1640 media containing 10% FBS was added and replaced 48 h later. Monoclonal cells were selected with NeoR. Then the cells were cultured for 24 h following transfection.
Reverse transcription-quantitative polymerase chain reaction (RT-qPCR)
Total RNA was extracted from the cells by TRIzol (cat. no. 15596026; Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. Prime Script RT reagent kit (DRR037A; Takara Bio, Inc., Otsu, Japan) were used for cDNA generation (42°C for 30–60 min, 70°C for 15 min). RT-qPCR was performed with Super Real PreMix (cat. no. FP204-01; Tiangen Biotech, Co., Ltd., Beijing, China) by the following program: 95°C for 3 min in 1 cycle; 95°C for 5 sec, 58°C for 30 sec, and 72°C for 30 sec in 35 cycles. To ensure the DNA production, a melting curve analysis was performed according to ABI Step One system. The relative gene expression was normalized to the internal standard β-actin using 2−ΔΔCq method (4,17,21). The primers are 5′-TGGAGTGCCAGAGCGTCACCT-3′ (forward) and 5′-ACTGGCAGTGACATGTGGTCTC-3′ (reverse).
Western blotting was used to verify successful transfection. Whole cell extracts of A549 and H1975 were prepared with a CellLytic™ M reagent (cat. no. C2978; Sigma-Aldrich; Merck KGaA, Darmstadt, Germany), the protein was quantified by a Bicinchoninic acid (Pierce; Thermo Fisher Scientific, Inc.) assay. Then, the protein samples (50 µg) were separated by 10% SDS-PAGE. The samples were blocked with goat serum (cat. no. ZLI-9022; OriGene Technologies, Inc.; 1:1) for 60 min at room temperature and detected by western blotting using rabbit polyclonal resistin antibody (cat. no. BS7730; Bioworld Technology, Inc.; 1:500), mouse anti-β-actin (cat. no. ab8226; Abcam, Cambridge, UK; 1:1,000), mouse anti-proliferating cell nuclear antigen (PCNA; cat. no. ab29; Abcam; 1:500), rabbit anti-Ki67 (cat. no. ab16667; Abcam; 1:1,000), rabbit polyclonal to caspase-3 (cat. no. ab13847; Abcam; 1:500), rabbit monoclonal to caspase-7 (cat. no. ab32522; Abcam; 1:1,000), mouse anti-matrix metalloproteinase (MMP)2 (cat. no. ab37150; Abcam; 1:1,000) and mouse anti-MMP9 (cat. no. ab38898; Abcam; 1:1,000) monoclonal antibodies at 4°C overnight, and the secondary antibody was goat anti-rabbit horseradish peroxidase (cat. no. ZDR-5306; ZSGB-BIO; OriGene Technologies, Inc.; 1:10,000) for rabbit polyclonal resistin antibody, rabbit anti-Ki67, rabbit polyclonal to caspase-3, rabbit monoclonal to caspase-7 or goat anti-mouse IgG(H+L)-HRP (cat. no. LK2003; Tianjin Sungene Biotech Co., Ltd.; 1:5,000) for mouse anti-β-actin, mouse anti-proliferating cell nuclear antigen, mouse anti-MMP2 and mouse anti-MMP9 monoclonal antibodies. The gray values (cat. no. C8420; Coomassie brilliant blue G-250, Beijing Solarbio Science & Technology Co., Ltd., Beijing, China) were analyzed by using the Odyssey V3.0 software (Thermo Fisher Scientific, Inc.).
Cell proliferation assay by colony formation
A549 and H1975 Cells were cultured in 6-well plates, 300 cells/well prior to being fixed in methyl hydrate room temperature for 10 min. Then the colonies were stained by 1% Crystal Violet Staining Solution at room temperature for 5 min and counted with a light microscope (Inverted microscope; Leica Microsystems GmbH, Wetzlar, Germany).
MTT assay was also used to observe and compare cell proliferation ability of A549 and H1975. A total of 2×103 cells were plated into a well of 96-well plates and 10 ml, 5 mg/ml MTT was added to each well and continued to culture for 4 h. Then following dimethyl sulfoxide addition, the plates were placed on a microplate autoreader (Bio-Rad, Laboratories, Inc., Hercules, CA, USA). Optical density was read at 570 nm wavelength and cell growth curves were determined according to the optical density value.
