Healthy Specific Pathogen-Free (SPF)-grade Sprague Dawley (SD) rats (6–8 weeks old) of both sexes was purchased from the Laboratory Animal Production Department, Southern Medical University. The rats were housed in separate cages in the barrier area of the Animal Experimental Center, Sun Yat-sen Eye Center, Sun Yat-sen University. The SD rats were acclimatized in a holding room at 23 °C ± 4 °C with relative humidity 50% ± 5% for 7 days prior to surgery. In accordance with the ARRIVE guidelines, every effort was made in this study to minimize the number of animals used and to reduce animal suffering. SD rats with normal anterior segments, ciliary bodies, and vitreous structure, free of abnormal hemorrhage and exudation, were selected for the experiment. Primary culture of LC astrocyte was prepared from the LC of adult SD rats, as described previously (Lopez et al., 2020), with minor modifications. According to the AVMA Guidelines on Euthanasia, SD rats were euthanized by intraperitoneal injection of 200 mg/kg sodium pentobarbital, and the eyes were subsequently used for primary culture of LC astrocytes. Briefly, the LC was dissected from the optic disc under sterile conditions. The tissues were seeded in 12-well plates containing 2 mL of Dulbecco modified eagle medium (DMEM; 11,965,084; Gibco, Grand Island, NY, United States) per well, supplemented with 20% fetal bovine serum (FBS; 10099141C; Gibco, Grand Island, NY, United States) and penicillin/streptomycin. After 10–14 days in culture (37 °C, 5% CO₂), non-astrocytic cells, such as neurons and LC cells, were removed by changing the medium. The isolated and purified LC astrocytes were cultured in astrocyte growth medium (CM-M188; Pricella, Wuhan, China) containing 5% FBS. Immunofluorescent staining for glial fibrillary acidic protein (GFAP) revealed that >98% of cells were GFAP-positive astrocytes. This study was approved by Ethics Committee of Affiliated Hospital of Yunnan University/the Second People’s Hospital of Yunnan Province (YNUCARE20210007), and was conducted in accordance with the Association for Research in Vision and Ophthalmology (ARVO) guidelines.

Primary VSMCs were isolated from the thoracic aorta media of SD rats, as described previously (Bertoni et al., 2020). The thoracic aortic media was identified and isolated under sterile conditions. The tissues were dissociated, and resulting cells were plated into 25-cm² culture flasks containing 4 mL of DMEM (11,965,084; Gibco, Grand Island, NY, United States) supplemented with 20% FBS (10099141C; Gibco, Grand Island, NY, United States). The cells reached confluence in approximately 4 weeks (37 °C, 5% CO₂). Over 98% of the cells were confirmed as positive for smooth muscle α-actin (α-SMA; ab124964; 1:300, Abcam, Cambridge, MA, United States).

OGD/R was performed in confluent LC astrocytes cultured in vitro. Cells were washed twice with phosphate-buffered saline (PBS), transferred to an oxygen-depleted, glucose-free AGM medium, and incubated in a hypoxic chamber (94% N2, 5% CO2, 1% O2, 37 °C) for the indicated durations (15 min, 30 min, 1 h, 2 h, 4 h, and 8 h). The cell culture chamber for hyperoxia was strictly regulated by an C21 oxygen and carbon dioxide controller (BioSpherix, Lacona, NY, United States). At the end of the OGD, the cultures were returned to standard medium and incubated under normal conditions (5% CO₂/95% air, 37 °C) for 12, 24, and 72 h, or 1 week, for reperfusion.

Non-contact co-culture of LC astrocytes and VSMCs was performed using Transwell, with LC astrocytes inoculated in the lower chamber and VSMCs in the upper chamber. Briefly, LC astrocytes were seeded to confluence on the bottom of 60-mm Transwell. Primary VSMCs were plated on 24-mm chamber in 6-well plates at a density of 100,000 cells per well. At the time of OGD/R, the chamber with VSMCs were placed in the Transwell containing the confluent monolayer of LC astrocytes for 12, 24, or 72 h, or for 1 week. After OGD/R, the chamber with VSMCs were removed and processed for immunofluorescence, while the cell lysates of LC astrocytes were collected for Western blot analysis.

For inhibitor studies, 100 nm/mL rapamycin (S1039; Selleck, Houston, TX, United States), an mTOR inhibitor (Querfurth and Lee, 2021), was incubated with LC astrocyte for 1 h prior to the addition of OGD/R stimuli, as described previously (Zha et al., 2021). In the Con groups, 0.01% DMSO was used instead of the rapamycin to pre-treated LC astrocyte cultures before the OGD/R exposure. Cell morphogenesis and protein expression were observed.

Cell viability following various OGD treatments was measured using the Cell Counting Kit-8 assay (CCK-8; CK04; Dojindo, Kumamoto, Japan), according to the manufacturer’s protocol. Briefly, cells in 96-well plates (AM-2 ML-RD-S; Corning, Corning, NY, United States) were cultured under OGD for different durations. A 10-μL solution of CCK-8 solution was added to each well, and cells were incubated in a humidified incubator (37 °C, 5% CO2) for 2 h. The optical density at 450 nm was measured using a xMark microplate reader (Bio-Rad, Hercules, CA, United States).

