Study on the Role of the AMPK/PGC-1α Pathway in Cold-Induced Vascular Endothelial Cell Apoptosis and Uterine Damage

Simple Summary

Primary dysmenorrhea may be associated with uterine vasoconstriction and local ischemia. However, its specific molecular regulatory mechanisms remain incompletely elucidated. Through animal and cellular experimental systems, this study observed that cold exposure induces energy metabolism disorders and enhanced oxidative stress, thereby disrupting uterine microcirculatory homeostasis and promoting vascular endothelial cell apoptosis. Results suggest that AMPK hyperphosphorylation and upregulation of PGC-1α signaling may participate in this process, constituting a key regulatory link in cold-exposure-induced uterine vascular abnormalities. Overall, this study provides new insights into the molecular mechanisms by which cold exacerbates primary dysmenorrhea, though further research is needed to clarify the interactions and regulatory patterns between the AMPK-PGC-1α pathway and other metabolic networks.

Abstract

Cold exposure may influence reproductive health through vascular changes, yet its mechanisms remain underexplored. This study aimed to investigate the impact of cold exposure on uterine blood vessels and the expression of the AMPK/PGC-1α gene and protein in adult female SD rats. A primary dysmenorrhea model was established in female Sprague Dawley rats and subjected to continuous cold exposure. Changes in body weight, ear temperature, and estrous cycle were observed. Superoxide dismutase (SOD) activity and adenosine triphosphate (ATP) levels were measured to assess oxidative stress. Uterine tissue morphology was assessed via small animal ultrasound, microcirculation observed using RFLSI imaging, and vascular morphology along with caspase-3 and AMPK expression evaluated histologically and immunohistochemically. CD31 and TUNEL double immunofluorescence were used to assess vascular endothelial apoptosis levels. Western blot was used to analyze Bax, BCL-2, and pAMPK/AMPK expression levels. In vitro injury models were used to treat human umbilical vein endothelial cells (HUVECs) with cold stimulus using the AMPK inhibitor Compound C. RT-PCR quantified Bax, AMPK, p53, and PGC-1α expression. Hypothermia-exposed rats exhibited significantly reduced body weight and ear temperature (p < 0.05), prolonged estrous cycle (p < 0.01), and decreased uterine index (p < 0.01), accompanied by reduced SOD and ATP levels (p < 0.01, p < 0.05). Ultrasound and flow imaging revealed decreased uterine blood flow velocity in the hypothermia group (p < 0.01). Histomorphology revealed disorganized uterine cell arrangement, reduced uterine vessel count (p < 0.01), and increased mean vessel area (p < 0.01) in cold-exposed uteri. Immunofluorescence detection revealed increased vascular endothelial cell apoptosis (p < 0.05). Western blot results showed that proapoptotic protein Bax was upregulated (p < 0.01), Bcl-2 was downregulated (p < 0.05), p-AMPK and p-AMPK/AMPK ratio were elevated (p < 0.01) after cold exposure; Rt-qPCR results indicated that Bax and P53 mRNA were increased (p < 0.01), while PGC-1α expression was elevated (p < 0.01). Rt-qPCR results showed elevated Bax and p53 mRNA (p < 0.01), along with increased AMPK and PGC-1α expression (p < 0.01) in the cold-exposed group. In human umbilical vein endothelial cells (HUVECs), compound C attenuated cold-induced effects (p < 0.01) and downregulated Bax and AMPK expression (p < 0.01). Cold exposure exacerbates uterine oxidative stress and energy imbalance, disrupts microcirculatory homeostasis, and induces endothelial cell apoptosis. Excessive phosphorylation of AMPK may co-activate PGC-1α, jointly contributing to cold-induced uterine dysfunction and exacerbated dysmenorrhea. This study reveals potential signaling pathways underlying cold-induced uterine vascular abnormalities, providing novel theoretical foundations and targeted intervention strategies for the prevention and treatment of primary dysmenorrhea.

1. Introduction

Primary dysmenorrhea is a common gynecological condition characterized by cramping pain in the lower abdomen occurring before, during, or after menstruation, without any significant pelvic organic pathology. It is often accompanied by symptoms such as nausea, vomiting, and diarrhea [1]. The prevalence rate among women of reproductive age ranges from 45% to 95%, and severe cases can significantly impact quality of life [2]. Currently, the clinical management of primary dysmenorrhea remains primarily symptomatic. Nonsteroidal anti-inflammatory drugs (NSAIDs, such as ibuprofen) serve as first-line treatment, effectively inhibiting prostaglandin synthesis and alleviating pain. However, long-term use may lead to gastrointestinal adverse reactions [3]. Hormonal contraceptives (such as combined oral contraceptives) regulate the menstrual cycle by suppressing ovulation and reducing menstrual flow [4]. However, they may cause irregular bleeding and potentially induce headaches and nausea. Although treatment methods for primary dysmenorrhea are currently available, its pathogenesis remains incompletely understood.