Apoptosis analysis by flow cytometry
The A549 and H1975 cells were dosed 24 h following plating and then tested according to the protocol of Biolegend kit (cat. no. 640906, Biolegend, Inc., San Diego, CA, USA). Cells were resuspended in Annexin V binding buffer at a concentration of 106 cells/ml. Following transferring 100 µl cell suspension to 5 ml test tube, 5 ml of Annexin V-fluorescein isothiocyanate and 10 µl of propidium iodide solution were added to the cell suspension. A total of 400 µl binding buffer was added to each tube 15 min later, the apoptosis was analyzed using a flow cytometer (CytExpert analysis software 2.0; Beckman Coulter, Inc., Brea, CA, USA).
Cell scratch assays
The A549 and H1975 cells were seeded to full confluence in 6-well plates overnight. A scratch was introduced in the middle of the well with a sterile pipette tip the following day. The medium was discarded and replaced. The rate of migration towards the center of the wound was determined using calipers in the image under a light microscope 48 h later.
Cell invasion assays
The invasion assays were performed with an 8.0-µm pore inserts in a 24-well Transwell chambers (Corning Incorporated, Corning, NY, USA). The A549 and H1975 cells (2×105/well) and Dulbecco's modified Eagle's medium were added to the upper chamber of Transwell coated with Matrigel (BD Biosciences, Franklin Lakes, CA, USA). The RPMI-1640 with 10% FBS was added to the lower chamber and incubated at 37°C for 24 h. Cells that had migrated to the bottom of the filter were stained with a three-step stain kit (Thermo Fisher Scientific, Inc.) at room temperature for 5 min. The cells were counted by light microscope from each chamber. A total of 5 fields of view were counted.
Xenografts assays in vivo
All surgery was performed under sodium pentobarbital anesthesia. A total of 3% sodium pentobarbital (50 mg/kg) by intraperitoneal injection was used. Athymic BALB/c nude mice (16 female, aged 5 weeks, 25 g) were provided by Slac Laboratory Animal Co., Ltd. (Shanghai, China). Mice were housed in a pathogen-free animal facility at 18–29°C. The humidity is 40–70%. The mice had access to food and water with 12-h light/dark cycle. They were randomly divided into 2 groups according to the A549 cell groups described above. There were 8 mice/group. A total of 0.1 ml serum-free RPMI-1640 with 2×106 cells were subcutaneously injected into the right flank of each mouse. The control mice were injected with stable A549 cell lines transfected with empty vector. The tumors were measured by vernier caliper on day 14, 17, 21, 23, 26 and 29. Mice were sacrificed at 29 days post inoculation. The final volume of tumor tissues was calculated with the following equation: Tumor volume (mm3) = tumor length (mm) × tumor width (mm) × tumor height (mm)/2.
Each experiment was repeated three times. All statistical analyses were performed using SPSS 20.0 software (IBM, Corps., Armonk, NY, USA). The Spearman method was used to analyze the correlation of resistin expression with clinicopathological variables. Kaplan-Meier method was used to perform survival analysis and evaluate the differences between survival curves by log-rank test. The hazard ratio and confidence interval was calculated by univariate and multivariate Cox regression model. The experiments' results in vitro and in vivo were recorded as the mean ± standard deviation. A student's two-sided t-test was used to compare values of test and control samples. P<0.05 was considered to indicate a statistically significant difference.
The expression of resistin in lung adenocarcinoma tissues
Lung adenocarcinoma tissues exhibited different levels of resistin expression and adjacent normal lung tissues/non-cancer tissues stained negative. A total of 41.4% of cancer tissues demonstrated high resistin expression (58.6% demonstrated low resistin expression; Table I). The expression of resistin was different between ≤65 and >65 years, tumor size ≤3 and >3 cm, non-lymph node metastasis and lymph node metastasis as well as early stage and advanced stage (Table I).
The association between resistin expression and clinicopathological characteristics
The expression of resistin in lung adenocarcinoma tissues was significantly, positively correlated with tumor size, lymph node status and clinical stage (P<0.05) and significantly, negatively correlated with progression-free survival (PFS) and overall survival (OS; P<0.01). There is no correlation with age at diagnosis, smoking, drinking, body mass index (BMI) and blood type (Table II).
