Skip to main content
NCBI home page
Springer logoLink to Springer
. 2026 Jan 31;31(1):6. doi: 10.1007/s10911-026-09600-3

Synergistic Anticancer Effects of Apatinib and PD-L1 Inhibition in Breast Cancer

Danyang Han 1,✉, Juanjuan Xu 1, Cairu Guo 1
PMCID: PMC12948880  PMID: 41619044

Abstract

Studies in breast cancer have demonstrated that apatinib exhibits both antiangiogenic and antitumor effects, while PD-L1 inhibitors have similarly shown meaningful clinical benefit. Building upon these observations, this study evaluated the potential synergistic antitumor effects of combining apatinib with a PD-L1 inhibitor and examined the mechanistic basis for their interaction in breast cancer. Notably, we found that this regimen could significantly suppress the proliferation, migration, and invasion of MCF-7 and MDA-MB-231 cells, and promote cell apoptosis. In addition, the levels of p-ERK, NF-κB, and Slug were markedly reduced in vitro. Collectively, these findings support the potential clinical utility of combining apatinib and PD-L1 inhibition, as evidenced by consistent in vitro and in vivo synergy.

Keywords: PD-L1 inhibitor, Apatinib, Breast cancer, Apoptosis

Introduction

Breast cancer (BC) is the most common malignancy among women worldwide and accounts for approximately 15% of all female cancer-related deaths [1]. Apatinib, a novel tyrosine kinase inhibitor, suppresses tumor growth by selectively targeting and inhibiting vascular endothelial growth factor (VEGF) receptor 2, thereby blocking the VEGF/VEGFR2 signaling pathway [2]. Extensive research has demonstrated the substantial antitumor efficacy of apatinib across multiple cancer types, including breast cancer [3–5], pointing to its potent inhibitory effects on BC cells. Programmed death-ligand 1 (PD-L1), also known as B7 homology 1 (B7-H1) or cluster of differentiation 274 (CD274), is a 40-kDa type I transmembrane protein belonging to the B7 family. Notably, PD-L1 expression is markedly upregulated not only on tumor cells but also on macrophages and dendritic cells. This increased expression enables tumor cells to evade immune surveillance and promotes immune escape by engaging the programmed death 1 (PD-1) receptor on T cells, thereby suppressing T-cell immune function through the PD-1/PD-L1 axis [6]. Interestingly, PD-L1 inhibitors have been shown to exert significant antitumor effects even in the absence of T cells. In recent years, multiple PD-1/PD-L1 targeting antibodies have received Food and Drug Administration (FDA) approval for cancer treatment, either as monotherapies or in combination with chemotherapy [7]. Nevertheless, previous studies have shown that PD-L1 blockade alone may, in some settings, lead to tumor progression and unfavorable clinical outcomes [8, 9]. Reports indicate that PD-1/PD-L1 monotherapy reduces cancer cell death by only 20%-40%, depending on the tumor type [10–12]. Consequently, optimizing immunotherapy strategies has re-emerged as a major focus in cancer research.

Herein, the antitumor effects of apatinib and a PD-L1 inhibitor were evaluated both individually and in combination in vitro and in vivo. Moreover, the mechanisms underlying their synergistic activity were also explored to help optimize their combined application in appropriate cancer patients.

Materials & Methods

Cell Culture and Chemicals

MDA-MB-231 and MCF-7 breast cancer cell lines were obtained from the China Center for Type Culture Collection (CCTCC). Cells were cultured in DMEM supplemented with 10% fetal bovine serum at 37 °C in a humidified atmosphere containing 5% CO₂, and all cell lines were confirmed to be mycoplasma-free. Apatinib and PD-L1 inhibitor (SHR-1316 ,Adebrelimab) are provided by Hengrui Medicine Co. Ltd.

Viability Assay

Cell viability was assessed by MTT assay. Briefly, MCF-7 and MDA-MB-231 cells were seeded in 96-well plates at a density of 4,000 cells/well and subsequently exposed to DMSO (control), serially diluted apatinib (starting from 200 µg/mL), serially diluted PD-L1 inhibitor (starting from 10 mg/mL), or their combinations for 36–48 h. Following treatment, 10 µL of MTT solution was added to each well, and cells were incubated for an additional 4 h. After aspirating the medium, the resulting formazan crystals were dissolved in 150 µL of DMSO. Absorbance at 570 nm was measured using a microplate reader (Bio-Tek, Norcross, GA, U.S.A). The IC50 value for each drug was calculated using a non-linear regression model in SPSS software. Subsequent combination experiments employed concentrations based on these IC50 values. The synergistic effect of the drug combination was analyzed by calculating the Combination Index (CI) using CompuSyn (ComboSyn Inc.) software. Synergy was defined as a combination index (CI) value < 1.0, antagonism as CI > 1.0, and additivity as CI = 1.0. All experiments were performed in triplicate, and data were plotted using GraphPad Prism.

Wound Healing and Transwell Assays

Cell migration was evaluated using a scratch wound healing assay. Briefly, cells were seeded in six-well plates at a density of 1 × 10⁶ cells per well and cultured overnight to reach approximately 90% confluence. A uniform scratch was then created by a sterile pipette tip. After washing to remove detached cells, the cells were incubated in fresh medium containing the designated treatments (DMSO, apatinib, PD-L1 inhibitor, or their combination). The scratch area was photographed at 0 and 24 h post-scratching under a microscope, and the relative wound closure was quantified using ImageJ software (NIH, Bethesda, MA, U.S.A.).