One potential underlying cause of primary dysmenorrhea may be a history of prolonged exposure to cold temperatures. Its primary pathophysiological mechanism involves excessive prostaglandin production in the endometrium, leading to spastic contractions of the uterine smooth muscle, uterine vasoconstriction, and local ischemia, thereby causing uterine dysfunction [5]. Research using a rat cold stress model has confirmed that cold stress can disrupt reproductive hormones, endothelin/nitric oxide, and microcirculation. Lv et al. investigated the mechanism of uterine microvascular injury induced by cold stress. LFWJD intervention restored the estrous cycle in rats, improved uterine pathology and blood flow, regulated vascular endothelial growth factor, and exerted protective effects by inhibiting endoplasmic reticulum stress-related pathways and apoptosis [6]. Additionally, studies utilizing a rat model of primary dysmenorrhea demonstrated that intervention with Taohong Siwu Decoction improves pain behavior and pathological damage to reproductive organs in the model rats. This effect is mediated through the regulation of factors such as PGE₂ and TNF-α, as well as the PI3K/AKT signaling pathway [7]. Further experimental animal studies demonstrate that primary dysmenorrhea is associated with oxidative stress and apoptosis. Peony pollen can alleviate these pathological processes by inhibiting inflammatory responses and regulating the COX-2/PGE₂ pathway, thereby effectively relieving primary dysmenorrhea [8]. However, the specific mechanisms by which cold temperatures influence dysmenorrhea, particularly the molecular pathways mediating cold-exposure-induced uterine vascular dysfunction, remain to be elucidated. Therefore, investigating the association between persistent cold exposure and uterine vascular homeostasis, especially evaluating the role of endothelial-cell-apoptosis-related signaling pathways, is crucial for revealing the pathophysiology of cold-aggravated dysmenorrhea.

The AMPK/PGC1α pathway, as a core regulatory pathway for energy metabolism and cell survival, not only participates in regulating vascular endothelial function and suppressing oxidative stress damage but also plays a crucial role in mediating apoptosis regulation across various diseases [9,10]. Previous studies have demonstrated that activation of the AMPK/PGC1α pathway can reduce cell apoptosis by regulating the expression of downstream apoptosis-related molecules [11]. This study employed rats with a primary dysmenorrhea model as subjects. Combining in vivo cold exposure intervention with in vitro cellular experiments, it employed behavioral assessment, pathological histological observation, biochemical indicator detection, and techniques such as Western blot and PCR to This study focuses on investigating the dynamic changes in uterine vascular function and apoptosis-related signaling pathways under sustained cold exposure. The aim is to clarify the regulatory role of apoptosis in cold-exposure-induced exacerbation of dysmenorrhea and to reveal the key molecular mechanisms by which cold exposure mediates uterine vascular abnormalities and apoptotic imbalance (Figure 1). These findings provide new theoretical foundations and potential therapeutic targets for the prevention and treatment of primary dysmenorrhea. This study hypothesizes that cold exposure induces endothelial cell apoptosis, and that impaired uterine microcirculation is associated with abnormalities in the AMPK/PGC-1α pathway. Using a rat model of cold-induced dysmenorrhea combined with in vitro experiments on human umbilical vein endothelial cells (HUVECs), we aim to: (1) elucidate the effects of cold exposure on uterine microvascular perfusion and endothelial integrity; (2) analyze the correlation between AMPK/PGC-1α signaling alterations and cold-induced endothelial cell apoptosis; (3) explore potential molecular pathways involved to establish a theoretical foundation for future research on cold-related reproductive disorders.

2. Materials and Methods

2.1. Primary Experimental Reagents and Instruments

Instruments: qRT-PCR instrument (qTOWER 2.2, Analytik Jena, Jena, Germany); microcirculation analyzer (TECHMAN, Chengdu, China); tissue panoramic quantitative analysis system (TissueFAXS plus, TissueGnostics, Vienna, Austria); laser speckle flow imaging system (RFLSI III, RWD, Guangzhou, China); Small Animal Ultrasound (SigmaVET, Shanghai, China); Microplate Reader (SpectraMax M2, Molecular Devices, San Jose, CA, USA); Inverted Fluorescence/Optical Microscope (BX51, Olympus Corporation, Tokyo, Japan); Centrifuge (BY-300C, Beijing Baiyang Medical Equipment Co., Ltd., Beijing, China); Dehydrator (JT-12S, Wuhan Junjie Electronics Co., Ltd., Wuhan, China) and Tissue Embedding Machine (JB-P7, Wuhan Junjie Electronics Co., Ltd., Wuhan, China); Paraffin Microtome (RM2235, Leica Microsystems GmbH, Wetzlar, Germany); Electrophoresis and Electrophoretic Transfer System (Mini Trans-Blot^® Electrophoretic Transfer Cell, Thermo Fisher Scientific, Waltham, MA, USA).

