Estradiol Promotes Tumor Progression in ERα‐Low Endometrial Cancer via the GPER/SphK1 Pathway

ABSTRACT

Endometrial cancer (EC) is the most common gynecological malignancy in postmenopausal women. Patients with low estrogen receptor alpha (ERα) expression frequently develop aggressive pathological subtypes and have poor prognosis. The role of estradiol (E2) in ERα‐low EC remains poorly understood. This study investigated the tumor‐promoting effects of E2 using clinical data, in vitro functional assays, and a mouse xenograft model. We found that serum E2 levels were elevated in ERα‐low patients, of whom 67.7% showed high tumor expression of G protein‐coupled estrogen receptor (GPER). High GPER expression correlated with increased E2 levels, enhanced sphingosine kinase 1 (SphK1) activity, and higher Ki67 proliferation index. In vitro, E2 promoted proliferation, migration, and invasion of HEC‐1A cells (low ERα, high GPER). These effects were suppressed by pharmacological inhibition or siRNA knockdown of GPER or SphK1. Mechanistically, E2 activated the ERK1/2 pathway via the GPER/SphK1 axis, leading to upregulation of Cyclin D1, Cyclin E1, and MMP‐9. In vivo, E2 stimulated xenograft tumor growth, an effect mitigated by inhibitors of GPER, SphK1, and ERK. Our findings demonstrate that E2 drives progression of ERα‐low endometrial cancer through the GPER/SphK1 signaling pathway, revealing potential therapeutic targets for this high‐risk subgroup.

Estradiol (E2) promotes tumor progression in ERα‐low endometrial cancer through the GPER/SphK1/ERK1/2 signaling axis. E2 binding to GPER activates SphK1 and downstream ERK1/2, upregulating Cyclin D1, Cyclin E1, and MMP‐9 to drive cell proliferation, migration, and invasion. This mechanism is supported by clinical findings (35.0% ERα‐low; 67.7% GPER‐high with elevated E2, increased SphK1 activity, and higher Ki67 index), in vitro validation in HEC‐1A cells (ERα‐low/GPER‐high), and in vivo confirmation in a mouse xenograft model, where G15, DMS, or PD98059 suppress tumor growth.

1. Introduction

Endometrial cancer is historically classified into two main types. Type I, representing the majority of cases, is estrogen‐dependent and primarily consists of endometrioid adenocarcinoma arising from unopposed estrogen exposure. In contrast, Type II tumors arise from atrophic endometrium and encompass aggressive histologies such as serous carcinoma, clear cell carcinoma, and carcinosarcoma. Their distinct molecular profiles suggest different etiological pathways [1]. However, this binary classification is increasingly challenged by overlapping risk factors [2] and comparable circulating estradiol (E2) levels [3]. Furthermore, a significant proportion of both Type I (96%) and Type II (82%) tumors express estrogen receptor alpha (ERα) [4], indicating that estrogen signaling may contribute to Type II pathogenesis.

Elevated estrogen levels are found in endometrial tumors compared to normal tissue, irrespective of ERα status [5]. The comparable circulating E2 levels in Type I and II cancers [3] suggest that E2 may exert oncogenic effects even in tumors with low classical ER expression, potentially through alternative receptors. G protein‐coupled estrogen receptor (GPER) is a transmembrane receptor that mediates rapid, non‐genomic estrogen signaling upon E2 binding [6]. It is highly expressed in various malignancies, including endometrial cancer [7, 8, 9]. Although ER negativity is more common in Type II cancers [4], GPER expression is consistent across subtypes and independent of ER status [10]. Notably, many ERα‐low tumors exhibit high GPER expression, supporting the hypothesis that E2 may drive progression specifically through GPER in this context.

In breast cancer, GPER expression is negatively correlated with or independent of ERα [11], and elevated estrogen promotes growth of ERα‐negative tumors such as triple‐negative breast cancer [12]. Similarly, in endometrial cancer, high GPER expression correlates with poor prognostic factors—deep myometrial invasion, high grade, aggressive histology, advanced stage—and predicts reduced survival [13, 14, 15]. Despite these associations, the functional roles of E2 and GPER across pathological subtypes of low‐ER endometrial cancer remain unclear.

This study focuses specifically on ERα‐low endometrial cancer, moving beyond the conventional Type I/II dichotomy. Elevated E2 levels were observed in patients with low ERα expression, particularly those with high GPER expression. Using in vitro and in vivo models, we demonstrate that E2 promotes tumor progression via the GPER/SphK1 signaling axis, identifying this pathway as a potential therapeutic target for this aggressive EC subtype.

