Introduction
Rapid progress in genomic analysis and sequencing technology has opened doors to a new era of precision treatment, and targeted therapy has become one of the most important treatment regimens for cancer. Gastric cancer (GC) ranks third among the leading causes of cancer deaths worldwide; however, targeted therapies for GC are very limited [
1]. Although clinical trials for GC treatment have continued to emerge, most of them have failed [
2]. Among the molecular targets studied, human epidermal growth factor receptor 2 (HER2) is the most widely used, and its clinical significance has been clearly described. According to reports, about ~ 12% to ~ 20% of GC patients are HER2-positive, with rates in the adenocarcinoma of esophagogastric junction (AEG) as high as 32.2%, indicating the requirement for an effective treatment strategy for HER2-positive GC [
3,
4].
Trastuzumab is considered a first-line treatment for HER2-positive advanced gastric cancer (AGC); however, the objective response rate (ORR) is reportedly only 47% [
5]. Moreover, trastuzumab cardiotoxicity remains an issue, and trastuzumab resistance is an unavoidable problem, given the lack of a standard treatment for trastuzumab-resistant HER2-positive AGC. Previous studies show that a critical drug-resistance mechanism of trastuzumab is the compensatory signal transduction of other HER receptors [
6,
7]. Therefore, this suggests that the pan-HER inhibitor pyrotinib might be an effective treatment strategy for HER2-positive AGC.
Pyrotinib is a novel oral, potent, and irreversible pan-HER tyrosine kinase inhibitor (TKI), which inhibits the epidermal growth factor receptor (EGFR)/HER1, HER2 and HER4. Preclinical studies have revealed the excellent antitumor activity of pyrotinib both in vitro and in vivo in HER2-positive advanced solid tumors [
8,
9]. In a phase II randomized controlled trial (RCT) of pyrotinib in HER2-positive advanced breast cancer patients, a combination of pyrotinib and capecitabine significantly increased the ORR (78.5% vs. 57.1%) and extended progression-free survival (PFS) (18.1 vs. 7.0 months) as compared with combined lapatinib and capecitabine therapy. Moreover, the patients tolerated this treatment regimen well and benefitted from pyrotinib, irrespective of prior treatment with trastuzumab [
10]. This trial motivated the acceptance of pyrotinib by the China Food and Drug Administration (CFDA) for advanced breast cancer treatment, and based on its remarkable antitumor efficacy in HER2-positive breast cancer patients, we evaluated its application for the treatment of HER2-positive GC in the present study.
Currently, a phase I trial of pyrotinib (NCT03480256) is underway in HER2-positive AGC patients [
11]. Although no data have been announced, there have been previous clinical applications of pyrotinib in patients with HER2-positive AGC. Pyrotinib was reportedly effective in a HER2-positive AGC patient demonstrating resistance to trastuzumab and lapatinib [
12], underscoring pyrotinib efficacy in patients showing trastuzumab resistance. Because GC is highly heterogeneous, the efficacy of pyrotinib alone is limited in HER2-positive AGC patients; therefore, combining pyrotinib with other targeted drugs might improve its utility.
Apatinib is a small-molecule TKI that selectively inhibits vascular endothelial growth factor receptor 2 (VEGFR2), and is also effective against Ret, c-kit, and c-src [
13]. Apatinib reportedly exhibits a wide range of antitumor activity in advanced solid tumors [
14,
15], and to date, it is the first small-molecule inhibitor approved for AGC treatment. Although no clinical trials have reported apatinib efficacy in HER2-positive AGC, studies reported partial response in a trastuzumab-resistant HER2-positive AGC patient that subsequently experienced 8 months of PFS following apatinib treatment [
12]. These data suggest apatinib as a potential therapeutic option for HER2-positive AGC and trastuzumab resistance.
Both pyrotinib and apatinib are currently used independently for GC treatment; however, detailed studies on their combinatorial use for treating HER2-positive AGC have not been found. Therefore, we hypothesized that a combination of the two inhibitors would show a synergistic effect in HER2-positive AGC. Additionally, because drug resistance is a common limitation of targeted therapies, and resistance to anti-HER2 therapies has already been reported in HER2-positive AGC patients, understanding the mechanisms behind acquired pyrotinib resistance and identifying alternate treatment strategies are of great importance. Therefore, this study confirmed the role of combined application of pyrotinib and apatinib for the treatment of HER2-positive GC using in vitro and in vivo models. Furthermore, we established a pyrotinib-resistant cell line to examine the underlying mechanisms of acquired pyrotinib resistance, thereby laying the foundation for clinical research in HER2-positive AGC patients.