Reagents: Estradiol benzoate injection (210106, Shanghai Quanyu Biotechnology Animal Pharmaceutical Co., Ltd., Shanghai, China); Oxytocin injection (140062778, Jiangxi Bolai Pharmaceutical Co., Ltd., Jiujiang, China); Compound C (S730603, Selleck Chemicals, Houston, TX, USA); AMPK polyclonal antibody (10929-2-AP, Proteintech, Wuhan, China); p-AMPK monoclonal antibody (CST #2535S, Cell Signaling Technology, Danvers, MA, USA); Bax antibody (Cat. No. 00073982, Proteintech, Wuhan, China); Bcl-2 Antibody (Catalog No. 00099041, Proteintech, Wuhan, China); β-actin Antibody (20536-1-AP, Proteintech, Wuhan, China); TUNEL Fluorescent Detection Kit (20010096, Beyotime Biotechnology, Shanghai, China); GFP Monoclonal Antibody (VD299006, Thermo Fisher, Waltham, MA, USA); DAPI (JR3369359-6, Abcam, Cambridge, UK); ATP and SOD reagents (05--2021, Lunchangshuo Bio, Xiamen, China); Cell line: HUVECs (provided by Hebei University of Chinese Medicine): endothelial model cells.

2.2. Experimental Animals and Grouping

Thirty healthy SPF-grade female Sprague Dawley (SD) rats, aged 8 weeks, weighing 220 ± 20 g, sexually mature but not pregnant, were provided by Beijing Huafukang Biological Technology Co., Ltd. (Beijing, China) (Experimental Animal Use Permit No.: SYXK(Ji)2020-0004). Rats were housed at the Animal Experiment Center of North China University of Science and Technology under environmental conditions of 20 ± 5 °C temperature and 50 ± 5% humidity, with a 12 h light–dark cycle. They had free access to standard laboratory chow and drinking water. The sample size was determined based on the effect size from a preliminary experiment examining the impact of cold stimulation on SOD levels in rat uteri. Using G*Power 3.1.9.7 with α = 0.05 and 1 − β = 0.80, the sample size was calculated and adjusted to account for a 10–20% dropout rate, ultimately establishing the sample size for each group. Prior to the experiment, rats underwent a 7-day acclimatization period. Eligible animals were randomly assigned using a random number table to either the normal group (NOL, n = 12) or the cold exposure model group (Mod, n = 18). The Model group received daily cold exposure from Day 1 to Day 10 of modeling by immersing the hindlimbs and lower abdomen in a 0 ± 1 °C ice–water mixture for 20 min per session. Concurrently, rats received subcutaneous injections of estradiol benzoate for 10 consecutive days: 0.5 mg/rat on days 1 and 10, and 0.2 mg/rat on days 2–9. Control rats received subcutaneous injections of an equivalent volume of physiological saline. Concurrently, their hind limbs and lower abdomen were immersed in water at 37 ± 1 °C for 20 min daily. Successful model establishment was confirmed by observing cold–wet stagnation-related symptoms in rats, including shivering, arched backs with erect hair, sneezing and curling up, lethargy, loose stools with reduced food intake, and pale purple discoloration of the mouth, ears, nose, paws, and tail [12]. On day 11 of modeling, all rats in the modeling groups received an intraperitoneal injection of oxytocin (2 U/rat), while the control group received an equivalent volume of saline solution via intraperitoneal injection. If rats exhibited pain responses such as abdominal, trunk, and hindlimb contractions or curling after injection, this further confirmed the successful establishment of the cold-induced dysmenorrhea model [13]. Observation continued until Day 21, excluding deceased and non-compliant rats, resulting in a final model group of 12 rats. All rats were confirmed to be in estrus via vaginal smear prior to tissue collection. General condition was recorded by independent observers using a predefined scoring sheet (Appendix A), with both experimenters and data analysts blinded to group assignments. All experimental protocols were approved by the Ethics Committee of North China University of Science and Technology (Approval No.: SQ20230139), and procedures strictly adhered to the ARRIVE guidelines.

2.3. Ultrasound and Laser Speckle Flow Imaging (LSR) Detection

After establishing the model and undergoing 21 days of treatment, the estrous cycle of rats was first monitored via vaginal smears. Only individuals in the late estrus phase were selected for the detection process. On the day of detection, enrolled rats were anesthetized with 2.5% isoflurane inhalation and fixed in a supine position on a temperature-controlled operating table. Prior to ultrasound imaging, abdominal hair was removed using a chemical depilatory agent and rinsed with pre-warmed saline. Transverse section imaging was performed using a 30 MHz high-frequency probe, with the focus precisely positioned at the uterine tissue plane. Endometrial thickness and uterine cavity dimensions were recorded. All ultrasound parameters were collected by the same skilled operator to minimize operator bias. For each rat, 3–5 clear frames were acquired per imaging plane.