2. Materials and Methods

2.1. Clinical Samples

This study enrolled 177 postmenopausal patients with pathologically confirmed endometrial cancer who underwent surgical resection at Tianjin First Central Hospital between June 2022 and December 2024. A control group comprised 60 postmenopausal women admitted for benign gynecological conditions during the same period. Endometrial cancer tissues were obtained intraoperatively by members of our research team, with all specimens collected as intracavitary tumor tissues devoid of the myometrial layer. Venous blood samples and endometrial cancer tissues were immediately transferred to the hospital's biospecimen repository following collection, where they were processed and preserved in liquid nitrogen. Archived tissues and blood samples were subsequently retrieved from the biospecimen repository for analysis. The study protocol was approved by the Ethics Committee of Tianjin First Central Hospital, and written informed consent was obtained from all participants prior to enrollment.

2.2. Serum E2 Measurement

Serum estradiol (E2) levels were measured by electrochemiluminescence immunoassay (Roche Diagnostics, Suzhou, China). Assays were performed according to the manufacturer's instructions. E2 concentrations were interpolated from a standard curve. The reference E2 level for postmenopausal women is < 10 pg/mL. Values below the detection limit (5 pg/mL) were recorded as 5 pg/mL for statistical analysis.

2.3. Immunohistochemistry (IHC)

Immunohistochemical analysis was routinely performed on all endometrial cancer tissue samples in the Department of Pathology. All tissues were assessed for ERα, Ki67, and P53 expression, with a subset of tissues assessed for the mismatch repair proteins MLH1, PMS2, MSH2, and MSH6 (a negative result indicates mismatch repair deficiency, i.e., dMMR). These sections were all re‐evaluated. For patients exhibiting low ERα expression (defined as IHC score ≤ 4 [16]), additional immunostaining for GPER (1:100, cat. no. ab260033, Abcam) was conducted. Two pathologists, blinded to clinical data, independently evaluated the immunostaining results using a semi‐quantitative scoring system [16]. This system assessed both the proportion of positively stained tumor cells (scored from 0 to 4) and the staining intensity (scored from 0 to 3). A final IHC score was calculated by multiplying these two individual scores. Based on this product, expression levels were initially categorized as: Negative (0–1), Weakly positive (2–4), Positive (5–8), or Strongly positive (9–12). For the purpose of statistical analysis, a final score of ≤ 4 was defined as low expression, while a score > 4 was defined as high expression.

2.4. SphK1 Assay

Sphingosine kinase 1 (SphK1) activity was measured using a commercial fluorescent substrate assay kit (Echelon Biosciences Inc., #E‐K068). Tissue or cell lysates were normalized to 20 μg of protein per sample. Assays were performed according to the manufacturer's instructions. Signal intensity was inversely correlated with SphK1 activity. Results are expressed as the fluorescence difference between blank control and sample wells, reflecting relative SphK1 activity levels.

2.5. Cell Culture and Reagents

HEC‐1A cells were routinely maintained in phenol red‐containing MEM (ATCC‐modified; Shanghai Yuanpei Biotechnology) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin‐streptomycin. For estrogen‐dependent assays, cells were switched to phenol red‐free MEM containing 10% charcoal‐stripped fetal bovine serum (CS‐FBS; Gibco) to eliminate potential estrogenic contaminants. 17β‐Estradiol (E2), the sphingosine kinase inhibitor DMS (N,N‐dimethylsphingosine), and the ERK1/2 inhibitor PD98059 were purchased from Sigma‐Aldrich. The GPER antagonist G15 was obtained from Cayman Chemical, while the ER antagonist ICI 182780 and the SphK1 agonist K6PC‐5 were from MedChemExpress. For experiments involving pharmacological modulators, cells were pretreated with the corresponding agents for 30 min prior to E2 stimulation.