Increased PFS and OS were observed in the patients with low resistin expression and low expression groups as determined by the log-rank test. Comparing the low resistin expression group with the high one, the prognosis of the former was improved (P<0.01; Fig. 1B and 1C). Survival analysis demonstrated that factors were significantly associated with PFS and OS, including smoking, age, lymph node status, resistin expression, clinical stage and chemoradiotherapy (P<0.05). The last three one were independent risk factors of PFS and OS in patients with lung adenocarcinoma (Table III).
The influence of resistin on biological behavior of A549 and H1975 cell lines
To test the influence of resistin on biological behavior of A549 and H1975 cell lines, the resistin overexpression cell lines was established through resistin plasmids. Although the expression of resistin in the lung adenocarcinoma cell lines was demonstrated, in the reported studies, the level of resistin in lung adenocarcinoma cells A549 and chondrosarcoma cells was low (17,21). The results of western blotting also indicated that the bands of resistin in cell lines A549 and H1975 were weak. In addition, one study reported that the resistin gene was transfected into the PC-3 cells to assess the effect of overexpression of resistin in prostate cancer cell line PC-3 (14). Same as above, the lung adenocarcinoma cells are not resisitin-dominant expression cells and secretory cells, the resistin level using overexpression was manipulated rather than being knocked-down in A549 cells and H1975 cells. The level of resistin was detected by RT-qPCR and western blotting in A549 and H1975 cell lines, which demonstrated a significantly increased level compared with the control (P<0.05; Fig. 2). Then, colony formation and MTT assays were performed to test the proliferation of A549 and H1975 cells, and the results demonstrated that compared with the control cells, stable overexpressed resistin cells obviously increased the ability of proliferation in vitro (Fig. 3A and B). Furthermore, to investigate the mechanism, the expression level of proliferation-associated proteins including PCNA and Ki67 were tested, and the results demonstrated that the protein expression of PCNA and Ki67 significantly increased in the group resistin compared with the control (P<0.05; Fig. 3C and D). Taken together, the results of the present study indicated that resistin could regulate the proliferation by increasing the associated proteins including Ki67 and PCNA in vitro.
The mRNA and protein expression levels of resistin in the established stable A549 and H1975 cell lines. According to whether cells had been transfected with an empty vector or overexpression resistin plasmid, cells were divided into the control group or resistin group, respectively. The resistin mRNA and protein level of the groups was detected by RT-q-PCR and western blotting. (A) The mRNA expression level of resistin was demonstrated by RT-q-PCR. (B) The protein expression level of resistin is presented by western blotting. *P<0.05 vs. the control. Reverse transcription-quantitative polymerase chain reaction, RT-qPCR.
Influence of resistin on the proliferation of A549 and H1975 cells. (A) Colony formation and (B) MTT arrays were performed to investigate the proliferation of tumor cells. The colony numbers and OD value in resistin group were higher compared with the controls. The expression of the proliferation associated proteins (C) PCNA and (D) Ki67 were analyzed by western blotting. *P<0.05 vs. the control. OD, optical density; PCNA, proliferating cell nuclear antigen.
Flow cytometry was used to detect alterations in cell apoptosis and the results indicated that the resistin overexpressing A549 and H1975 cells demonstrated significantly decreased apoptotic ability (P<0.05; Fig. 4A). Furthermore, the results were confirmed by detecting the level of the apoptosis associated proteins, including caspase-3 and caspase-7. A significantly decreased level in the resistin overexpression group was demonstrated compared with the control group (P<0.05; Fig. 4B and C).
Influence of resistin on the cell apoptosis. (A) Flow cytometric analysis demonstrated the number of apoptotic cells in the resistin group was significantly decreased compared with the control group. The results of western blotting demonstrated that the expression of the apoptosis associated proteins (B) caspase-3 and (C) caspase-7 were significantly decreased in the resistin group compared with the control group. *P<0.05 vs. the control.
In addition, a scratch test and Transwell invasion assay was used to detect alterations in cell migration and invasion. The results were as follows: Compared with the control cells, the resistin overexpression cells demonstrated significantly reduced scratches and more invading cells through the membrane (P<0.05; Fig. 5A and B). Resistin promoted A549 and H1975 cell migration and invasion in vitro. Furthermore, to investigate the mechanism of the above, the expression level of the proteins that reflect invasion and metastasis including MMP2 and MMP9 was tested. The expression of MMP2 and MMP9 was demonstrated to be significantly increased in the resistin group compared with the control (P<0.05; Fig. 5C and D). In summary, the results indicated that resistin could strengthen the invasive capacity of cancer cells by regulating the associated proteins including MMP2 and MMP9.
