Introduction
Human epidermal growth factor receptor 2 (HER2) is amplified in 15% to 20% of breast cancers, and its overexpression is associated with adverse prognosis [
1]. Trastuzumab, a humanized monoclonal antibody directed against HER2, was approved in 1998 for the treatment of HER2-overexpressing breast cancer. Mechanisms of action of trastuzumab include inhibition of HER2 dimerization, direct induction of cell growth arrest and apoptotic cell death, inhibition of HER2 shedding, and recruitment of immune effector cells to mediate tumor lysis [
2]. This latter mechanism, designated as antibody-dependent cellular cytotoxicity (ADCC), has been shown to be dependent on expression of Fc receptors (FcRs) by innate immune cells [
3]. Recent studies by Park
et al. also demonstrated the involvement of adaptive immune cells in the action of anti-HER2/neu antibody [
4]. As a single agent or in combination with chemotherapy, trastuzumab has shown remarkable efficacy [
5]. However, not all patients respond to trastuzumab and some patients whose breast cancer initially responds to treatment eventually experience progression, corresponding to primary and acquired resistance to trastuzumab, respectively [
5].
Different mechanisms of resistance to trastuzumab have been reported. Downregulation of phosphatase and tensin homolog (PTEN), a phosphatase whose activation contributes to trastuzumab activity, has been shown to confer trastuzumab resistance both
in vitro and
in vivo [
6]. Patients with PTEN deficiency displayed poorer responses to trastuzumab-based therapy than those with active PTEN [
6]. Shedding of the extracellular domain of HER2 protein by proteolytic cleavage has been shown to neutralize the antitumor effects of trastuzumab [
7]. Elevated circulating levels of HER2 have also been correlated with disease progression in patients treated with trastuzumab-based therapy [
8]. Furthermore, masking of the HER2 antigens by the glycoprotein mucin 4 (MUC4) has been shown to reduce binding of anti-HER2 antibodies
in vitro [
9], whereas increased MUC4 levels have been observed
in vivo in tumors that were resistant to anti-HER2 therapies [
10]. Additionally, activation of other signaling pathways, notably insulin-like growth factor 1 (IGF-1) receptor, has also been reported to inhibit trastuzumab-mediated growth inhibition in breast cancer cells [
11].
The aforementioned mechanisms of resistance are related to alterations in the tumor cells themselves and do not take into account the impact of the tumor microenvironment. This latter phenomenon is highly complex in terms of cellular composition with different cell types, including adipocytes, preadipocytes, endothelial cells, pericytes and immune cells. Several studies have shown that immune suppressor cells, such as tumor-associated macrophages, myeloid-derived suppressor cells and regulatory T cells, are recruited to the tumor sites and promote immune evasion [
12-
14]. However, the implication of resident cells, notably adipocytes, in tumor resistance to trastuzumab remains largely unknown.
Adipocytes are the most abundant cells in the breast adipose tissue. It has been shown that adipocytes are not simply energy storage depots but also active sources of various paracrine and endocrine factors, termed
adipocytokines, such as leptin, adiponectin, tumor necrosis factor (TNF)-α and interleukin (IL)-6 [
15]. Given their abundance and their proximity to tumor cells, adipocytes have been shown to affect tumor behavior, including tumor growth and metastasis [
16]. Moreover, recent studies suggested the implication of adipocytes in resistance to radiotherapy [
17] and chemotherapy [
18]. Furthermore, obesity, which is associated with increased fat mass and inflammatory adipose tissue phenotype, has been shown to be an important risk factor for breast cancer in postmenopausal women [
19]. Chemotherapy and endocrine therapy have also been observed to be less effective in patients with breast cancer and obesity [
20].
Adipocytes derive from precursors, termed
preadipocytes, through a tightly controlled differentiation process. These preadipocytes are known to possess a secretion profile distinct from that of adipocytes [
21]. Increased evidence has shown that preadipocytes contribute to tumor progression and resistance to chemotherapy [
22].
In this study, we evaluated whether preadipocytes and adipocytes could contribute to resistance to trastuzumab. We show that preadipocytes and adipocytes inhibited trastuzumab-mediated ADCC in breast cancer cells via secretion of soluble factors, resulting in a direct protective effect on tumor cells.