Cell invasion was assessed using a Matrigel-coated Transwell system. Cells were suspended in serum-free medium at a density of 1 × 106 cells/mL, and 200 µL of the suspension was added to the upper chambers (Corning Inc.,Coring, NY, USA) of a 24-well Transwell insert (8 μm pore size) pre-coated with Matrigel (Sigma-Aldrich, St. Louis, MO, USA). The lower chamber was filled with 600 µL of medium containing 10% fetal bovine serum as a chemoattractant. Following 24 h of incubation at 37 °C, non-invaded cells on the upper surface of the membrane were gently removed. The invaded cells on the lower surface were fixed with 4% paraformaldehyde, stained with 0.1% crystal violet, and counted in three randomly selected fields per well under a light microscope.

Cell Cycle and Apoptosis Analysis Using Flow Cytometry

Cell cycle distribution was analyzed by flow cytometry. Briefly, MCF‑7 and MDA‑MB‑231 cells were seeded in six‑well plates and cultured overnight. Cells were then treated with DMSO, apatinib, PD‑L1 inhibitor, or their combination for 24 h. After treatment, cells were harvested by trypsinization, washed with PBS, and fixed in 70% ice‑cold ethanol at ‑20 °C for at least 1 h. Fixed cells were washed with PBS, treated with RNase A (100 µg/mL) at 37 °C for 30 min, and stained with propidium iodide (PI, 50 µg/mL) at 4 °C for 30 min in the dark. DNA content was analyzed on a BD FACSCalibur flow cytometer, and the proportions of cells in G0/G1, S, and G2/M phases were determined using ModFit LT software.

Apoptosis was assessed using an Annexin V‑FITC/PI apoptosis detection kit (Solarbio, CA1020) according to the manufacturer’s instructions. After 24 h of drug treatment as described above, cells were collected, washed twice with cold PBS, and resuspended in 1× binding buffer. Cells were then incubated with Annexin V‑FITC and PI for 15 min at room temperature in the dark. The stained samples were analyzed immediately by flow cytometry. Quadrant analysis was performed to distinguish viable (Annexin V‑/PI‑), early apoptotic (Annexin V+/PI‑), late apoptotic (Annexin V+/PI+), and necrotic (Annexin V‑/PI+) cell populations.

Western Blotting

Following treatment with DMSO, apatinib, PD‑L1 inhibitor, or their combination for 24 h, MCF‑7 and MDA‑MB‑231 cells were harvested. Cytoplasmic and nuclear proteins were fractionated using a Nuclear and Cytoplasmic Protein Extraction Kit (Cowin Bio, CW0199) according to the manufacturer’s protocol. Protein concentrations were determined using a BCA assay (Solarbio, Beijing, China). Equal amounts of protein were separated by SDS‑PAGE and transferred onto PVDF membranes (Millipore, Massachusetts, U.S.A). The membranes were blocked with 5% non‑fat milk for 1 h at room temperature and then incubated overnight at 4 °C with the following primary antibodies purchased from Cell Signaling Technology (CST): anti‑ERK (CST‑4695, 1:1000), anti‑p‑ERK (CST‑4370, 1:1000), anti‑NF‑κB (CST‑8242, 1:1000), and anti‑SLUG (CST‑9585, 1:1000). Rabbit anti‑β‑actin antibody (Bioss, bsm‑52262R, 1:5000) was used as a loading control. After washing three times with TBST, the membranes were incubated with an HRP‑conjugated goat anti‑rabbit (Bioss, bs‑0295G, 1:5000) for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence (ECL) detection kit (Tanon, 180‑5001) and imaged with a Tanon 2500 chemiluminescence system. Band intensities were quantified using ImageJ software (NIH, Bethesda, MA, U.S.A).

In Vivo Experiments

BALB/c nude female mice were purchased from Changzhou Covens Experimental Animal Co. Ltd. Female nude mice (4–6 weeks old) were maintained under pathogen-free conditions with controlled temperature (20–26 °C), humidity (40–60%), and a 12-hour light/dark cycle. MDA-MB-231 cells (1 × 10⁶) were subcutaneously injected into the abdominal region in a 0.2 mL suspension. Following injection, mice were monitored regularly, and once tumors were established, the mice were randomly assigned to four experimental groups (n = 6 per group). In the treatment regimens, apatinib was administered orally at 50 mg/kg daily, while the PD-L1 inhibitor (Adebrelimab) was given intravenously at 10 mg/kg on days 3, 6, and 10. Based on the results of the pre-experiment and the existing pharmacokinetic literature [12], this spaced dosing regimen was designed to maintain the effective drug concentration while reducing potential cumulative toxicity. Specifically, starting the administration on the third day is to allow the tumor to grow to an appropriate volume, facilitating subsequent experiments. In the combination group, both treatments were administered according to these schedules. Tumor volume was measured twice weekly and calculated using the formula: Tumor volume = (length × width × 2)/2, with assessments based on tumor size and body weight. All experimental procedures were conducted in accordance with approved ethical guidelines (approval number: LWLL-2024-11-05-01).

Statistical Analysis

All experiments were performed in triplicate. Statistical analyses were conducted using IBM SPSS version 25.0 (SPSS, Chicago, IL, USA). Data are presented as mean ± standard deviation (SD). Differences between experimental and control groups were evaluated using one-way ANOVA, with P < 0.05 considered statistically significant.