Influence of resistin on the migration and invasion of A549 and H1975 cells. (A) A scratch assay demonstrated that the width in resistin group was decreased. (B) Transwell assay was used to measure the invasion of tumor cells. Compared with the controls, the number of cells within the lower chamber in resistin group was significantly increased (magnification, ×400). The results of western blot demonstrated that the expression of (C) MMP2 and (D) MMP9 proteins was upregulated in the resistin group. *P<0.05 vs. the control.
The results above indicated that resistin could promote A549 and H1975 cell proliferation, migration and invasion as well as suppress apoptosis in vitro.
The promotional effect of resistin for lung cancer in vivo
The biological roles of resistin in lung cancer tumorigenesis were further examined by xenograft studies in nude mice. A549 cells transfected with resistin plasmids or an empty vector were inoculated subcutaneously into the upper back of nude mice. Following four weeks the tumor volume of resistin group was significantly larger compared with the control group (P<0.05), as presented in Fig. 6A and B. To validate the expression of resistin in mice tumors, immunoblotting analysis was performed. The results revealed that the resistin protein level was significantly increased in the resistin plasmids group (P<0.05; Fig. 6C). Taken together, these indicated the tumorigenic effect of resistin in vivo.
The enhancing effect of resistin for lung adenocarcinoma in vivo. (A) The tumors in different groups were harvested following 4 weeks. The tumor size of resistin group was notably larger compared with the controls. (B) The growth curve of tumors was recorded 14, 17, 21, 23, 26 and 29 days following the injection of cells. The tumors grew faster in the resistin group and the size of tumors was also larger compared with those in the control group. (C) Western blotting was performed to confirm the effect of resistin overexpression in tumor tissues of nude mice. These results indicated the tumorigenic effect of resistin in vivo. *P<0.05 vs. the control.
In the present study, the resistin expression of 70 patients with lung adenocarcinoma was analyzed by immunohistochemistry. Resistin expression was demonstrated to be increased in lung adenocarcinoma tissues compared with the paired adjacent normal lung tissues. It was in accordance with previous reports that NSCLC patients exhibited a higher blood level of resistin in contrast to controls (15–17). Compared with the non-cancerous regions, Kuo et al (17) also demonstrated a higher level of resistin in the marginal areas of human lung cancer tissue by immunofluorescence staining. It was also similar to that presented in cancer tissues of breast cancer, colorectal cancer, pancreatic ductal adenocarcinoma and prostate cancer (5,13–14,22).
It was also demonstrated that the expression of resistin in lung adenocarcinoma tissues increased as the size of tumor and clinical stage of the cancer progressed. The resistin level was increased in patients with lymph node metastasis compared with the ones without lymph node metastasis. As well as in breast cancer tissues, the resistin expression was positively associated with tumor stage, tumor size and lymph node status (13,23). In pancreatic ductal adenocarcinoma, resistin expression was strongly and positively associated with tumor stage (5). The blood level of resistin was also elevated with progression in tumor stage in patients with gastric cancer and colorectal cancer (7,24). In general, resistin is correlated with poor clinicopathological status.
It was also demonstrated that there was no correlation of resistin expression with sex, age at the point of diagnosis, smoking, drinking and blood type in patients with lung adenocarcinoma in the present study. To summarize the existing reports, resistin was associated with sex in certain types of cancer, which mainly occurred in females, including breast cancer and endometrial cancer (10,11); but in certain cancer that mainly occurs in males, resistin level demonstrated no significant sex differences, including esophageal (6), gastric (7,25–26), colorectal (1–2,8,27–28) and pancreatic cancer (29). Karapanagiotou et al (16) also demonstrated that serum resistin level was unassociated with sex, age and BMI in NSCLC patients. In the present study, the expression of resistin in lung adenocarcinoma tissues also demonstrated no sex differences prior to and following being divided into low and high expression groups. This may be due to there being no significant sex differences for resistin in lung adenocarcinoma patients; or that the morbidity of lung cancer is increasing annually in female in recent years, the difference between male and female is decreasing; or that the sample size is small in the present study. Further research with an expanded sample size is required.