Methods
Cell culture
BT-474, SK-BR-3, MDA-MB-453, and MDA-MB-361 human breast cancer cell lines, all obtained from the American Type Culture Collection (Manassas, VA, USA), were cultured in complete RPMI 1640 medium (supplemented with 10% fetal calf serum (FCS), 2 mM l-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin). Human multipotent adipose-derived stem cells (hMADS) were provided by Dr Christian Dani, UMR 6543 CNRS, Nice, France, with approval granted by the local ethics committee of the University of Nice. hMADS were cultured in complete Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2.5 ng/ml basic fibroblast growth factor (FGF2). The immortalized human mammary epithelial hTERT-HME1 cells were cultured in complete DMEM/F-12 medium supplemented with 1% nonessential amino acids, 50 μg/ml gentamicin, 3.5 μg/ml insulin, 100 ng/ml epidermal growth factor and 500 ng/ml hydrocortisone. All cells were maintained at 37°C in presence of 5% CO2, except in hypoxic experiments, in which cells were maintained in 1% O2.
Induction of human multipotent adipose-derived stem cell differentiation
The differentiation of hMADS cells was previously described [
23,
24]. Differentiated hMADS (#hMADS) were used at day 14 of differentiation. The differentiation yield was estimated to range between 70% and 80% and was verified by Oil Red O staining, as described previously [
25]. hMADS and #hMADS were incubated in culture media containing 10% FCS for 2 days, and the conditioned media (CM) were harvested after centrifugation at 300 ×
g for 5 minutes and frozen at −20°C before use.
Retroviral transduction of NK-92 natural killer cells
NK-92, the human natural killer (NK) cell line [
26], generously provided by Conkwest (Del Mar, CA, USA), was grown in complete RPMI 1640 culture medium. NK-92-CD16 cells were obtained by transduction of pMX/CD16 plasmid [
27], using retroviral supernatant as described in Additional file
1.
Isolation and differentiation of human adipose-derived stem cells
Adipose tissues were provided by Dr Emmanuel Delay (Centre Léon Bérard, Lyon, France). They were obtained by liposuction from abdominal fat of patients undergoing plastic surgery. Written patient consent was obtained, and this protocol was approved by the Lyon research ethics committee. Adipose tissue samples (5 to 10 g) were rapidly digested with 100 collagen digestion units/ml collagenase (Sigma-Aldrich, St Louis, MO, USA) at 37°C with agitation for 30 minutes. Digestion was stopped by addition of complete DMEM/F-12 medium. After centrifugation at 300 ×
g for 7 minutes, cells corresponding to the stromal vascular fraction were seeded for adherence and amplification in complete DMEM/F-12 medium supplemented with 10 ng/ml FGF2. Adipose-derived stem cells (ASCs) were used at passage 2 or 3 and verified by flow cytometry to be CD14−, CD45−, CD73+, CD90+, CD105+ and HLA-ABC+ (human leukocyte antigen abacavir). Differentiation of ASCs into adipocytes was performed as described previously [
28].
Antibody-dependent cellular cytotoxicity assay
Human mammary epithelial hTERT-HME1 and hMADS cells were seeded at 1.2 × 105 cells/ml/well in 12-well plates. The medium of 12-day #hMADS was changed and supplemented with 10% FCS. Two days later, target cells were labeled with 12.5 μM calcein-AM (Sigma-Aldrich) for 30 minutes at 37°C and added at 105 cells/0.5 ml/well. NK-92-CD16 cells were added at 5 × 105 cells/0.5 ml/well (effector to target (ratio = 5:1), with trastuzumab at 1 μg/ml final concentration. After 4 hours of incubation at 37°C, the supernatants in each well were harvested and measured for fluorescence signals at 485/535 nm. Triton X-100 was added to the control wells to estimate total cell lysis. Cytotoxicity was calculated using the following formula: percent cytotoxicity = (experimental lysis − spontaneous lysis) ÷ (maximal lysis − spontaneous lysis) × 100.
Flow cytometry
Cells were labeled with corresponding antibodies for 15 minutes at room temperature and analyzed with a BD LSR II flow cytometer using BD FACSDiva software (BD Biosciences, San Diego, CA, USA) and FlowJo software (Tree Star, Ashland, OR, USA). The antibodies used are detailed in Additional file
1.
Enzyme-linked immunosorbent assay
Supernatants obtained from NK cells cultured alone or from ADCC assays in the presence of adipose cells were subjected to enzyme-linked immunosorbent assay (R&D Systems, Minneapolis, MN, USA) according to the manufacturer’s instructions.
Fluorescence microscopy
BT-474 cells were seeded overnight onto Nunc Lab-Tek chambered coverglasses (Thermo Fisher Scientific, USA). CM from #hMADS or hMADS cells were added, and cells were incubated for additional 4 hours at 37°C before being labeled on ice with anti-ErbB 2 Affibody fluorescein isothiocyanate (FITC) (Abcam, Cambridge, UK) for 15 minutes. After the washing, cells were observed with a confocal Zeiss LSM 780 microscope (Carl Zeiss, Oberkochen, Germany).