Results

Effects of Apatinib and PD-LI Inbibitor on Tbe Proliferation of MCF‑7 and MDA‑MB‑231 Cells

The antiproliferative effects of apatinib and the PD‑L1 inhibitor on breast cancer cells were evaluated using the MTT assay. As shown in Fig. 1A, both apatinib and the PD‑L1 inhibitor reduced cell viability in a dose‑dependent manner. The half‑maximal inhibitory concentration (IC50) were 15 µg/mL for apatinib, and 2 mg/mL or 3 mg/mL for the PD‑L1 inhibitor in MCF‑7 and MDA‑MB‑231 cells, respectively. Based on these IC50 values, the drugs were tested in combination under three different sequential regimens: apatinib followed by PD‑L1 inhibitor, PD‑L1 inhibitor followed by apatinib, and simultaneous treatment. The combination treatments resulted in significantly enhanced growth inhibition compared to either drug alone (Fig. 1B). Notably, the sequential treatment of apatinib followed by the PD‑L1 inhibitor exhibited the strongest antiproliferative effect among the three regimens. Based on this result, this schedule was selected for subsequent in vivo experiments. Synergy was quantitatively assessed using the Chou‑Talalay method, and the calculated combination index (CI) values was always below 1, confirming the synergistic interaction between apaptinib and PD-L1 inhibitor (Fig. 1C). Furthermore, dose-response curves generated from serial dilutions around the IC₅₀ of each drug (Fig. 1D, E) demonstrated a concentration-dependent effect. Quantitative analysis of these data (Fig. 1F) further supported the concentration‑dependent suppression of cell proliferation. Consistent results were obtained in MDA‑MB‑231 cells, as shown in Fig. 1G–L, which validated the findings in a second breast cancer cell line. Collectively, the antiproliferative effect of PD‑L1 inhibition on breast cancer cells is potentiated by co‑treatment with apatinib. The sequential administration of apatinib followed by the PD‑L1 inhibitor yields the most potent synergy, underscoring the therapeutic potential of this drug combination schedule.

Fig. 1.

Fig. 1

Open in a new tab

Apatinib and PD-L1 inhibitors synergistically suppress breast cancer cell growth. A–F Effects on MCF-7 cells. A Dose-response curves of MCF-7 cell viability following 48-h treatment with increasing concentrations of apatinib or PD-L1 inhibitor, as determined by MTT assay. B Cell viability under monotherapy, concurrent combination (A + P), or sequential treatment regimens. C Combination index (CI) plot generated by CompuSyn software, demonstrating synergy (CI < 1) between apatinib and the PD-L1 inhibitor. D, E Proliferation of MCF-7 cells treated with a concentration gradient of PD-L1 inhibitor (D) or apatinib (E) for 48 h. F Quantification of the data presented in panel (D). G–L Effects on MDA-MB-231 cells. G Dose-response curves of MDA-MB-231 cell viability after drug treatment. H Cell viability under the indicated treatment regimens. I CI plot confirming synergistic interaction in MDA-MB-231 cells. J, K Proliferation of MDA-MB-231 cells treated with a concentration gradient of (J) PD-L1 inhibitor or (K) apatinib. (L) Quantification of the data from panel (J)

Combined Apatinib and PD-L1 Inhibition Significantly Suppresses Breast Cancer Cell Invasion and Metastasis

The effects of apatinib and PD-L1 inhibitor on cell migration were evaluated using wound-healing assays in MCF-7 and MDA-MB-231 cells. In the wound healing assay, treatment with either apatinib or the PD‑L1 inhibitor significantly suppressed the migration of MCF‑7 cells in a concentration‑dependent manner. The relative wound closure rates were reduced to 16.65 ± 5.03% and 22.25 ± 2.49% by high‑concentration apatinib and PD‑L1 inhibitor, respectively, compared to the control (set as 100%, P < 0.01; Fig. 2A). The PD‑L1 inhibitor alone exhibited a weaker inhibitory effect than apatinib at equivalent concentrations. Notably, the combination treatment resulted in the most potent suppression, decreasing wound closure to 7.78 ± 3.27% (P < 0.001), indicating a synergistic anti‑migratory effect. A similar inhibitory trend was confirmed in MDA‑MB‑231 cells (Fig. 2B). The quantitative data summarized in Fig. 2C and D confirm the superior in vitro growth-inhibitory activity achieved by the synergistic drug combination in MCF-7 and MDA-MB-231 cell.

Fig. 2.

Fig. 2

Open in a new tab

Apatinib and PD-L1 inhibitor suppress breast cancer cell migration. Representative images and quantitative analysis from wound‑healing (migration) assays in (A, B) MCF‑7 and (C, D) MDA‑MB‑231 cells following treatment with apatinib, PD‑L1 inhibitor, or their combinatio. A-C Representative images of the scratch wound (0 and 24 h) and invaded cells. B, D Quantification of relative wound closure and the number of invaded cells, respectively. The histogram data represents the mean ± SD of three independent experiments. *p < 0.05 and **P < 0.01 **p < 0.001 compared with the control. PBS, Phosphate bufferd saline, A, Apatinib; P, PD‑L1 inhibitor

Consistent with the migration results, the Transwell assay demonstrated that both agents effectively attenuated cellular invasion. In MCF‑7 cells, the number of invading cells was significantly decreased in all treatment groups (P < 0.05) except the low‑concentration PD‑L1 inhibitor group, with the combination group showing the most profound reduction (66.67 ± 2.69%, P < 0.001; Fig. 3A). In MDA‑MB‑231 cells, significant inhibition was observed in the combination group (74.46 ± 3.57% reduction, P < 0.001) and the high‑concentration monotherapy groups (P < 0.05), but not in the low‑concentration groups (Fig. 3B). Consistent with this trend, the quantitative results in Fig. 3C and D verified the enhanced anti‑invasive efficacy of the synergistic combination in both cell lines.

Fig. 3.