2.6. Plasma Membrane Isolation

Plasma membrane fractions were isolated from HEC‐1A cells by differential centrifugation. Cells were homogenized in ice‐cold buffer (10 mM Tris‐HCl pH 7.4, 0.25 M sucrose, 1 mM EDTA, protease inhibitors). All steps were performed at 4°C. Nuclei and cell debris were pelleted at 600 × g for 10 min. The supernatant was sequentially centrifuged at 10,000 × g for 20 min and then at 100,000 × g for 60 min to obtain crude membranes. These were further purified on a discontinuous sucrose density gradient (20%/40%/60%) by centrifugation at 100,000 × g for 2 h. The plasma membrane‐enriched fraction at the 20%/40% interface was collected and pelleted. Purity was validated by Western blot using Na⁺/K⁺‐ATPase (plasma membrane), COX IV (mitochondria), and Calnexin (ER) markers, confirming minimal contamination. For GPER detection, samples were mixed with Laemmli buffer and incubated at 37°C for 30 min without boiling.

2.7. siRNA Transfection

HEC‐1A cells at 60%–80% confluence in 6‐well plates were transfected with SphK1 siRNA (sc‐44114, Santa Cruz Biotechnology) or GPER siRNA (sc‐60743, Santa Cruz Biotechnology) according to the manufacturer's instructions. siRNA was mixed with transfection reagent for 30 min and then added to cells. After 6 h incubation, the medium was replaced with complete medium (10% FBS, 1% penicillin‐streptomycin). Cells were harvested for analysis 48–72 h post‐transfection.

2.8. Western Blot Analysis

Total protein extracts or plasma membrane fractions were resolved on 10% SDS‐PAGE gels and transferred onto PVDF membranes. The membranes were blocked with 5% skimmed milk for 2 h at room temperature, followed by overnight incubation at 4°C with primary antibodies targeting the following proteins: β‐actin (1:1000; cat. no. 4970; Cell Signaling Technology, Inc.), ERα (1:1000; cat. no. 8644; Cell Signaling Technology, Inc.), GPER (1:1000; cat. no. ab260033; Abcam), Na⁺/K⁺‐ATPase (1:1000; cat. no. 3010; Cell Signaling Technology, Inc.), ERK1/2 (1:1000; cat. no. 4695; Cell Signaling Technology, Inc.), phosphorylated (p)‐ERK1/2 (1:2,000; cat. no. 4370; Cell Signaling Technology, Inc.), Cyclin D1 (1:1000; cat. no. 55506; Cell Signaling Technology, Inc.), Cyclin E1 (1:1000; cat. no. 20808; Cell Signaling Technology, Inc.), and MMP‐9 (1:1000; cat. no. 13667; Cell Signaling Technology, Inc.). After washing with TBST, the membranes were incubated with the corresponding HRP‐conjugated secondary antibodies (1:1000; cat. no. A0208; Beyotime Biotechnology Co. Ltd.) for 1 h at room temperature. Protein bands were visualized using a ChemiDoc Imaging System (Bio‐Rad) with ECL reagent (Millipore) and quantified using ImageJ software.

2.9. Cell Proliferation Assay

Cell proliferation was assessed by the MTT assay. Cells were seeded in 96‐well plates at 5 × 10³ cells/well. After treatments, 20 µL MTT solution was added and incubated at 37°C for 4 h. The formazan crystals were solubilized with 200 µL DMSO, and the absorbance was measured at 490 nm. Cell viability was expressed as a percentage of controls.

2.10. Cell Migration Assay

Cell migration was evaluated using a wound healing assay. Cells were seeded in 6‐well plates and grown to 90% confluence. A scratch was made with a sterile pipette tip, and the initial wound area was photographed. Cells were then treated with phenol red‐free MEM supplemented with 1% FBS. After the indicated treatments, the same wound areas were rephotographed. Migration rate was quantified by measuring the reduction in wound area.

2.11. Cell Invasion Assay

Cell invasion was assessed using Matrigel‐coated Transwell chambers. Cells in serum‐free medium were seeded in the upper chamber. The lower chamber contained medium with 20% FBS as a chemoattractant. After 48 h, non‐invading cells on the upper surface were removed with a cotton swab. Invaded cells on the lower membrane were fixed, stained with 0.1% crystal violet, and counted under a microscope.

2.12. In Vivo Xenograft Model

Female BALB/c nude mice (4–6 weeks old, Beijing Vital River) were ovariectomized. Two weeks later, HEC‐1A cells (2 × 10⁶ in 0.2 mL) were inoculated subcutaneously into the dorsal region. Tumor dimensions were measured every 3 days with a caliper, and volume was calculated as V = πab ²/6.