Microarray
BT-474 cells or SK-BR-3 cells were exposed to #hMADS-CM for 2 hours. RNA was extracted using the QIAamp RNeasy Mini Kit (QIAGEN, Valencia, CA, USA). After RNA amplification with the Illumina TotalPrep RNA Amplification Kit (Life Technologies, Carlsbad, CA, USA), cRNA was hybridized on HumanHT-12 v4 Expression BeadChips (Illumina, San Diego, CA, USA). Scanning was performed with an Illumina iScan microarray scanner, and data were analyzed with GeneSpring and Ingenuity software (Agilent Technologies, Santa Clara, CA, USA). Our microarray data have been deposited in the Gene Expression Omnibus database under accession number [GEO:GSE52660].
Reverse transcription and quantitative PCR
BT-474 cells were exposed to #hMADS-CM or hMADS-CM for the Additional file
2: Figure S6. Cells were harvested, and RNA was extracted using the RNeasy Mini Kit (QIAGEN). Reverse transcription (RT) was performed using random primers (Life Technologies). Quantitative PCR (qPCR) was performed with primers (QIAGEN) of the Additional file
2: Figure S6 using the LightCycler Nano Instrument (Roche Life Science, Indianapolis, IN, USA).
Western blot analysis
BT-474 cells were exposed to either #hMADS-CM or control medium for the Figure
5D. Proteins were extracted in radioimmunoprecipitation assay buffer and subjected to SDS-PAGE and immunoblot analysis. The antibodies used were anti-phospho-Akt and anti-Akt (Cell Signaling Technology, Beverly, MA, USA) and anti-tubulin (Sigma-Aldrich).
In vivo studies
All animal procedures were performed in accordance with European Union directive 86/609/EEC. Experiments were performed under individual permit and in animal care facilities accredited by the French Ministry of Agriculture. The study was approved by the local animal ethics committee (Université Claude Bernard Lyon I, protocol number BH-2012-40). The study was conducted using severe combined immunodeficiency (SCID) mice, with four used per group. Each mouse was given a subcutaneous injection of 1 ml of abdominal adipose tissue obtained from patients undergoing plastic surgery to form a lipoma. After 1 week, BT-474 tumors were grafted subcutaneously in contact with the lipoma. Treatments were initiated when the tumor volume was 100 mm3, with intraperitoneal administration of antibodies (rituximab 30 mg/kg or trastuzumab 25 mg/kg, twice per week for 3 weeks). Tumor growth was directly measured using a caliper, based on the difference of consistencies between the lipoma and the tumor.
Statistical analysis
All experiments were performed at least three times. Mean ± SD values of representative experiments are shown. Statistical significance was evaluated using paired Student’s t-tests on the means of at least three independent in vitro experiments. Unpaired Student’s t-tests were used for in vivo experiments. P-values <0.05 were deemed significant.
Discussion
Adipocytes constitute the most abundant cell type in the adipose tissue proximal to breast cancer cells and actively participate in tumor progression through the paracrine secretion of various adipocytokines [
33]. Recent studies have suggested that adipocytes promote cancer resistance to chemotherapy and radiotherapy [
17,
18]. However, the role of these cells in resistance to targeted therapy remains unknown. In this study, using an original coculture system, we investigated the implication of adipocytes and preadipocytes in trastuzumab-mediated ADCC in HER2-overexpressing breast cancer cells.
We demonstrate for the first time that adipocytes and preadipocytes inhibit trastuzumab-mediated tumor lysis by NK cells
in vitro and that adipose tissue inhibits the antitumor effect of trastuzumab
in vivo. This highlights the importance of adipose tissue in the resistance of cancer to targeted therapy using monoclonal antibodies. A recent study by Crozier
et al. showed that increased body mass index (BMI) is associated with shorter disease-free survival in HER2-overexpressing breast cancer patients, although trastuzumab improved clinical outcome, regardless of BMI [
34]. However, BMI is only a conventional indicator of the body fat mass and does not reflect the paracrine effect of the local adipose tissue in the microenvironment. Indeed, we observed a reduced antitumor effect of trastuzumab in mice in which the tumor was in direct contact with the lipoma, but not in mice in which the tumor was distantly localized from the lipoma (data not shown). Additionally, we observed an enhancement of the inhibition of ADCC by adipose cells under hypoxic conditions. Because hypoxia is one of the hallmarks of the tumor microenvironment [
29] and has also been associated with obesity [
35], this latter result underlines the potential clinical relevance of our findings on the impact of adipose tissue on targeted therapy for breast cancer. Furthermore, because adipocytes are hypertrophic with altered functions in obesity [
36,
37], a comparative study between adipocytes from lean and obese individuals could provide better understanding on the link between obesity and cancer resistance to therapies.