Fig. 3

Open in a new tab

Apatinib and PD-L1 inhibitor inhibit breast cancer cell invasion. Transwell invasion assays showing the effect of drug treatment on MCF‑7 (A, B) and MDA‑MB‑231 (C, D) cells. Cells were treated for 24 h with PBS (control), apatinib (1.5 µg/ml or 15 µg/ml), or PD‑L1 inhibitor (0.2 mg/ml or 2 mg/ml for MCF‑7, 0.3 mg/ml or 3 mg/ml for MDA‑MB‑231). A-C Representative images of invaded cells. B, D Quantification of the number of invaded cells per field. Data are presented as mean ± SD from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001 compared with the control. PBS, Phosphate‑buffered saline; A, Apatinib; P, PD‑L1 inhibitor

The Combination of PD-L1 Inhibitor and Apatinib Differentially Induced Apoptosis, as Opposed to Cell Cycle Arrest

As shown in Fig. 4 (Fig. 4A, B), treatment with apatinib, PD‑L1 inhibitor, or their combination did not induce significant changes in the cell cycle distribution of MCF‑7 or MDA‑MB‑231 cells. This observation was further confirmed by quantitative assessment (Figs. 4C, D), which indicated no statistically significant difference in cell cycle progression between the treated groups and the control.

Fig. 4.

Fig. 4

Open in a new tab

Effects of apatinib and PD-L1 inhibitor on cell cycle. Cell cycle distribution was analyzed by flow cytometry following propidium iodide (PI) staining in MCF‑7 and MDA‑MB‑231 cells. A Cell cycle analysis of MCF-7 cells by PI staining and flow cytometry after treatment with PBS, apatinib (1.5–15 µg/ml), or PD-L1 inhibitor (0.2–2 µg/ml) for 24 h. B Quantification of the percentage of cells in each cell cycle phase (G0/G1, S, and G2/M) in MCF-7. C Cell cycle analysis of MDA-MB-231 cells by PI staining and flow cytometry following treatment with PBS, apatinib (1.5–15 µg/ml), or PD-L1 inhibitor (0.3–3 µg/ml) for 24 h. D Quantification of the percentage of cells in each cell cycle phase (G0/G1, S, and G2/M) in MDA-MB-231. PBS, Phosphate‑buffered saline; A, Apatinib; P, PD‑L1 inhibitor

Next, the effects of apatinib and PD-L1 inhibition on apoptosis were evaluated in MCF-7 and MDA-MB-231 cells using Annexin V-FITC/PI double staining. As quantified in Fig. 5A and B, all drug treatments significantly increased the total apoptosis rate compared to the control. In MCF‑7 cells, the total apoptotic population rose from 3.04 ± 0.61% (control) to 24.14 ± 0.81% with high‑dose apatinib, 31.55 ± 2.99% with high‑dose PD‑L1 inhibitor, and 44.90 ± 4.03% with the combination treatment (P < 0.05 for all vs. control). Notably, the combination regimen induced a synergistic increase in early apoptosis (Annexin V⁺/PI⁻), which reached 33.95 ± 2.55% in MCF‑7 cells. A similar trend was observed in MDA‑MB‑231 cells (Fig. 5C, D), where the combination treatment elevated the total apoptosis rate to 32.09 ± 4.42% (vs. 3.86 ± 1.02% in control, P < 0.01). In this cell line, the synergistic effect was primarily reflected in a marked increase in late apoptosis (Annexin V⁺/PI⁺), which accounted for 28.99 ± 3.27% of cells following combination therapy. Collectively, these results demonstrate that both apatinib and PD‑L1 inhibitor promote apoptosis in breast cancer cells in a dose‑dependent manner, and that their combination generates a significantly enhanced pro‑apoptotic effect, with distinct patterns of early and late apoptosis induction in MCF‑7 and MDA‑MB‑231 cells, respectively.

Fig. 5.

Fig. 5

Open in a new tab

Apatinib and PD-L1 inhibitor induce apoptosis in breast cancer cells. Apoptosis was assessed by Annexin V-FITC/PI double staining and flow cytometry in MCF‑7 and MDA‑MB‑231 cells. A Apoptosis of MCF-7 cells was assessed by Annexin V-FITC/PI double staining and flow cytometry after treatment with PBS, apatinib (1.5–15 µg/ml), or PD-L1 inhibitor (0.2–2 µg/ml). B Quantification of the total apoptotic cell populations (early + late apoptosis) in MCF-7. C Apoptosis of MDA-MB-231 cells was assessed by Annexin V-FITC/PI double staining and flow cytometry under the same conditions as panel A. D Quantification of the total apoptotic cell populations (early + late apoptosis) in MDA-MB-231. Data are presented as mean ± SD of three independent experiments. PBS, Phosphate‑buffered saline; A, Apatinib; P, PD‑L1 inhibitor

The Combination of Apatinib and the PD-L1 Inhibitor Converges To Inhibit the p-ERK/NF-κB/Slug Pathway

To elucidate the molecular mechanism underlying the synergistic antitumor effects, the activity of key signaling proteins was examined by Western blot. Following 48 h of combination treatment, the protein levels of phosphorylated ERK (p-ERK), NF-κB, and the epithelial-mesenchymal transition (EMT) regulator Slug were significantly downregulated in both MCF-7 and MDA-MB-231 cells compared to the control (Fig. 6A, B). In contrast, treatment with either apatinib or the PD-L1 inhibitor alone did not produce a statistically significant reduction in p-ERK or NF-κB levels in MCF-7 (Fig. 6C) or MDA-MB-231 cells (Fig. 6D). Notably, while monotherapy groups showed no significant effect (P > 0.05), the combination therapy consistently yielded significant inhibition (P < 0.05). These results indicate that the observed synergistic effect is associated with concurrent inhibition of the p-ERK and NF-κB pathways and the synergistic drug combination potently suppresses the p-ERK/NF-κB/Slug signaling axis. Based on these findings, a schematic model was proposed to illustrate how the co-targeting strategy collaboratively inhibits breast cancer cell proliferation, migration, and invasion (Fig. 6E).