One day before inoculation, estrogen‐supplemented mice received a subcutaneous 17β‐estradiol tablet (1.7 mg, 60‐day release). Inhibitor groups received daily intraperitoneal injections of DMS or PD98059, or subcutaneous G15 injections (50 nmol) twice weekly for 4 weeks. Mice were then euthanized; tumors were excised, weighed, and stored for analysis. All procedures were approved by the Animal Care and Use Committee of Tianjin First Central Hospital.

2.13. Statistical Analysis

Data are presented as mean ± SD. Statistical analyses were performed using SPSS 22 or GraphPad Prism 9. Group comparisons were performed using Student's t‐test or one‐way ANOVA. Categorical variables were compared using Pearson's χ ² test or Fisher's exact test. Correlations were assessed by Pearson's correlation analysis. p < 0.05 was considered statistically significant.

3. Results

3.1. Clinical and Pathological Characteristics

Among the 177 enrolled patients, 21.47% presented with Type II endometrial cancer (EC), and 36.16% had high‐grade disease. Patients with Type II tumors were significantly older than those with Type I tumors (66.63 ± 8.21 vs. 62.92 ± 7.79 years, p = 0.011). No significant differences were observed between the two histotypes with respect to gravidity, parity, hypertension, diabetes, or body mass index (BMI). Notably, patients with low‐grade tumors had a significantly higher BMI compared to those with high‐grade tumors (26.42 ± 5.74 vs. 24.29 ± 4.27, p = 0.005).

3.2. Features by ERα Expression Level

Low ERα expression was observed in 35.03% of the tumor tissues. Compared to the high ERα group, patients with low ERα expression exhibited a higher prevalence of Type II histology, a higher Ki67 proliferation index, an increased P53 mutation rate, and more advanced FIGO 2023 stage (Table 1). No significant differences were found between the two groups with respect to age, pregnancy history, or BMI.

Table 1.

Association between ERα expression and clinicopathological features in endometrial carcinoma patients.

Variable	Subtype	Low‐ERα (n = 62)	High‐ERα (n = 115)	p‐value
Ki67 index	(Mean ± SD)	51.61 ± 23.67	41.54 ± 19.71	0.005
Histological type	Type I	35 (56.5)	104 (90.4)	< 0.001
	Type II	27 (43.5)	11 (9.6)
Grade	Low‐grade	20 (32.3)	93 (80.9)	< 0.001
	High‐grade	42 (67.7)	22 (19.1)
P53 Mutation	No	42 (67.7)	104 (90.4)	< 0.001
	Yes	20 (32.3)	11 (9.6)
dMMRa	No	36 (58.1)	86 (74.8)	0.973
	Yes	7 (11.3)	17 (14.8)
	Unknown	19 (30.6)	12 (10.4)
Stage	I/II	42 (67.7)	99 (86.1)	0.004
	III/IV	20 (32.3)	16 (13.9)
LNM	No	52 (83.9)	106 (92.2)	0.089
	Yes	10 (16.1)	9 (7.8)

Note: Data are presented as n (%) unless otherwise indicated.

Abbreviations: dMMR, deficient mismatch repair; ERα, estrogen receptor alpha; LNM, lymph node metastasis; SD, standard deviation.

^a Patients with unknown dMMR status (Low‐ERα: n = 19; High‐ERα: n = 12) were excluded from the statistical analysis for this variable.

3.3. E2, GPER, and SphK1 in Patients With Low ERα Expression

To investigate the potential role of E2 in low ERα endometrial cancer, we measured serum E2 levels. Concentrations were significantly higher in patients with low ERα expression than in controls (12.86 ± 10.81 vs. 7.83 ± 4.73 pg/mL, p = 0.001). Among the 62 patients with low ERα expression, 42 (67.7%) exhibited high GPER expression (Figure 1A–F). Compared to the low GPER subgroup, the high GPER subgroup showed significantly elevated E2 levels, increased SphK1 activity, and a higher Ki67 index (Table 2). No significant differences were observed between GPER subgroups regarding P53 status, mismatch repair protein expression, disease stage, or lymph node metastasis.

Figure 1.