Methods
Cell lines and reagents
Human GC cell lines (NCI-N87, SGC7901, MGC-803, and MKN45) were purchased from the Cell Bank of Chinese Academy of Sciences (Shanghai, China). Human HER2-positive GC SNU216 cells were a generous gift from Professor Ruihua Xu, of Sun Yat-sen University Cancer Center (Guangzhou, China). NCI-N87-AR is a pyrotinib-resistant cell line obtained by continuous exposure of NCI-N87 cells to pyrotinib. All cells were cultured in RPMI-1640 (Hyclone, Provo, UT) containing 10% fetal bovine serum (FBS; Gibco, MD, USA) in an incubator with 5% CO2 and proper humidity, at 37 ℃. Pyrotinib (SHR1258) and apatinib (YN968D1) were obtained from Hengrui Medicine Co., Ltd (Lianyungang, China). Reagents were dissolved in dimethyl sulfoxide (DMSO) and stored at − 20 ℃. Recombinant human SCF was purchased from MedChemExpress LLC. (New Jersey, USA) and maintained at − 20 ℃ until further use.
Cell viability assay
Cells (4000–8000 cells/100μL per well) were seeded in 96-well plates and incubated at 37 °C overnight. After 72 h of treatments, the Cell Counting Kit (CCK)-8 reagent (Promotor, Wuhan, China) was added at a concentration of 10%, and the absorbance of the samples was measured at 450 nm using a microplate reader (BioTek, VT, USA). The experiment was performed in triplicate and data were representative of three separate experiments.
Cells were cultured in 6-well plates at a density of 800–1000 cells/well. The culture medium was renewed to normal after 72 h of different treatments and incubated for 2 weeks. The colonies were fixed and stained with 0.1% Crystal Violet, and colony (> 50 cells/colony) counts of the samples were then compared to that of the control group. The experiments were performed in triplicate and data were representative of three separate experiments.
Transwell assay
Cells (5 × 104) suspended in 200 µL serum-free medium were transferred into the upper chamber of a 24-well insert containing a membrane with an 8-μm pore size (Corning, New York, USA), and 500 μL medium containing 20% FBS was added to the bottom chamber. After incubation for 72 h, the cells that crossed the membrane were fixed and stained with 0.1% Crystal Violet for 30 min. For each sample, 10 fields of view were counted and the average was calculated (100 ×). The final data were obtained from three independent experiments.
Cell apoptosis analysis by flow cytometry
5 × 10
5 cells were obtained, washed twice with PBS, and re-suspended in 200 μL of 1 × binding buffer (BD Biosciences, NJ, USA), stained with 5 µL of FITC Annexin V, and incubated in the dark for 15 min, at 37 ℃. 5 µL of propidium iodide (PI) and 300 μL of 1 × binding buffer were then added 5 min prior to testing in the dark, at room temperature. Samples were measured by flow cytometry on an LSRFortessa cell analyzer (BD Biosciences), and the results were analyzed using FlowJo software (v.10.0; TreeStar, CA, USA). The final data were obtained from three independent experiments (Table
1).
Table 1
Synergism analysis in NCI-N87 xenograft models
NCI-N87 | 1.5–150 | 31.19 | 0.2–20 | 3.95 | 1.5/0.2–150/20 | 4.54 | 0.26 (0.48) |
RNA extraction and quantitative real-time PCR (qRT-PCR)
Total RNA was extracted using RNAiso Plus (Takara, Dalian, China), and reverse-transcribed to cDNA with the RevertAid first-strand cDNA synthesis kit (Thermo Fisher) according to a standard protocol. Real-Time qPCR was performed using the 7900HT Fast real-time PCR system (Thermo Fisher). GAPDH and β-actin were used as control and the 2−ΔΔCt method was used to quantify the relative mRNA expression of the genes. The sequences of primers are enlisted in the Supplementary Table 2. The experiments were performed in triplicate and the data were obtained from three separate experiments.
RNA sequencing and data analysis
RNA (1 μg) with an RNA Integrity number above 6.5 was used for following library preparation. Libraries with different indices were multiplexed and loaded onto an Illumina HiSeq instrument (Illumina, CA, USA) according to manufacturer’s instructions. The sequences were processed and analyzed by GENEWIZ (NJ, USA). A P value < 0.05 was used as the cut-off criterion.
Protein extraction and western blot analysis
Total protein was extracted with a radioimmunoprecipitation assay (RIPA) buffer (Beyotime, Shanghai, China) supplemented with phenylmethylsulphonyl fluoride (PMSF) and phosphatase inhibitors (Servicebio, Wuhan, China). Protein quantification was then performed using the BCA reagent (Beyotime), following the manufacturer's instructions. The extracted proteins were loaded into 10% SDS-PAGE for separation, and transferred to a 0.45 μm polyvinylidene fluoride (PVDF) membrane (Millipore, Massachusetts, USA). Primary antibodies were added, and the membrane was incubated at 4 ℃ overnight. Subsequently, the membrane was incubated with secondary antibodies (Promotor) at room temperature for 1 h. Post incubation, the SuperSignal West Pico Chemiluminescent Substrate (Thermo Fisher) was added. The immuno-reactive bands obtained were analyzed using the G: BOX Chemi X system (Syngene, Cambridge, UK). The primary antibodies used are enlisted in the Supplementary Table 3.