Fig. 6.

Fig. 6

Open in a new tab

Molecular mechanisms underlying the effects of Apatinib and PD-L1 inhibitor in MCF-7 and MDA-MB-231 cells. MCF-7 and MDA-MB-231 cells were treated with PBS, apatinib (1.5–15 µg/ml), PD-L1 inhibitor (0.2–2 µg/ml for MCF-7; 0.3–3 µg/ml for MDA-MB-231), or their combination for 24 h. Western blot analysis was performed to assess the expression of ERK, p-ERK, NF-κB, and Slug in nuclear and cytosolic fractions of MCF-7 (A) and MDA-MB-231 (B) cells. C, D Quantification of Western blot results for MCF-7 (C) and MDA-MB-231 (D) cells. E Schematic representation of the inhibitory effects of apatinib and PD-L1 inhibition on breast cancer cell proliferation, migration, and invasion. Data are presented as mean ± SD (n = 3). *P < 0.05, **P < 0.01, and ***P < 0.001 compared the control group. PBS, Phosphate‑buffered saline; A, Apatinib; P, PD‑L1 inhibitor

Apatinib and PD-L1 Inhibitors Synergistically Inhibit Tumor Growth in Vivo

The antitumor efficacy of apatinib and the PD‑L1 inhibitor, alone or in combination, was further evaluated in a BALB/c nude mouse xenograft model. Animals in the combination group received daily apatinib, with PD‑L1 inhibitor administration initiated on day 3 (Fig. 7A). This intermittent dosing schedule (every 3 days for 3 cycles) is a validated model for evaluating PD-1/PD-L1 blockade [13]. Our specific timing was designed based on timing therapy, aiming to intervene during the early establishment of the tumor immune microenvironment. This optimized schedule, in synergy with the continuous administration of apatinib, maximized the observed therapeutic synergy without increasing toxicity. The results showed that the tumor growth of all treatment groups was significantly inhibited compared with the control group(Fig. 7B). While apatinib monotherapy induced a modest reduction in tumor volume, the combination treatment resulted in the most pronounced inhibition throughout the study period. By day 21, the mean tumor volume in the control group reached 874 ± 102 mm³, whereas in the combination group it was limited to 312 ± 45 mm³ (P < 0.01 vs. control; Fig. 7D). Correspondingly, tumor weight at the endpoint was lowest in the combination group (Fig. 7C), demonstrating superior tumor suppression compared to either monotherapy (P < 0.05). No significant loss in body weight was observed across treatment groups (Fig. 7F), indicating that the regimens were well tolerated. Collectively, these in vivo results confirm that the combination of apatinib and PD‑L1 inhibitor exerts significantly enhanced antitumor activity compared to single‑agent treatment, supporting its potential therapeutic utility for breast cancer intervention.

Fig. 7.

Fig. 7

Open in a new tab

Synergistic antitumor efficacy of apatinib combined with PD-L1 inhibitor in breast cancer xenograft models. In vivo anticancer effect of apatinib (A) and PD-L1 inhibitor (P) in breast cancer xenograft models. A Schematic of the treatment schedule in the xenograft model. Red arrows indicate oral administration of apatinib, and blue arrows indicate intravenous injection of PD-L1 monoclonal antibodies. B Representative images of tumors from nude mice randomly assigned to receive PBS, apatinib, PD-L1 inhibitor, or combination treatment (A + P) for 21 days. C Tumor weights were measured on day 21 post-treatment. D The tumor growth volume was recorded every three days throughout the treatment period. E Body weight changes of mice were measured every three days during the treatment period. Data are presented as mean ± SD of three independent experiments. *P < 0.05, **P < 0.01, and ***P < 0.001 compared the control group. PBS, Phosphate‑buffered saline; A, Apatinib; P, PD‑L1 inhibitor

Discussion

Breast cancer remains a leading cause of cancer-related mortality worldwide [14]. While immunotherapy targeting the PD-1/PD-L1 axis has shown promise in a subset of patients [15, 16], response rates are limited, driving the search for rational combinations. This study demonstrates that the combination of apatinib, a VEGFR2 tyrosine kinase inhibitor, with a PD-L1 inhibitor exerts superior antitumor activity against breast cancer cells both in vitro and in vivo. Notably, a sequential administration schedule of apatinib followed by the PD-L1 inhibitor proved most effective. Mechanistically, this synergy is associated with the cooperative suppression of the p-ERK/NF-κB/Slug signaling pathway, a key regulator of cell proliferation, survival, and epithelial-mesenchymal transition (EMT) [17–20].

Our findings align with the growing evidence supporting the combination of antiangiogenic agents and immune checkpoint inhibitors [21]. Previous studies indicate that VEGF-driven angiogenesis contributes to an immunosuppressive tumor microenvironment, and its inhibition can potentiate immunotherapy [22, 23]. For instance, Schmittnaegel et al. reported that antiangiogenic agents can enhance the efficacy of anti-PD-L1 therapy across multiple tumor types [24]. Previous studies also suggest that this combination promotes antitumor immunity by inducing the formation of high endothelial venules (HEVs) [25]. Research suggests that combining apatinib with PD-L1 monoclonal antibodies can enhance antitumor efficacy by modulating tumor vasculature and promoting immune-cell infiltration, as demonstrated in preclinical studies [26–28]. Consistent with this, our data show that the apatinib and PD-L1 inhibitor combination achieved significantly greater tumor suppression than either monotherapy, without exacerbating toxicity as indicated by stable body weight. This aligns with clinical observations from phase 1b studies of avelumab in metastatic breast cancer, supporting the relevance of this approach [29, 30].