Several studies have pointed out the alteration of NK phenotype or NK functions by the tumor microenvironment [
38,
39]. Moreover, NK cells have been shown to express receptors for different adipokines, and their cytotoxicity has been found to be modulated by factors such as leptin and adiponectin [
40,
41]. Here, using NK-92-CD16 cells, we observed a modification of the secretion of IFN-γ, but not any modification of NK cell markers or NK cell cytotoxicity. These results seem to be in contrast to the work by DelaRosa
et al., who showed that human ASCs impaired NK cell cytotoxicity and NK cell markers, in particular decreased CD16 expression [
42]. However, in DelaRosa’s studies, the NK cells were continuously coincubated with ASCs for 72 hours, whereas in our studies, the incubation times were much shorter (4 hours or 18 hours). Moreover, the use of the NK-92 cell line may be not representative of the situation
in vivo. Therefore, a possible alteration of NK cells by adipose cells cannot be excluded in our studies. We also found that inhibition of ADCC by adipose tissue was not due to titration or degradation of the therapeutic antibody itself.
Our results suggest that adipose-derived factors reduce the sensitivity of HER2+ breast tumor cells to trastuzumab-mediated ADCC. This is supported by the facts that the ability of adipose cells to inhibit ADCC was variable, depending on the breast tumor cell lines studied, and that BT-474 cells preincubated overnight in #hMADS-CM showed decreased sensitivity to ADCC. We show that exposure of BT-474 cells to #hMADS-CM rapidly upregulated the expression of several genes involved in cell survival. However, the phenotypic alterations of tumor cells induced by exposure to adipocyte-conditioned medium are likely to be complex because downregulation of selected target genes by siRNA did not reverse the adipocyte-induced inhibition of ADCC. Importantly, the protective effect of adipocytes also seemed to apply to other targeted therapies because we observed a protection of BT-474 cells from T-DM1 cytotoxicity in the presence of #hMADS-CM. Likewise, it would be interesting to investigate the impact of adipocytes on the resistance of cancer cells to the adaptive immune system, notably resistance to cytotoxic T cell-mediated cytotoxicity. Further studies are required to better understand the roles of adipose tissue in cancer resistance to therapies.
In an attempt to identify the adipose secreted factors involved in inhibition of ADCC, we tested the ability of known adipocytokines to inhibit ADCC under our experimental conditions. Neither leptin, adiponectin, vaspin, IL-6, IGF-1, autotaxin, TNF-α, nor transforming growth factor β was found to reproduce the inhibition observed with the CM (Additional file
12: Table S3). As the secretomes of adipocytes and preadipocytes are extremely diverse [
43], a number of other adipose derived factors could be tested. We hypothesize that the inhibition of ADCC could be mediated by multiple factors that act at the same time on cancer cells.
In a previous study, Dirat
et al. [
44] showed that there is crosstalk between adipocytes and tumor cells and that adipocytes are modified by tumor cells to acquire a typical phenotype, named
cancer-associated adipocytes, that further enhances tumor cell invasion. In our short-duration coculture, we did not observe any delipidation of adipocytes by tumor cells (data not shown). However, we speculate that the inhibitory effect of ADCC observed with naive adipocytes could be further enhanced by cancer-associated adipocytes.
An important finding in our studies is that abdominal adipose tissue from plastic surgery could inhibit the antitumor effect of trastuzumab
in vivo. It is worth noting that the biology of adipose tissue can differ substantially, depending on its localization in the human body [
45], suggesting that further studies on mammary adipose tissue obtained from mammoplasties could provide more information on the impact of adipose tissue in the breast. Likewise, the impact of adipose tissue on the efficacy of other therapies also requires further investigation. Nonetheless, our data, together with those of others [
46,
47], raise the question of the safety of lipotransfer in patients undergoing breast reconstruction.
Competing interests
CD is a recipient of research grants from Roche. The other authors declare that they have no competing interests.
Authors’ contributions
MND participated in the conception and design of the study, performed the experiments, analyzed the results and drafted the manuscript. AC, ELM and DM performed the experiments, interpreted data, prepared the figures and were involved in drafting the manuscript. KC analyzed the microarray results and contributed to critical revision of the manuscript. SVW and BC generated the NK-92-CD16 cell line, participated in drafting the manuscript and provided critical comments on the manuscript. CD participated in the conception and design of the study and drafted the manuscript. All authors read and approved of the final manuscript.