GPER expression and clinicopathologic correlations in low ERα endometrial cancer. (A–F) Representative immunohistochemical staining of GPER in low‐grade endometrioid (A, B), high‐grade endometrioid (C, D), and high‐grade serous (E, F) endometrial adenocarcinoma. Upper panels, ×200; Lower panels, ×400. (G, H) Correlations of serum E2 levels with tumor SphK1 activity (G) and tumor SphK1 activity with Ki67 index (H). (I) Ki67 index in patients stratified by serum E2 levels (cutoff: 10 pg/mL; low E2, n = 33; high E2, n = 29). (J, K) SphK1 activity in tumors stratified by FIGO stage (J) (I/II, n = 42; III/IV, n = 20) and lymph node metastasis (K) (no LNM, n = 52; yes LNM, n = 10). Data are presented as mean ± SD. *p < 0.05, **p < 0.01 (Student's t‐test).

Table 2.

Association between GPER expression and clinicopathological features in patients with low‐ERα endometrial carcinoma.

Variable	Subtype	Low‐GPER (n = 20)	High‐GPER (n = 42)	p‐value
E2 level	(Mean ± SD)	8.60 ± 7.41	14.89 ± 11.63	0.031
SphK1 activity	(Mean ± SD)	224.48 ± 63.01	286.02 ± 100.04	0.005
Ki67 index	(Mean ± SD)	42.25 ± 22.97	56.07 ± 22.94	0.030
Histological type	Type I	12 (60.0)	23 (54.8)	0.697
	Type II	8 (40.0)	19 (45.2)
Grade	Low‐grade	9 (45.0)	11 (26.2)	0.139
	High‐grade	11 (55.0)	31 (73.8)
P53 mutation	No	13 (65.0)	29 (69.0)	0.750
	Yes	7 (35.0)	13 (31.0)
dMMRa	No	15 (75.0)	21 (50.0)	0.170
	Yes	1 (5.0)	6 (14.3)
	Unknown	4 (20.0)	15 (35.7)
Stage	I/II	16 (80.0)	26 (61.9)	0.154
	III/IV	4 (20.0)	16 (38.1)
LNM	No	19 (95.0)	33 (78.6)	0.100
	Yes	1 (5.0)	9 (21.4)

Note: Data are presented as n (%) unless otherwise indicated.

Abbreviations: dMMR, deficient mismatch repair; ERα, estrogen receptor alpha; GPER, G protein‐coupled estrogen receptor; LNM, lymph node metastasis; SD, standard deviation.

^a Patients with unknown dMMR status (Low‐GPER: n = 4; High‐GPER: n = 15) were excluded from the statistical analysis for this variable.

E2 levels were positively correlated with SphK1 activity (Figure 1G), and SphK1 activity was positively correlated with the Ki67 index (Figure 1H). However, no significant correlation was found between E2 levels and the Ki67 index (r = 0.185, p = 0.185). Using the postmenopausal reference cutoff of 10 pg/mL, patients were stratified into high‐ and low‐E2 groups. The high‐E2 group displayed a significantly higher Ki67 index (Figure 1I). Furthermore, SphK1 activity was significantly increased in patients with advanced‐stage tumors (Figure 1J) or lymph node metastasis (Figure 1K).

3.4. E2 Promotes Malignant Phenotypes via GPER in HEC‐1A Cells

We first examined ERα and GPER expression in four endometrial cancer cell lines. HEC‐1 A cells exhibited low ERα expression but high GPER expression, and were therefore selected for subsequent functional studies (Figure 2A). E2 promoted HEC‐1 A cell proliferation in a dose‐ and time‐dependent manner (Figure 2B). At a concentration of 1 × 10⁻⁷M, E2 significantly enhanced cell proliferation, migration, and invasion after 48 h of treatment. These effects were markedly suppressed by the GPER antagonist G15 (Figure 2C–G). In contrast, the ER antagonist ICI 182780 did not significantly affect E2‐induced proliferation or invasion (Figure 2H–J), suggesting that the pro‐malignant effects of E2 are mediated predominantly through GPER rather than classical ERs. This notion was further supported by GPER knockdown using specific siRNA, which effectively abrogated E2‐induced malignant phenotypes (Figure 3A–F).

Figure 2.