The novelty of our work lies in the elucidation of a specific intracellular signaling axis—p-ERK/NF-κB/SLUG—as a convergent point for the synergistic action of this drug combination. While apatinib is known to inhibit VEGFR2 and downstream pathways like MAPK/ERK [17], and PD-L1 signaling has been linked to Ras/ERK activation, their combined effect on this specific pathway network in breast cancer was not fully characterized [31]. Our Western blot analyses revealed that only the combination treatment can lead to significant concurrent downregulation of p-ERK, NF-κB, and the EMT-transcription factor Slug. This effect is mediated through a pathway in which the inhibition of ERK and NF-κB signaling converges to suppress Slug expression, a key driver of metastasis, thereby attenuating cellular migratory and invasive capacities [32]. This finding extends the work of Zhao et al. who highlighted the role of ERK inhibition in suppressing NF-κB-driven metastasis, by positioning Slug as a key downstream effector in the context of this particular drug synergy [19].

Furthermore, we optimized the treatment sequence. The superior efficacy of administering apatinib prior to the PD-L1 inhibitor can be rationalized by the “vascular normalization” hypothesis [33]. Apatinib may first remodel the tumor vasculature, improving perfusion and immune cell delivery, thereby creating a more favorable microenvironment for the subsequently administered immunotherapy. This conceptual demonstrating that antiangiogenics can enhance the efficacy of immune checkpoint blockade.

Despite promising results, our study has limitations. The research was conducted primarily in two cell lines and one immunocompromised mouse xenograft model, which lacks a fully functional immune system to fully recapitulate the immunomodulatory effects of PD-L1 blockade. Future investigations should employ immunocompetent syngeneic models or humanized mouse models to validate the immune-mediated mechanisms of the combination. Additionally, while we identified the p-ERK/NF-κB/SLUG pathway, other signaling cascades or immune-related factors may contribute to the observed synergy and warrant further exploration.

In conclusion, our data provide strong preclinical evidence that the sequential combination of apatinib and a PD-L1 inhibitor represents a potent therapeutic strategy against breast cancer. The synergy is mechanistically grounded in the inactivation of the p-ERK/NF-κB/SLUG pathway, offering a novel molecular insight for this combination. These findings warrant further clinical investigation to assess the efficacy and safety of this optimized regimen in patients with breast cancer.

Acknowledgements

The authors sincerely thank Luoyang Central Hospital for providing the experimental facilities and fostering an academic environment conducive to learning and exchange during the course of this research.

Authors’ Contributions

All authors provided substantial intellectual input throughout the study, participating in its design, analysis and interpretation of the data, and the development of the manuscript. Specifically, Juanjuan Xu was primarily responsible for the study’s conceptualization and methodology. Her investigation included performing the in vitro experiments and conducting subsequent data analysis of the manuscript. Cairu Guo contributed to the investigation by conducting the in vivo experiments and performing data validation. All authors have read and approved the final submitted version of the manuscript.

Funding

This study received no financial support from public, commercial, or non-profit organizations.

Data Availability

No datasets were generated or analysed during the current study.

Declarations

Ethical approval and consent to participate

This study was approved by the Medical Ethics Committee of Luoyang Central Hospital (Approval No.: LWLL-2024-11-05-01), and all procedures were conducted in accordance with the institutional animal guidelines.