Xenograft models
Female nude mice (4–6-weeks old; BALB/c nu-nu) were purchased from the Hunan SJA Laboratory Animal Co., Ltd. (Changsha, China), and raised in a specific pathogen-free laboratory. Cells (5 × 10
6 cells/100 μL) were injected subcutaneously into the left posterior side of each mouse, and mice were randomized into different groups (
n = 6) when the tumors reached about ~ 100 mm
3. The size of the tumor and the weight of the nude mice were measured every 3 days, and the tumor volume was calculated as follows: [(short diameter)
2 × (long diameter)]/2. Mice were sacrificed after 21 days. Tumor samples were collected, photographed, and stained. The fixed dose ratio of apatinib/pyrotinib was determined from the median effective dose (ED
50) of the two monotherapies to prevent bias. The combination index (CI) was calculated to determine the combined effect, with CI < 1, CI > 1, and 1 used to defined synergism, antagonism, and additivity, respectively. To determine the statistical relevance of CI < 1 [i.e., ln(CI) < 0], the variance of ln(CI) was calculated according to a previous described method [
16]. All institutional and national guidelines for the care and use of laboratory animals were followed.
Immunohistochemistry (IHC)
Immunohistochemistry was performed following the manufacturer’s instructions. In brief, tissue samples from xenograft models (n = 6 per group) were formalin-fixed overnight at 25 ℃, prior to embedding in paraffin. All tissue sections were obtained as slices from the paraffin blocks and the slices were incubated with primary antibodies against SCF (1:200, ab52603, Abcam, Cambridge, UK), Ki67 (1:200, #9027, Cell Signaling Technology, MA, USA), and CD31(1:2000, #3528, Cell Signaling Technology, MA, USA). The TdT-mediated dUTP nick-end labeling (TUNEL) assays were performed using a TUNEL in situ cell death detection kit (Sigma-Aldrich, MO, USA), according to manufacturer’s instructions. The results were then evaluated by two professional pathologists independently. The assays were performed by Biossci Biotechnology Co., Ltd. (Wuhan, China).
Statistics
Chart generation and statistical calculations were performed using the GraphPad Prism software (v8.0; CA, USA), and SPSS (v19.0; NY, USA) was used for statistical analysis. All descriptive statistics were presented as the mean ± standard deviation (SD). Statistical comparisons between two experimental groups were performed using Student's t test. If multiple groups were compared, the one-way analysis of variance (ANOVA) was performed first. The least significant difference t test was applied if the overall difference was statistically significant. The main objective of statistical analysis in isobologram studies was to generate an upper confidence limit for the CI to determine whether the observed synergy was statistically relevant (CI < 1). A P < 0.05 was considered statistically significant for all tests.
Discussion
Currently, in precision medicine, HER2-targeted therapy is widely used in the treatment of a variety of tumors. However, tumor heterogeneity and the occurrence of primary or acquired drug resistance limit the efficacy of anti-HER2 therapy alone. Therefore, a commonly accepted treatment strategy involves combining anti-HER2 therapy with other targeted drugs, resulting in enhanced therapeutic benefits due to synergy, reduced treatment-related toxicity due to lower drug doses, and decreased or no drug resistance.
The interaction between drugs is usually confirmed in vitro by isobologram analysis [
19]. Since we were uncertain whether interaction of pyrotinib with apatinib in this study would be affected by tumor microenvironments, we performed in vivo analysis to allow a clinically relevant assessment. Therefore, we refer to a method derived from isobologram analysis, and use the CI to determine the presence of synergy in combined pyrotinib and apatinib treatment in vivo [
20]. Our analysis revealed a CI < 1, which indicated synergy and was consistent with our hypothesis.
We initially considered the anti-vascular effect of apatinib during our investigation of the synergistic mechanism. Co-culture of apatinib-treated HUVEC cells with pyrotinib-treated HER2-positive GC cells revealed no significant difference in cell survival, proliferation, or malignant properties in the GC cells relative to controls. Because apatinib is a multi-target TKI in addition to its targeting VEGFR2, we speculated that apatinib and pyrotinib may act on tumor cells together.
Researchers have reported that the PI3K/AKT and MAPK pathways are the most important downstream signaling pathways of HER2 receptor [
21]. Our data showed that HER2-positive GC cells exposed to combined pyrotinib and apatinib treatment did not exhibit significant decreases in phosphorylated levels of HER2 (p-HER2) and p-VEGFR2 relative to those observed following monotherapies, whereas p-AKT, p-mTOR and p-ERK were significantly reduced following combined treatment (Fig.