The GPER‐SphK1 signaling axis mediates E2‐induced progression in HEC‐1A cells. (A) Expression of ERα and GPER in four endometrial cancer cell lines (Ishikawa, RL95‐2, HEC‐1A, and HEC‐1B) by Western blotting. β‐Actin and Na⁺/K⁺‐ATPase served as loading controls. (B) Dose‐ and time‐dependent effects of E2 on cell viability. HEC‐1A cells were treated with increasing concentrations of E2 (10⁻⁹ to 10⁻⁵M) for 24, 48, or 72 h, and viability was assessed by MTT assay. (C) Effects of G15 (10 µM), DMS (10 µM), and PD98059 (PD, 20 µM) on E2‐induced (1 × 10⁻⁷M, 48 h) cell viability. (D, E) Effects of G15, DMS, and PD98059 on E2‐induced migration. Representative images of wound‐healing assays at 0 and 48 h are shown in (D), with quantitative analysis in (E). (F, G) Effects of G15, DMS, and PD98059 on E2‐induced invasion. Representative images of Transwell assays are shown in (F), with quantitative analysis in (G). (H, I) Effects of the ER antagonist ICI 182780 (1 µM) on E2‐induced proliferation (H) and invasion (I). ICI 182,780 did not attenuate E2‐enhanced proliferation or invasion, excluding the involvement of classical ERs and further supporting that E2 acts through GPER. Data are presented as mean ± SD. *p < 0.05, **p < 0.01 versus control; ^# p < 0.05, ^## p < 0.01 versus E2 group (one‐way ANOVA).

Figure 3.

Genetic knockdown of GPER or SphK1 abrogates E2‐induced malignant phenotypes in HEC‐1A cells. (A) Validation of siRNA‐mediated knockdown of GPER and SphK1 by Western blotting. Na⁺/K⁺‐ATPase and β‐actin served as loading controls. (B) Effects of GPER or SphK1 knockdown on E2‐induced cell viability. Cells were transfected with si‐GPER or si‐SphK1 and treated with E2 (1 × 10⁻⁷M, 48 h); viability was assessed by MTT assay. (C, D) Effects of GPER or SphK1 knockdown on E2‐induced migration. Quantitative analysis of wound‐healing assays is shown in (C), and representative images at 0 and 48 h are shown in (D). (E, F) Effects of GPER or SphK1 knockdown on E2‐induced invasion. Quantitative analysis of Transwell assays is shown in (E), and representative images are shown in (F). Data are presented as mean ± SD. *p < 0.05, **p < 0.01 versus si‐NC; ^# p < 0.05, ^## p < 0.01 versus si‐NC + E2 (one‐way ANOVA).

3.5. E2 Activates SphK1 via GPER

E2 rapidly and transiently increased SphK1 activity in HEC‐1 A cells. Activation was detectable within 5 min, peaked at 15 min, and returned to baseline by 60 min (Figure 4A). This effect was significantly inhibited by the GPER antagonist G15 (Figure 4B), indicating that E2‐induced SphK1 activation is GPER‐dependent.

Figure 4.

E2 activates the GPER/SphK1/ERK axis and regulates downstream targets in HEC‐1 A cells. (A) Time course of E2‐induced SphK1 activation. Cells were treated with E2 (1 × 10⁻⁷M) for indicated times. SphK1 activity peaked at 15 min and returned to baseline by 60 min. (B) G15 inhibits E2‐induced SphK1 activation. Cells were pretreated with G15 (10 µM) for 30 min prior to E2 stimulation (15 min). (C–E) The SphK1 agonist K6PC‐5 rescues G15‐mediated suppression of E2‐induced proliferation (C) and invasion (D, E). Quantitative analysis of invasion is shown in (D), and representative images of Transwell assays are shown in (E). (F) Effects of G15, DMS, and PD98059 on E2‐induced ERK1/2 phosphorylation. All three inhibitors blocked E2‐induced p‐ERK. (G–I) Regulation of Cyclin D1 (G), Cyclin E1 (H), and MMP‐9 (I) expression by E2 and the indicated inhibitors (G15, DMS, PD98059). E2 upregulated these proteins, and this effect was reversed by all three inhibitors. Data are presented as mean ± SD. *p < 0.05, **p < 0.01 versus control; ^# p < 0.05, ^## p < 0.01 versus E2 group (one‐way ANOVA).

3.6. SphK1 Mediates E2‐Induced Malignant Behaviors

We next examined whether SphK1 mediates E2‐induced malignant behaviors in HEC‐1 A cells. Treatment with the SphK inhibitor DMS or siRNA‐mediated SphK1 knockdown significantly suppressed E2‐enhanced proliferation, migration, and invasion (Figure 2C–G; Figure 3B–F). Conversely, treatment with the SphK1 agonist K6PC‐5 effectively rescued the G15‐induced inhibition of cell proliferation and invasion (Figure 4C–E). Collectively, these results demonstrate that E2 promotes malignant phenotypes through GPER‐dependent activation of SphK1.