Competing Interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  • 1.Ferrara N. Vascular endothelial growth factor: molecular and biological aspects [J]. Curr Top Microbiol Immunol. 1999;237:1–30. 10.1007/978-3-642-59953-8_1. [DOI] [PubMed] [Google Scholar]
  • 2.Huang M, Huang B, Li G, Zeng S. Apatinib affect VEGF-mediated cell proliferation, migration, invasion via blocking VEGFR2/RAF/MEK/ERK and PI3K/AKT pathways in cholangiocarcinoma cells. BMC Gastroenterol. 2018;18(1):169. 10.1186/s12876-018-0870-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Wu YP, Wu JJ, Tian SM, Jin T, Li C, Xie K. Apatinib with EGFR-TKIs in advanced wild gene-type NSCLC: A case report. Med (Baltim). 2019;98(1):e13924. 10.1097/MD.0000000000013924. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Jin J, Du J, Wu Y. Apatinib monotherapy for advanced VEGFR-2-negative nasopharyngeal carcinoma: A case report. Med (Baltim). 2019;98(1):e13491. 10.1097/MD.0000000000013491. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Liu Z, Wang C, Wu G, Cheng J. MRI analysis of hydrogel-loaded apatinib for local therapy of hepatocellular carcinoma model in nude mice. Biochem Biophys Res Commun. 2019;509(2):529–34. 10.1016/j.bbrc.2018.12.120. [DOI] [PubMed] [Google Scholar]
  • 6.Zou W, Chen L. Inhibitory B7-family molecules in the tumour microenvironment. Nat Rev Immunol. 2008;8(6):467–77. 10.1038/nri2326. [DOI] [PubMed]
  • 7.Liu B, Song Y, Liu D. Recent development in clinical applications of PD-1 and PD-L1 antibodies for cancer immunotherapy. J Hematol Oncol. 2017;10(1):174. 10.1186/s13045-017-0541-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Dedecker H, Teuwen LA, Vandamme T, Domen A, Prenen H. The role of immunotherapy in esophageal and gastric cancer. Clin Colorectal Cancer. 2023;22(2):175–82. 10.1016/j.clcc.2023.03.001. [DOI] [PubMed] [Google Scholar]
  • 9.Diggs LP, Hsueh EC. Utility of PD-L1 immunohistochemistry assays for predicting PD-1/PD-L1 inhibitor response. Biomark Res. 2017. 10.1186/s40364-017-0093-8. 5,12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Topalian SL, Hodi FS, Brahmer JR, Gettinger SN, Smith DC, Mcdermott DF, Powderly JD, Carvajal RD, Sosman JA, Atkins MB, Leming PD, Spigel DR, Antonia SJ, Horn L, Drake CG, Pardoll DM, Chen L, Sharfman WH, Anders RA, Taube JM, Mcmiller TL, Xu H, Korman AJ, Jure-Kunkel M, Agrawal S, Mcdonald D, Kollia GD, Gupta A, Wigginton JM, Sznol M. Safety, activity, and immune correlates of anti-PD-1 antibody in cancer. N Engl J Med. 2012;366(26):2443–54. 10.1056/NEJMoa1200690. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Hamid O, Robert C, Daud A, Hodi FS, Hwu WJ, Kefford R, Wolchok JD, Hersey P, Joseph RW, Weber JS, Dronca R, Gangadhar TC, Patnaik A, Zarour H, Joshua AM, Gergich K, Elassaiss-Schaap J, Algazi A, Mateus C, Boasberg P, Tumeh PC, Chmielowski B, Ebbinghaus SW, Li XN, Kang SP, Ribas A. Safety and tumor responses with lambrolizumab (anti-PD-1) in melanoma. N Engl J Med. 2013;369(2):134–44. 10.1056/NEJMoa1305133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Hodi FS, O’day SJ, Mcdermott DF, Weber RW, Sosman JA, Haanen JB, Gonzalez R, Robert C, Schadendorf D, Hassel JC, Akerley W, Van Den Eertwegh AJ, Lutzky J, Lorigan P, Vaubel JM, Linette GP, Hogg D, Ottensmeier CH, Lebbé C, Peschel C, Quirt I, Clark JI, Wolchok JD, Weber JS, Tian J, Yellin MJ, Nichol GM, Hoos A, Urba WJ. Improved survival with ipilimumab in patients with metastatic melanoma. N Engl J Med. 2010;363(8):711–23. 10.1056/NEJMoa1003466. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Schoenherr C, Pietzsch S, Barca C, Müller FE, Bahr FS, Kasten M, Zeug A, Erschow S, Falk CS, Ponimaskin E, Thackeray JT, Hilfiker-Kleiner D, Ricke-Hoch M. Immune-checkpoint-inhibitor therapy directed against PD-L1 is tolerated in the heart without manifestation of cardiac inflammation in a preclinical reversible melanoma mouse model. Front Mol Med. 2025;4:1487526. 10.3389/fmmed.2024.1487526. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Bonotto M, Gerratana L, Poletto E, Driol P, Giangreco M, Russo S, Minisini AM, Andreetta C, Mansutti M, Pisa FE, Fasola G, Puglisi F. Measures of outcome in metastatic breast cancer: insights from a real-world scenario. Oncologist. 2014;19(6):608–15. 10.1634/theoncologist.2014-0002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Liu K, Ren T, Huang Y, Sun K, Bao X, Wang S, Zheng B, Guo W. Apatinib promotes autophagy and apoptosis through VEGFR2/STAT3/BCL-2 signaling in osteosarcoma. Cell Death Dis. 2017;8(8):e3015. 10.1038/cddis.2017.422. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Emens LA. Breast cancer immunotherapy: facts and hopes. Clin Cancer Res. 2018;24(3):511–20. 10.1158/1078-0432.CCR-16-3001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Yu X, Fan H, Jiang X, Zheng W, Yang Y, Jin M, Ma X, Jiang W. Apatinib induces apoptosis and autophagy via the PI3K/AKT/mTOR and MAPK/ERK signaling pathways in neuroblastoma. Oncol Lett. 2020;20(4):52. 10.3892/ol.2020.11913. [DOI] [PMC free article] [PubMed]
  • 18.Qiu XY, Hu DX, Chen WQ, Chen RQ, Qian SR, Li CY, Li YJ, Xiong XX, Liu D, Pan F, Yu SB, Chen XQ. PD-L1 confers glioblastoma multiforme malignancy via Ras binding and Ras/Erk/EMT activation. Biochim Biophys Acta Mol Basis Dis. 2018;1864(5 Pt A):1754–69. 10.1016/j.bbadis.2018.03.002. [DOI] [PubMed] [Google Scholar]