1l). Therefore, we speculated that apatinib blocked PI3K/AKT and MAPK pathways through other pathways unrelated to HER2 and VEGFR2 levels.
To investigate the mechanisms associated with pyrotinib resistance and identify potential treatment strategies, we compared the differences in pyrotinib-resistant cells and their parent cells.
KITLG encodes a ligand for the tyrosine kinase c-kit, commonly known as stem cell factor (SCF) [
22‐
25]. SCF binding to its receptor (c-kit) promotes phosphorylation of the PI3K regulatory subunit p85α encoded by
PIK3R1 to subsequently activate the protein kinase B (i.e., AKT1). Additionally, SCF is involved in mediating MAPK signaling. The primary function of SCF in hematopoietic cells is to enhance the effect of other growth factors, and thereby induce cell proliferation [
23,
26,
27]. It also plays a similar role in promoting cell growth in solid tumors. Previous reports indicate that an SCF/c-kit autocrine loop increases cell growth via EGF family ligands by enhancing signal transduction in the PI3K and ERK pathways in breast cancer [
28,
29]. Other researchers found that the activation of the SCF/c-kit pathway promoted the property of cell proliferation and invasion in colorectal and pancreatic cancers via the PI3K/AKT and/or MAPK pathways [
30,
31]. In the present study, our data were consistent with these findings and demonstrated that up-regulation of SCF levels was associated with acquired pyrotinib resistance. Activation of PI3K/AKT and MAPK signaling via the SCF/c-kit interaction induced the proliferation and survival of pyrotinib-resistant cells.
Previous studies indicate that activation of alternate pathways represents an important resistance mechanism against anti-HER2 drugs [
7,
32,
33]. The crosstalk between the EGFR/HER2 and SCF/c-kit pathways occurs at multiple levels, with PI3K/AKT and ERK representing crucial downstream signaling nodes. Because SCF activates PI3K/AKT and MAPK signaling through its receptor c-kit, this suggests that combining pyrotinib with c-kit inhibitors might effectively address the observed drug resistance.
The c-kit protein, which is the SCF receptor, is a member of the type III receptor tyrosine kinase family and stimulates continuous proliferation and loss of anti-apoptotic signals in tumor cells through various pathways [
23]. Imatinib, a classical c-kit inhibitor, was found to be effective for combating acquired pyrotinib resistance both in NCI-N87-AR cells and xenografts. These data indicate that the SCF/c-kit signaling do play an important role in overcoming acquired pyrotinib resistance. In addition to imatinib, apatinib, which has been proven to be effective in AGC, can also effectively inhibit c-kit phosphorylation. In our study, apatinib blocked the downstream pathways of SCF/c-kit, and therefore re-sensitized pyrotinib-resistant cells to pyrotinib both in vitro and in vivo. Since apatinib also has anti-angiogenic effects, combining it with pyrotinib seems to be a better choice than that of imatinib, which could be verified by the results of in vivo immunohistochemical staining.
Interestingly, we found that the inhibitory effect of apatinib on c-kit and the downstream pathways was also observed in NCI-N87 and SNU216 cells (Supplementary Fig.
2, Fig.
1l), indicating that apatinib might show synergy with pyrotinib by targeting c-kit and inhibiting the PI3K/AKT and MAPK signaling.
The complexity of the mechanisms associated with anti-HER2 resistance suggests the existence of other mechanisms related to acquired pyrotinib resistance [
34‐
36]. A previous study reported that application of exosomes produced by HER2-positive GC cells treated with pyrotinib promotes HUVEC proliferation, invasion, and translocation, whereas administration of apatinib counteracted these effects [
37]. Although this suggested a potential mechanism of pyrotinib resistance, the hypothesis requires further confirmation both in vitro and in vivo.
This study has limitations. Although our findings are compelling, different pyrotinib-resistant cell lines need to be established, and large-scale clinical studies need to be conducted for confirmation of the results and further elucidation of the associated mechanisms. Furthermore, clinical trials are required to establish the safety and efficacy of this drug combination in vivo.
In conclusion, our study revealed that apatinib exhibited synergistic antitumor effects with pyrotinib and reversed acquired pyrotinib resistance in HER2-positive GC by targeting the SCF/c-kit/PI3K/AKT and SCF/c-kit/MAPK signaling pathways (Fig.
6g). Additionally, our findings provided evidence for the safety of the combined therapy in vivo using an animal model. These results suggest the efficacy of combined application of pyrotinib and apatinib in HER2-positive AGC patients as a novel treatment strategy that can also be applied for patients with acquired pyrotinib resistance characterized by elevated SCF expression.
Publisher's Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.