3.7. The GPER/SphK1 Axis Activates ERK1/2 Signaling

E2 treatment (15 min) induced ERK1/2 phosphorylation, which was blocked by either G15 or DMS (Figure 4F). In contrast, Akt and JNK phosphorylation remained unaffected. E2 upregulated the expression of Cyclin D1, Cyclin E1, and MMP‐9, and these effects were inhibited by G15, DMS, or the ERK inhibitor PD98059 (Figure 4G–I).

3.8. In Vivo Validation in a Xenograft Model

E2 significantly promoted the growth of HEC‐1 A xenografts in ovariectomized mice, as evidenced by increased tumor weight and volume. This effect was suppressed by treatment with G15, DMS, or PD98059 (Figure 5A–C). Analysis of xenograft tissues revealed that E2 increased SphK1 activity, which was attenuated by G15 or DMS but not by PD98059 (Figure 5D), confirming that GPER acts upstream of SphK1. E2 also enhanced ERK1/2 phosphorylation and elevated the protein levels of Cyclin D1, Cyclin E1, and MMP‐9 in tumor tissues. These changes were reversed by treatment with G15, DMS, or PD98059 (Figure 5E–H).

Figure 5.

In vivo validation of the GPER/SphK1/ERK signaling axis in the HEC‐1A xenograft model. (A, B) Effects of G15, DMS, and PD98059 on E2‐driven tumor growth. Final tumor weight is shown in (A), and tumor volume kinetics over time are shown in (B). (C) Representative images of dissected tumors from each treatment group at the end of the experiment. (D) SphK1 activity in tumor tissues following the indicated treatments. (E) ERK1/2 phosphorylation levels in tumor tissues, determined by Western blotting. (F–H) Expression of Cyclin D1 (F), Cyclin E1 (G), and MMP‐9 (H) in tumor tissues, determined by Western blotting. Data are presented as mean ± SD. *p < 0.05, **p < 0.01 versus control; ^# p < 0.05, ^## p < 0.01 versus E2 group (one‐way ANOVA).

4. Discussion

This study provides integrated clinical and experimental evidence that E2 promotes the progression of ERα‐low endometrial cancer through the GPER/SphK1/ERK signaling axis. Moving beyond the traditional Type I/II classification, we focused on the clinically relevant subgroup characterized by low ERα expression, which is associated with aggressive tumor features and poor prognosis.

Consistent with previous reports linking ER‐negative status in the No specific molecular profile (NSMP) subgroup to aggressive characteristics—including high‐grade histology, non‐endometrioid subtypes, and L1CAM overexpression [17]—our clinical data revealed that 35.03% of the endometrial cancer tissues examined exhibited low ERα expression. These patients displayed a higher prevalence of Type II tumors, high‐grade histology, an elevated Ki67 index, an increased P53 mutation rate, and more advanced disease stage, collectively reflecting enhanced tumor aggressiveness and poorer prognosis. Notably, serum E2 levels in these patients were significantly elevated compared to those in controls, challenging the conventional view that such tumors are largely estrogen‐independent. This observation raises the possibility that E2 may exert its effects through alternative pathways, such as those mediated by GPER—a membrane‐associated estrogen receptor known to facilitate rapid non‐genomic signaling. Within this ERα‐low subgroup, high GPER expression was prevalent (67.7%) and significantly correlated with elevated E2 levels, enhanced SphK1 activity, and increased proliferative capacity (Ki67 index), implicating GPER and SphK1 as potential key mediators in this context.

Building on previous evidence that GPER mediates rapid estrogenic signaling in endometrial cancer cells [15, 18], we hypothesized that SphK1 functions as a critical downstream effector linking GPER to ERK activation. This hypothesis was rigorously tested in vitro using HEC‐1 A cells, a model that mirrors the low ERα/high GPER phenotype. E2 robustly stimulated proliferation, migration, and invasion in these cells. Through the use of specific pharmacological inhibitors (G15, DMS) and genetic knockdown approaches, we demonstrated that these oncogenic effects are dependent on both GPER and its downstream effector SphK1.