Following ultrasound examination, a laser speckle blood flow imaging system was employed with a laser wavelength of 785 nm and an imaging resolution of 639 × 480 pixels. The CCD camera maintained a vertical distance of 25 cm from the uterine surface. Each measurement captured 10–20 s of continuous imaging synchronized with the respiratory cycle, selecting the period with minimal blood flow fluctuation. A circular region of interest (ROI) was delineated along the uterine serosa surface in real-time imaging, avoiding major vascular trunks and shadowed areas. Measurements were taken once at each uterine horn bilaterally, with the system automatically calculating the mean blood flow velocity (mm/s) and relative perfusion units (PU) for each ROI. For each animal, we collected three sets of ROI data from the left and right uterine horns, respectively, and used the mean values for statistical analysis. Data processing was performed using the system’s native software and MATLAB R2020b. Minor respiratory displacement was corrected via an information-theory-based image registration algorithm, and frames with severe artifacts exceeding 10 pixels of displacement were discarded. Left and right uterine horns were analyzed independently, with the average values from both sides combined.

2.4. Observation Indicators and ELISA Detection

Following behavioral observation of the rats, they were anesthetized with isoflurane, and blood samples were collected from the abdominal aorta. On the cold table, both uteri were dissected, weighed to calculate the uterine index, and divided into three portions. These were respectively fixed in 4% paraformaldehyde, preserved in 15% sucrose, and rapidly frozen in liquid nitrogen. Samples were stored at −80 °C for subsequent analysis. Monitor the body weight and ear temperature of rats on days 7, 14, and 21. SOD and ATP levels in the uterus were measured using enzyme-linked immunosorbent assay (ELISA) according to the manufacturer’s instructions.

2.5. Histological Examination of Rat Uterus

Uterine tissue was fixed in 4% paraformaldehyde for over 72 h, followed by sequential dehydration with graded ethanol, clearing with xylene, and paraffin embedding. Sections 4 μm thick were prepared using a microtome. Perform routine hematoxylin-eosin staining on the sections. TissueFAXS panoramic tissue cell quantitative analysis system was used to scan sections. Target areas were selected and magnified at 200× to observe and analyze uterine tissue cell morphology, staining characteristics, and pathological changes. Vascular quantitative analysis utilized sections from the same anatomical location in the middle segment of rat uterine horns. Vessels were identified based on morphological features revealed by HE staining. ImageJ software (version 1.x)was employed for quantification with uniform threshold settings: with grayscale values 150–220 defining vascular lumens. Micro-lumens < 20 μm² were excluded, and only complete vascular lumens with clear boundaries were included. Vessel counts were performed, and lumen area and mean lumen area were measured and calculated.

Paraffin-embedded uterine tissue sections measuring 4 μm thick were dewaxed in water, followed by antigen retrieval and endogenous peroxidase blocking. Subsequently, the sections were incubated with primary antibodies against caspase and AMPK overnight at 4 °C. Secondary antibody incubation was performed at room temperature for 1 h, followed by DAB substrate staining. Sections were then dehydrated, cleared, and mounted. Quantitative analysis of positive cells was conducted using ImageJ software to measure the integral optical density (IOD) and average optical density (AOD, AOD = sum of IOD/sum of area).

2.6. Immunofluorescence and TUNEL Apoptosis Double-Staining of Rat Uterine Tissue

Place frozen uterine sections in cold air to air-dry. Fix in 10% neutral buffered formalin (NBF) for 20 min, then wash three times with PBS. Delineate staining areas using a PAP immunohistochemistry pen. Add balanced buffer and incubate at room temperature for 10 min. After aspirating the buffer, add 1:2500 diluted CD31 rabbit anti-rat primary antibody and incubate overnight at 4 °C in a humidified chamber protected from light. Wash three times with PBS. Add Alexa Fluor 594-labeled red fluorescent goat anti-rabbit IgG secondary antibody (dilution 1:500) and incubate at room temperature in the dark for 2 h. Wash three times with PBS, each wash for 5 min. Proceed according to the TUNEL apoptosis detection kit instructions, then mount with DAPI-containing anti-fluorescence quenching mounting medium. Acquire images in blue, red, and green channels using an upright fluorescence microscope for overlay analysis.

2.7. Cell Culture and Cold Exposure of HUVECs

Human umbilical vein endothelial cells (HUVECs) were cultured and grouped as follows: HUVEC cells were cultured in high-glucose DMEM medium supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin. Cells were routinely cultured in a 37 °C incubator maintained at 95% air + 5% CO₂ with saturated humidity. To elucidate the regulatory role of AMPK in cold-induced cellular dysfunction, three groups were established, each with triplicate wells: ① Control group: Cells incubated at 37 °C for 2 h; ② Moderate-Oxygenation Damage (MOD) group: Cells incubated at 4 °C for 2 h to establish a moderate-oxygenation damage model; ③ Compound C (AMPK-specific inhibitor) + Moderate-Oxygenation Damage (CC + MOD) group: Cells pretreated with 10 μM Compound C followed by incubation at 4 °C for 2 h.