  • 19.Zhao W, Ma N, Wang S, Mo Y, Zhang Z, Huang G, Midorikawa K, Hiraku Y, Oikawa S, Murata M, Takeuchi K. RERG suppresses cell proliferation, migration and angiogenesis through ERK/NF-κB signaling pathway in nasopharyngeal carcinoma. J Exp Clin Cancer Res. 2017;36(1):88. 10.1186/s13046-017-0554-9. [DOI] [PMC free article] [PubMed]
  • 20.Pires BR, Mencalha AL, Ferreira GM, De Souza WF, Morgado-Díaz JA, Maia AM, Corrêa S, Abdelhay ES. NF-kappaB is involved in the regulation of EMT genes in breast cancer cells. PLoS ONE. 2017;12(1):e0169622. 10.1371/journal.pone.0169622. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Zhao S, Ren S, Jiang T, Zhu B, Li X, Zhao C, Jia Y, Shi J, Zhang L, Liu X, Qiao M, Chen X, Su C, Yu H, Zhou C, Zhang J, Camidge DR, Hirsch FR. Low-Dose apatinib optimizes tumor microenvironment and potentiates antitumor effect of PD-1/PD-L1 Blockade in lung cancer. Cancer Immunol Res. 2019;7(4):630–43. 10.1158/2326-6066.CIR-17-0640. [DOI] [PubMed] [Google Scholar]
  • 22.Zhang Y, Wang F, Sun HR, Huang YK, Gao JP, Huang H. Apatinib combined with PD-L1 Blockade synergistically enhances antitumor immune responses and promotes HEV formation in gastric cancer. J Cancer Res Clin Oncol. 2021;147(8):2209–22. 10.1007/s00432-021-03633-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Zhou C, Wang Y, Zhao J, Chen G, Liu Z, Gu K, Huang M, He J, Chen J, Ma Z, Feng J, Shi J, Yu X, Cheng Y, Yao Y, Chen Y, Guo R, Lin X, Wang Z, Gao G, Wang Q, Li W, Yang X, Wu L, Zhang J, Ren S. Efficacy and biomarker analysis of camrelizumab in combination with apatinib in patients with advanced nonsquamous NSCLC previously treated with chemotherapy. Clin Cancer Res. 2021;27(5):1296–304. 10.1158/1078-0432.CCR-20-3136. [DOI] [PubMed] [Google Scholar]
  • 24.Schmittnaegel M, Rigamonti N, Kadioglu E, Cassará A, Wyser Rmili C, Kiialainen A, Kienast Y, Mueller HJ, Ooi CH, Laoui D, De Palma M. Dual angiopoietin-2 and VEGFA Inhibition elicits antitumor immunity that is enhanced by PD-1 checkpoint Blockade. Sci Transl Med. 2017;9(385):eaak9670. 10.1126/scitranslmed.aak9670. [DOI] [PubMed] [Google Scholar]
  • 25.Allen E, Jabouille A, Rivera LB, Lodewijckx I, Missiaen R, Steri V, Feyen K, Tawney J, Hanahan D, Michael IP, Bergers G. Combined antiangiogenic and anti-PD-L1 therapy stimulates tumor immunity through HEV formation. Sci Transl Med. 2017;9(385):eaak9679. 10.1126/scitranslmed.aak9679. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Wang Q, Gao J, Di W, Wu X. Anti-angiogenesis therapy overcomes the innate resistance to PD-1/PD-L1 Blockade in VEGFA-overexpressed mouse tumor models. Cancer Immunol Immunother. 2020;69(9):1781–99. 10.1007/s00262-020-02576-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Hamzah J, Jugold M, Kiessling F, Rigby P, Manzur M, Marti HH, Rabie T, Kaden S, Gröne HJ, Hämmerling GJ, Arnold B, Ganss R. Vascular normalization in Rgs5-deficient tumours promotes immune destruction. Nature. 2008;453(7193):410–4. 10.1038/nature06868. [DOI] [PubMed] [Google Scholar]
  • 28.Huang Y, Yuan J, Righi E, Kamoun WS, Ancukiewicz M, Nezivar J, Santosuosso M, Martin JD, Martin MR, Vianello F, Leblanc P, Munn LL, Huang P, Duda DG, Fukumura D, Jain RK, Poznansky MC. Vascular normalizing doses of antiangiogenic treatment reprogram the immunosuppressive tumor microenvironment and enhance immunotherapy. Proc Natl Acad Sci USA. 2012;109(43):17561–6. 10.1073/pnas.1215397109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Zhang H, Sun J, Ju W, Li B, Lou Y, Zhang G, Liang G, Luo X. Apatinib suppresses breast cancer cells proliferation and invasion via Angiomotin Inhibition. Am J Transl Res. 2019;11(7):4460–9. [PMC free article] [PubMed] [Google Scholar]
  • 30.Dirix LY, Takacs I, Jerusalem G, Nikolinakos P, Arkenau HT, Forero-Torres A, Boccia R, Lippman ME, Somer R, Smakal M, Emens LA, Hrinczenko B, Edenfield W, Gurtler J, Von Heydebreck A, Grote HJ, Chin K, Hamilton EP. Avelumab, an anti-PD-L1 antibody, in patients with locally advanced or metastatic breast cancer: a phase 1b JAVELIN solid tumor study]. Breast Cancer Res Treat. 2018;167(3):671–86. 10.1007/s10549-017-4537-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Heidegger S, Wintges A, Stritzke F, Bek S, Steiger K, Koenig PA, Göttert S, Engleitner T, Öllinger R, Nedelko T, Fischer JC, Makarov V, Winter C, Rad R, Van Den Brink MRM, Ruland J, Bassermann F, Chan TA, Haas T, Poeck H. RIG-I activation is critical for responsiveness to checkpoint Blockade. Sci Immunol. 2019;4(39). 10.1126/sciimmunol.aau8943. [DOI] [PubMed]
  • 32.Kajita M, Mcclinic KN, Wade PA. Aberrant expression of the transcription factors snail and slug alters the response to genotoxic stress. Mol Cell Biol. 2004;24(17):7559–66. 10.1128/MCB.24.17.7559-7566.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Luo X, Zou W, Wei Z, et al. Inducing vascular normalization: A promising strategy for immunotherapy. Int Immunopharmacol. 2022;112:109167. 10.1016/j.intimp.2022.109167. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

No datasets were generated or analysed during the current study.

Articles from Journal of Mammary Gland Biology and Neoplasia are provided here courtesy of Springer

RESOURCES