Nevertheless, resistin expression level exhibited a negative correlation with PFS and OS by bulk analysis. Increased PFS and OS were observed in the patients with low resistin expression (−/+) and in the low resistin expression groups (− to +). The prognosis of the low expression group was improved compared with the high one (++ to +++). Multivariate Cox regression analysis demonstrated that resistin expression, clinical stage and chemoradiotherapy were independent prognostic factors of PFS and OS in the patients with lung adenocarcinoma.
To a certain extent, this was in line with a few reports. Higher resistin expression in cancer tissues was associated to poor prognosis in breast cancer patients (13,23). It may be an independent prognostic factor of breast cancer and pancreatic ductal adenocarcinoma (5,13).
With respect to lung cancer, one report demonstrated that the serum resistin levels exhibited a trend of association with time to relapse, but it was not a predictive factor for OS (15). Another study reported that it was unconnected with diagnosis and prognosis (16). However, the present study's results were mainly discussing resistin level within the tumor tissues. Most researchers believed that the levels of adipokine (including leptin, resistin and adiponectin) in blood circulation cannot accurately reflect their true levels in human body (30–31). Resistin level in cancer tissues or the tumor microenvironments was higher than in blood circulation and more closely correlated with tumorigenesis and tumor progression (30–31). It demonstrated that NSCLC cancer tissue-specificity and is more reliable than circulating level.
Furthermore, a series of in vitro and in vivo assays were performed. Resistin overexpression was verified in A549 and H1975 cell lines, and could promote cell proliferation, migration and invasion as well as inhibit apoptosis in vitro, and serve a tumorigenic function of lung adenocarcinoma in vivo. The present study demonstrated suggested that resistin may work by increasing proliferation associated proteins Ki67 and PCNA, while decreasing the apoptosis associated proteins caspase-3 and caspase-7. In 2015, it was demonstrated that resistin could promote chondrosarcoma metastasis and MMP2 expression through activation of the AMP-activated protein kinase/p38 signaling pathway and downregulation of microRNA-519d expression (21). Consistent with the above study, it was demonstrated that resistin may promoted lung adenocarcinoma migration and invasion by increasing MMP2 and MMP9. Resisitin also promoted breast cancer progression via Toll-like receptor 4 (TLR4)/nuclear factor (NF)-κB/signal transducer and activator of transcription 3 signaling (23). Recently resistin was reported to be strongly expressed in lung adenocarcinoma tissues. Resistin promoted lung adenocarcinoma metastasis via TLR4/Src/epidermal growth factor receptor/PI3K/NF-κB pathway (32). These studies have important referential significance and value for the potential molecular mechanisms of resistin in lung adenocarcinoma.
There were certain limitations in the present study. First, the study only investigated adenocarcinoma and not other pathological subtypes of NSCLC. Nevertheless, lung adenocarcinoma is considered as the main type of NSCLC at present. One study reported that the serum level of resistin was not correlated to the histological type of NSCLC (15). Furthermore, in the present study, the potential molecular mechanism was only investigated in vitro.
In conclusion, the expression of resistin in pathological tissues and its association with corresponding clinicopathological data in 70 consecutive patients with lung adenocarcinoma was studied for the first time to the best of our knowledge. It was demonstrated that high resistin expression was predominantly observed in lung adenocarcinoma tissues. It is associated with a more malignant clinicopathological status and poorer survival. Analysis demonstrates resistin expression is an independent prognostic factor for PFS and OS. Resistin could promote A549 and H1975 cell proliferation, migration and invasion while inhibit their apoptosis in vitro. Resistin also serves a tumorigenic function in vivo. The present study will be helpful to make clear the exact role of resistin in lung adenocarcinoma.
The present study was supported by the National Natural Science Foundation of China (grant no. 81401957) and Tumor translational medicine seed fund of Tianjin Medical University Cancer Institute and Hospital (grant no. 1317).
Availability of data and materials
The data used during the present study are available from the corresponding author upon reasonable request.
CCZ, JC, RFN and CGZ conceived and designed the study. CCZ, JC and YL collected the data. CCZ, JC and RFN performed the data analysis and interpretation. CCZ and JC wrote the manuscript and revised the important intellectual content. All authors edited and approved the final manuscript.
Ethics approval and consent to participate
The research involving human samples and animal experiments had been approved by the Ethics Committee of Tianjin Cancer Hospital (Tianjin, China). All experiments were conducted according to relevant national and international guidelines. Informed consent was obtained from all participants included in the study.
Patient consent for publication
Informed consent was obtained from all participants included in the study.
The authors declare that they have no competing interests.
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