2.8. Western Blot of Rat Uterine Tissue

Collect cell samples from different treatment groups, add RIPA lysis buffer for thorough lysis, then centrifuge at 4 °C at 12,000 rpm for 15 min and collect the supernatant. Determine protein concentration using the BCA protein quantification kit, mix the protein with 1× loading buffer, and denature at 95 °C for 5 min. Load denatured protein samples onto SDS-PAGE gels. Electrophorese at 80 V for 30 min, then adjust to 120 V for 60 min. Transfer proteins to PVDF membranes at 20 V constant voltage. After transfer, the PVDF membrane was blocked in 5% non-fat milk blocking solution on a shaking incubator at room temperature for 1 h. Subsequently, the corresponding primary antibodies (Bax: 1:1000, Bcl-2: 1:1000, p-AMPK: 1:1000, AMPK: 1:1000, β-Actin: 1:5000) and incubated overnight at 4 °C. The next day, the membrane was washed three times with TBST buffer for 10 min each. A 1:2000 diluted HRP-labeled secondary antibody was added and incubated on a shaking incubator at room temperature for 1 h. After another wash, we developed them using an ECL chemiluminescent kit. We captured the bands via an imaging system and analyzed band optical density values using ImageJ software. Finally, we expressed the relative expression levels of each protein as the optical density ratio of the target protein to the β-Actin internal control.

2.9. Rt-qPCR of Rat Uterine Tissue

Total RNA from uterine tissue was extracted using the RNAeasy™ Animal RNA Isolation Kit with Spin Column (Beyotime Biotechnology, Shanghai, China). Subsequently, the RNA was reverse transcribed into cDNA using the Bio-Rad reverse transcription kit.

The sequences of primers used were as follows (

Table 1. Primer sequences for qPCR.

Gene	Forward Primer (5′–3′)	Reverse Primer (5′–3′)
AMPK	TCAGTAGGCACACACATCGCAT	AGGAAGGTCTGTGGGGATTC
PGC-1α	TGGATGAAGACGGATTGCCC	TCTGAGTGCTAAGACCGCTG
P53	AAAGAAGAGCATTGCCCGGA	GTCAGGCCCCACTTTCTTGA
BAX	CTGGACAACAACATGGAG	AAGTAGAAAAGGGCAACC
GAPDH	ACTCTACCCACGGCAAGTTC	TGGGTTTCCCGTTGATGACC

):

After amplification, qPCR software (version 4.1)was used to generate amplification curves and melting curves for real-time qPCR. Using GAPDH as the internal control, the relative expression levels of the target gene in each sample were calculated using the 2^(−ΔΔCT) method.

2.10. Statistical Analysis

All experimental data were analyzed using IBM SPSS 25.0 statistical software. Quantitative data are expressed as mean ± standard deviation. When data met the conditions of normal distribution and homogeneity of variance, comparisons among multiple groups were performed using one-way analysis of variance (ANOVA), with pairwise comparisons between groups using the Least Significant Difference (LSD) test; comparisons between two groups used the independent samples t-test. Nonparametric tests were employed when data did not meet normality or homogeneity of variance assumptions. Repeated-measures ANOVA with a group × time interaction term was used to analyze changes in body weight and body temperature over time. Differences were considered statistically significant at p < 0.05.

3. Results

3.1. Changes in Physiological Status and Oxidative Stress Markers in Rats

To clarify the impact of model establishment on the baseline physiological state of rats, this study monitored weight dynamics across different groups. Repeated-measures ANOVA revealed a significant group × time interaction for body weight (p < 0.01). By day 21, MOD group rats exhibited significantly lower body weight compared to the NOL group (p < 0.01, Figure 2A,

Table 2. Ear temperature measurements at different time points in rats (°C, $\stackrel{-}{X}$ ± s).

Group	n	0–7 d	7–14 d	14–21 d
NOL	12	37.17 ± 0.32	36.70 ± 0.18	36.68 ± 0.22
MOD	12	35.73 ± 0.43 **	36.56 ± 0.30	33.44 ± 0.68 **
F		28.76	0.82	56.31
p		0.000	0.373	0.000

** p  <  0.01, NOL vs. MOD (n = 12).

). Ear temperature in the MOD group was also significantly reduced on day 21 (p < 0.01, Figure 2B,

Table 3. Body weight at different time points in rats (°C, $\stackrel{-}{X}$ ± s).

Group	n	0–7 d	7–14 d	14–21 d
NOL	12	213.17 ± 8.72	228.50 ± 11.36	230.83 ± 13.05
MOD	12	206.58 ± 5.57 **	216.17 ± 10.53 **	217.17 ± 10.97 **
F		12.85	15.62	18.37
p		0.000	0.000	0.000

** p  <  0.01, NOL vs. MOD (n = 12).

). Analysis of reproductive-related phenotypes and oxidative stress/energy metabolism indicators revealed that compared to the NOL group, MOD rats exhibited significantly prolonged estrous cycle duration (p < 0.01, Figure 2C) and markedly reduced uterine index (p < 0.01, Figure 2D), simultaneously exhibiting significantly decreased superoxide dismutase (SOD) activity (p < 0.01, Figure 2E) and a significant downward trend in adenosine triphosphate (ATP) levels (p < 0.05, Figure 2F).