Previous studies have shown that E2 and the GPER agonist G1 activate Akt or ERK via GPER in endometrial cancer cells [18, 19, 20]. In triple‐negative breast cancer, E2 and G1 rapidly activate ERK1/2 without affecting p‐Akt [21]. E2 activates SphK in breast cancer cells, which in turn activates ERK1/2 and Akt [22, 23]. However, SphK1 stimulates ERK1/2 but not Akt in MCF‐7 cells [24]. The effects of GPER and SphK1 on JNK are cell‐type dependent. In vascular endothelial cells, E2 and G1 activate JNK through GPER [25, 26], whereas GPER knockout activates JNK and promotes apoptosis in breast cancer cells [27]. SphK1 overexpression activates JNK and promotes tumor growth in glioblastoma [28], while SphK1 inhibition activates JNK and suppresses proliferation in prostate and lung cancers [29, 30]. Thus, ERK, Akt, or JNK can mediate GPER signaling in a context‐dependent manner. In the present study, we demonstrate that ERK, but not Akt or JNK, mediates GPER–SphK1 signaling in ERα‐low endometrial cancer cells, wherein E2 binding to GPER triggers rapid SphK1 activation and subsequent ERK1/2 cascade, leading to upregulation of Cyclin D1, Cyclin E1, and MMP‐9.

The in vivo xenograft model further substantiated these findings. E2 significantly accelerated tumor growth, an effect reversed by GPER antagonism, SphK1 blockade, or ERK inhibition. Molecular analyses of tumor tissues—revealing increased SphK1/ERK activity and upregulation of downstream target proteins—were consistent with the in vitro data, confirming the physiological relevance of this pathway.

Our findings align with and extend prior knowledge. The role of GPER in ER‐negative cancers [12, 18, 31] and the association of SphK1 with malignancy [32, 33, 34] are well established. This study connects these players into a coherent, functionally validated signaling axis specific to ERα‐low endometrial cancer. The consistent suppression of tumor‐promoting effects at multiple points along this axis (GPER, SphK1, ERK) underscores its central role and highlights its potential as a therapeutic vulnerability.

4.1. Limitations

Several limitations of this study should be acknowledged. First, although our clinical findings are promising, they were derived from a relatively modest cohort of ERα‐low patients (n = 62). Validation in larger, multi‐center cohorts is warranted to confirm these associations and enhance generalizability. Second, the key mechanistic findings were established in a single cell line (HEC‐1A) that recapitulates the low ERα/high GPER phenotype. While this cell line was specifically selected for its relevance to the clinical subgroup of interest, future studies should extend these observations to additional endometrial cancer cell lines—such as GPER‐overexpressing models in GPER‐low HEC‐1B cells—and patient‐derived organoids to confirm the broader applicability of this signaling axis. Third, although pharmacological inhibitors effectively targeted the pathway in vivo, future studies employing stable GPER or SphK1 knockout models would provide more definitive genetic evidence. Fourth, while we minimized stromal contamination by carefully collecting intraluminal tumor tissues devoid of the myometrium, we did not further purify tumor cells using laser capture microdissection or flow cytometry. Future studies employing these techniques would provide more refined measurements of tumor‐cell‐specific SphK1 activity. Fifth, while we have demonstrated pharmacological rescue using the SphK1 agonist K6PC‐5, future studies employing constitutively active SphK1 (CA‐SphK1) in GPER‐knockdown cells would provide definitive genetic evidence establishing SphK1 as the critical downstream effector of GPER. Finally, the precise molecular mechanism by which GPER activates SphK1 requires further investigation, representing a key direction for future research.

5. Conclusions

In conclusion, we demonstrate that ERα‐low endometrial cancer remains responsive to estrogenic stimulation via an alternative GPER/SphK1/ERK pathway. This pathway drives tumor progression by regulating key cell cycle and invasion molecules. Targeting components of this axis—particularly GPER or SphK1—represents a promising therapeutic strategy for this aggressive, high‐risk endometrial cancer subgroup.

Author Contributions

Xiuwen Wang: investigation, methodology, validation. Haofei Sun: resources, methodology, validation. Xin Wen: data curation, software. Weiwei Xin: investigation, methodology. Zhandong Hu: resources. Zhongwei Zhao: formal analysis, resources. Lizhi Zhang: conceptualization, writing – original draft, writing – review and editing, funding acquisition.

Conflicts of Interest

The authors declare no conflicts of interest.

Wang X., Sun H., Wen X., et al., “Estradiol Promotes Tumor Progression in ERα‐Low Endometrial Cancer via the GPER/SphK1 Pathway,” Journal of Biochemical and Molecular Toxicology 40 (2026): e71112. 10.1002/jbt.71112.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.