3.2. Microcirculation and Tissue Status in Rats

This study analyzed uterine and vascular characteristics in NOL and MOD groups via ultrasound, macroscopic morphology, and laser speckle flow imaging (RFLS). Ultrasound imaging revealed uniform uterine tissue echogenicity in the NOL group, whereas the MOD group exhibited localized hypoechoic areas within the uterine region (Figure 3A). Macroscopic morphological examination revealed a full uterus with regular contours, clear vascular distribution, and good perfusion in the NOL group. In contrast, the MOD group exhibited localized uterine atrophy (red arrows), tissue wrinkling, and dilated, congested vessels with disorganized texture (Figure 3C). Laser speckle flow imaging revealed predominantly high blood flow signals (red areas) in the NOL group with adequate local perfusion, whereas the MOD group exhibited significantly reduced blood flow signals dominated by low flow signals (blue areas) (Figure 3D).

3.3. Histomorphological Features of the Uterus

In the normal group, the epithelium of the uterine mucosa was intact, with clear boundaries distinguishing each layer, including the muscular and serosa layers. The myometrium displayed consistent thickness, and the single-layer columnar epithelium on the uterine glands was uniformly arranged. Additionally, the uterine serosa was vascular-rich and stable in structure, with smooth tube walls. The vessel walls were tightly adjoined to the surrounding tissues, and new capillaries were observed. In contrast, rats from the model group exhibited significant disparities. The endometrial epithelial cells of these patients were more dispersed, and the gland count decreased. The uterine serosa layer had fewer new capillaries, and the blood vessels were irregularly shaped with a thicker lumen. Furthermore, the integrity of the blood vessel wall was compromised, and the endothelial layer showed signs of damage (Figure 4A). Compared with the normal group, the model group presented a significant decrease in the total number of uterine blood vessels (p < 0.01). However, the total area and average single vessel area of the uterine blood vessels in the model group were significantly greater (p < 0.01) (Figure 4B–D).

Immunohistochemical staining revealed apoptosis in rat uterine tissue. Compared with the NOL group, the MOD group exhibited significantly elevated levels of Caspase-3 and AMPK (p < 0.01, Figure 4G,H). Notably, most of the apoptotic cells were concentrated in the blood vessels and glandular ducts (Figure 4E,F).

3.4. Immunofluorescence Detection of Apoptosis in Uterine Vascular Endothelial Cells

Using dual immunofluorescence staining with CD31 and TUNEL to detect apoptosis in vascular endothelial cells within uterine tissue, the fluorescence intensity of CD31-positive cells in the MOD group was significantly reduced compared to the NOL group (Figure 5(Ac,Bc)), while the number of TUNEL-positive apoptotic cells increased significantly (Figure 5(Ad,Bd)). TUNEL fluorescence signals showed high colocalization with CD31-labeled vascular areas, indicating that apoptotic cells were primarily enriched in vascular endothelial regions (Figure 5(Aa,Ba)).

3.5. Regulatory Effects of Low Temperature on Apoptosis-Related Proteins and the AMPK Pathway

Compared with the NOL group, the MOD group exhibited increased Bax expression (p < 0.01) and decreased BCL-2 expression (p < 0.05). In the CC + MOD group, Bax expression was reduced compared to the MOD group (p < 0.05), while BCL-2 expression levels increased (p < 0.01). The relative expression of p-AMPK and the p-AMPK/AMPK phosphorylation ratio were significantly higher in the MOD group than in the NOL group (p < 0.01). In contrast, both the relative expression of p-AMPK and the p-AMPK/AMPK ratio were significantly lower in the CC + MOD group than in the MOD group (p < 0.01, Figure 6A). Quantitative analysis of each protein is shown in Figure 6B.

3.6. Quantitative Reverse Transcription Polymerase Chain Reaction (RT–PCR) Analysis

Quantitative RT-PCR results revealed significantly elevated levels of Bax and p53 mRNA in the MOD group compared to the NOL group (p < 0.01). Concurrently, RT-PCR product levels of PGC-1α and AMPK were markedly increased in the MOD group (p < 0.01). Refer to Figure 7A–D for details.

4. Discussion

Primary dysmenorrhea is a common functional disorder of the reproductive system in women of childbearing age. Its core pathological mechanisms involve excessive or imbalanced release of prostaglandins and cyclooxygenase-derived substances from the endometrium, which induce dysfunctional uterine contractions and local ischemia while simultaneously heightening peripheral nerve sensitivity to pain [14]. In recent years, both clinical and animal studies have indicated that cold exposure significantly exacerbates dysmenorrhea symptoms, but the underlying molecular mechanisms remain unclear [7,15]. The AMPK/PGC1α pathway, as a key regulatory pathway for energy metabolism and cell survival, is extensively involved in modulating vascular endothelial function and regulating apoptosis, exerting protective effects in various ischemic diseases [16,17]. This study systematically evaluated the effects of sustained cold exposure on uterine vascular function, oxidative stress levels, apoptosis, and the AMPK/PGC1α pathway through a cold-induced primary dysmenorrhea model and cellular experiments. The aim is to reveal that cold exposure may contribute to uterine dysfunction and exacerbated dysmenorrhea by disrupting the regulatory balance of the AMPK/PGC-1α signaling network, thereby providing new theoretical foundations for clinical prevention and treatment.

This study found that cold-exposed rats exhibited significantly reduced body weight, decreased body temperature, prolonged estrous cycles, and diminished uterine indices. This indicates that low temperatures not only suppress metabolic activity but also induce significant oxidative stress and energy deficiency states. Cold stress disrupts energy metabolism in the uterus and ovaries by inhibiting the body’s energy metabolism and interfering with the hypothalamic–pituitary–ovarian axis function [18]. Additionally, SOD activity and ATP levels in uterine tissue from model group rats were significantly reduced, indicating that cold exposure induces oxidative stress damage in uterine tissue, thereby disrupting uterine smooth muscle contraction and exacerbating dysmenorrhea. Oxidative stress represents a key pathological mechanism in cold stress-mediated tissue injury. As a core enzyme in the body’s antioxidant defense system, SOD plays a crucial role. Cold stress disrupts the body’s oxidation-antioxidation balance, causing oxidative damage to multiple tissues by altering antioxidant status and promoting oxidative reactions [19,20]. As the direct energy source for cellular life activities, ATP homeostasis imbalance serves as a critical regulatory node in apoptosis. Cellular energy depletion triggered by insufficient energy supply can directly activate apoptosis signaling pathways, thereby mediating the initiation of the apoptosis program [21]. Szmidt et al. reviewed 175 cases of women with primary dysmenorrhea and found elevated levels of oxidative stress in these patients [22]. Furthermore, research indicates that Lacerda et al. established a rat model of primary dysmenorrhea, successfully reproducing pathological features such as excessive uterine contractions and heightened pain responses, while also inducing oxidative stress imbalance in uterine tissue [23].

Imbalance in uterine vascular homeostasis is a key pathological feature of primary dysmenorrhea. Ultrasound, laser speckle flow imaging, and histomorphological observations confirm that prolonged exposure to low temperatures significantly disrupts uterine vascular structure and perfusion function in rats. Cold stimulation triggers excessive sympathetic nervous system activation and dysregulation of endothelin-1 (ET-1) secretion, thereby inducing sustained uterine vasoconstriction and microcirculatory impairment [24]. ET1 not only directly acts on vascular smooth muscle cells to induce contraction, but also inhibits nitric oxide (NO) production, significantly impairing vasodilation function and leading to prolonged ischemic hypoxia [25]. Additionally, cold-induced metabolic suppression and oxidative stress can further exacerbate microcirculatory damage. Research indicates that cold exposure reduces blood flow, triggers cellular contraction, and elevates ROS levels. ROS mediate cold-induced vasoconstriction through specific pathways, with vascular tissues rather than nerve fibers, participating in this process [26]. From a pathophysiological perspective, the vascular homeostasis imbalance induced by low temperatures elevates prostaglandin levels, thereby intensifying uterine spasmodic contractions [27]. Song et al. demonstrated that electroacupuncture significantly reduced writhing scores, modulates prostaglandin (PGF2α, PGE2) and β-EP levels, and alleviates dysmenorrhea symptoms [28].

Cold-induced oxidative stress amplifies apoptosis. Dual immunofluorescence detection of CD31 and TUNEL revealed decreased fluorescence intensity in CD31-positive cells and a significant increase in TUNEL-positive cells within the model group. CD31, an endothelial cell-specific marker, is highly expressed at endothelial cell junctions. It maintains the integrity of endothelial cell connections and accelerates the restoration of the vascular permeability barrier following inflammatory or thrombotic challenges [29]. TUNEL detection identifies 3′-OH DNA breaks, and large-fragment/nucleosome-level DNA fragmentation is one of the hallmark events of apoptosis. TUNEL is widely used to identify and quantify apoptotic cells [30]. Research indicates that CD31 can trigger endothelial cell signaling events through monoclonal antibody cross-linking, promoting increased mRNA levels of protective genes A20 and A1 and activating the transcription factor Sp-1, thereby protecting endothelial cells from apoptosis [31]. Additionally, Cheung et al. found that CD31 expressed by endothelial cells can resist TNF-α-induced endothelial cell apoptosis by activating the Erk/Akt pathway, thereby inhibiting the transcription of proapoptotic genes and promoting the expression of anti-apoptotic genes [32]. Low temperatures may disrupt this balancing mechanism, leading to an increase in TUNEL-positive apoptotic cells.

Bax and Bcl-2 are key regulators of apoptosis in the mitochondrial pathway [33,34]. Bcl-2 inhibits caspase activation by stabilizing the mitochondrial membrane potential and preventing cytochrome c release [35]. Bax promotes increased mitochondrial permeability and induces apoptosis [36]. Previous studies have confirmed that prolonged exposure to low temperatures increases the expression of proapoptotic caspases-3 and Bax while decreasing the expression of anti-apoptotic Bcl-2, consistent with the observations in this experiment [37]. AMPK, as an energy-sensing factor, plays a central role in cellular stress responses and metabolic reprogramming [38]. When cellular ATP levels decrease and the AMP/ATP ratio increases, AMP-activated protein kinase (AMPK) is phosphorylated and activated to maintain energy homeostasis. However, under prolonged stress, its excessive activation may contribute to the transmission of proapoptotic signals [39]. He et al. investigated the effects of low temperatures on pigs via the gut-liver axis, finding that cold stress impairs growth performance, while sustained activation of AMPK leads to hepatic oxidative stress and apoptosis [40]. PGC1α is a key downstream target gene of AMPK that regulates cellular energy homeostasis and resistance to oxidative stress [17]. This study observed that cold exposure significantly activates AMPK and upregulates PGC-1α expression, suggesting that abnormalities in the AMPK/PGC-1α pathway correlate with cold-induced uterine tissue apoptosis and microcirculatory damage. Concurrently, cold exposure upregulates p53 and its downstream apoptotic effector molecule Caspase-3, further promoting uterine cell apoptosis. A study indicates that cold stimulation activates the AMPK/PGC-1α signaling pathway, thereby downregulating the Bcl-2/Bax ratio and reducing the expression levels of apoptosis-related proteins such as CytC, Caspase-3, Caspase-9, and Caspase-8. This mechanism inhibits cell apoptosis and mitigates cold-induced tissue damage. p53, a classic apoptotic transcription factor, drives the expression of proapoptotic molecules [41]. p53 is a classical apoptotic transcription factor that drives the expression of proapoptotic molecules [42]. Previous studies have demonstrated that low temperatures induce the expression of oxidative stress-related genes and upregulate apoptosis-related genes such as p53, caspase-9, and caspase-3 [43].

This study analyzed the effects of cold exposure on uterine microcirculatory function and the AMPK-PGC-1α signaling pathway, but certain limitations remain. First, the sample size was relatively small, and individual variations in sensitivity to cold stimulation were not fully reflected in the results. Second, the study did not distinguish between the physiological effects of acute and chronic cold exposure. Differing durations and intensities of cold exposure may trigger distinct regulatory responses in energy metabolism and apoptotic pathways. Finally, at the molecular level, this study primarily focused on the single AMPK-PGC-1α signaling pathway, lacking analysis of other pathways within the oxidative stress and metabolic regulation network.

Future research will expand the sample size to enhance statistical reliability and biological consistency of the results. Subsequently, studies will incorporate models comparing acute and chronic cold exposure to clarify the regulatory mechanisms of different cold exposure patterns, providing a more comprehensive reference for research. Additionally, investigations will be extended to explore more relevant pathways and analyze interactions among them. Through pathway interaction validation and core molecular functional interventions, the underlying molecular mechanisms of cold-exposure-induced uterine microcirculatory dysfunction will be comprehensively elucidated.

5. Conclusions

This study systematically investigated the effects of prolonged cold exposure on uterine vascular function, oxidative stress status, and the AMPK/PGC-1α signaling pathway. Comprehensive behavioral, imaging, and molecular analyses revealed that cold exposure significantly reduced body weight and core body temperature in female SD rats, prolonged the estrous cycle, and induced elevated oxidative stress levels and insufficient energy supply in uterine tissue, manifested as marked decreases in SOD and ATP. Corresponding ultrasound and laser speckle flow imaging confirmed impaired uterine microcirculatory perfusion, while histological examination revealed vascular structural disarray and aggravated endothelial layer damage. Immunofluorescence results indicated reduced CD31-positive cells and increased TUNEL-positive cells, suggesting markedly enhanced vascular endothelial cell apoptosis. Molecular-level detection showed that cold exposure significantly upregulated the proapoptotic protein Bax and the transcription factor p53, while downregulating the anti-apoptotic factor Bcl-2. This was accompanied by AMPK hyperphosphorylation and upregulates of PGC-1α expression. In vitro experiments further confirmed that intervention with the AMPK inhibitor Compound C attenuated cold-induced apoptosis and energy imbalance effects, indicating that abnormal AMPK activation plays a key role in cold-stress-induced cellular damage. Although this study was conducted using rat and human umbilical vein endothelial cell models, it holds significant reference value for human pathology. When women experience prolonged exposure to cold environments or develop abnormal thermoregulation due to lifestyle habits, they may exhibit uterine microcirculatory disorders and energy metabolism imbalances similar to those observed in this experiment. This process may represent a key physiological basis for exacerbated dysmenorrhea symptoms and increased sensitivity to cold.