To detect cell apoptosis, terminal deoxynucleotide transferase-mediated dUTP nick-end labeling (TUNEL; C10245; Thermo Fisher Scientific, Waltham, MA, United States) staining was performed, following the manufacturer’s protocol. After OGD treatment on coverslips, cells were fixed in 4% paraformaldehyde and permeabilized with 0.1% Triton X-100. TUNEL reaction buffer was added to each coverslip and incubated for 60 min at 37 °C in the dark. Coverslips were counterstained with DAPI. Coverslips were rinsed and mounted on glass microscope slides (Vector Laboratories, Burlingame, CA, United States), and apoptotic cells were imaged and counted in three different fields on each coverslip using BX53 fluorescence microscopy (Olympus, Tokyo, Japan).

To confirm the hypoxia or normoxia status in cells during OGD/R, an Oxylite oxygen-sensitive microelectrode probe (Oxford Optronix, Rochester Hills, MI, United States) was used to monitor the oxygen concentration in the cell supernatant, following the manufacturer’s instructions. The sterile oxygen-sensitive probe was immersed at least 2 mm below the liquid surface of the cell supernatant to measure and record the oxygen concentration. Recordings were taken for 1 min until the data stabilized, ensuring that the deviation remained below 10%. Four replicates were measured per group. A PO₂ value below 10 mmHg under hypoxic condition confirmed successful OGD.

The cell culture medium was collected following the OGD/R treatment. The release of prostaglandin E2 (PGE₂) from the cell supernatant was measured in duplicate using ELISA (PKGE004B; R&D Systems, Minneapolis, MN, United States) according to the manufacturer’s instructions. In brief, the calibration diluent (150 μL) was added to each well of the microplate, followed by the addition of the standard substance (150 μL) and the cell supernatant sample (150 μL), respectively. Afterwards, the primary antibody (50 μL) and the PGE2 conjugate (50 μL) were added to each well in sequence, and incubated at room temperature for 1 h and 2 h, respectively. After washing the microplate four times, substrate solution (200 μL) was added and incubated at room temperature for 30 min, followed by the addition of stop solution (100 μL). The absorbance was measured at a wavelength of 450 nm using a microplate reader (A51119700C; Thermo Fisher Scientific). The level of PGE2 in the sample was calculated based on the standard curve.

Cells grown on glass coverslips were washed twice with PBS, fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.1% Triton X-100 for 10 min, blocked with 5% FBS for 30 min at 37 °C, and then incubated at 4 °C for 24 h with following primary antibodies: rabbit polyclonal antibody to glial fibrillary acid protein (GFAP; ab7260; 1:5,000; Abcam, Cambridge, MA, United States), rabbit polyclonal antibody to cytosolic phospholipase A2 (cPLA2; ab73406; 1:500; Abcam, Cambridge, MA, United States), rabbit monoclonal antibody to mTOR (2,983; 1:200; Cell Signaling Technology, Danvers, MA, United States), rabbit monoclonal antibody to phosphorylated mTOR (p-mTOR; 5,536; 1:100; Ser2448; Cell Signaling Technology, Danvers, MA, United States), and rabbit polyclonal antibody to myosin phosphatase target subunit 1 (MYPT1; ab59235; 1:100; Abcam, Cambridge, MA, United States), or PBS (as negative control). After washing with PBS, cells were incubated at 37 °C for 1 h with the following secondary antibodies: goat anti-rabbit IgG H&L (ab150077; 1:500; Alexa Fluor® 488; Abcam, Cambridge, MA, United States). The cell nuclei were stained with DAPI, and fluorescent images were acquired using a Microphot-FX Research Microscope (Nikon, Tokyo, Japan). Five fields of view were randomly selected from each biological replicate. The overall mean fluorescence intensity of each field of view was analyzed using ImageJ software. The average value of the five fields of view represents the mean fluorescence intensity of this biological replicate.

Proteins were extracted from rat cells using a protein extraction kit (P0013G; Beyotime, Beijing, China). Protein concentrations were quantified using a BCA protein assay kit (23,225; Pierce, Rockford, IL, United States), and proteins were separated by 8%–12% SDS-PAGE electrophoresis, then transferred to PVDF membranes (1,620,177; Bio-Rad, Hercules, CA, United States). Membranes were incubated with the following primary antibodies: rabbit polyclonal antibody to GFAP (ab7260; 1:5,000; Abcam, Cambridge, MA, United States), rabbit polyclonal antibody to cPLA2 (ab73406; 1:1,000; Abcam, Cambridge, MA, United States), rabbit monoclonal antibody to mTOR (2,983; 1:1,000; Cell Signaling Technology, Danvers, MA, United States), rabbit monoclonal antibody to p-mTOR (5,536; 1:1,000; Ser2448; Cell Signaling Technology, Danvers, MA, United States), and subsequently incubated with goat anti-rabbit IgG H&L (ab150077; 1:500; Alexa Fluor® 488; Abcam, Cambridge, MA, United States). Rabbit monoclonal antibody to glyceraldehyde 3-phosphate dehydrogenase (GAPDH; 5,174; 1:1,000; Cell Signaling Technology, Danvers, MA, United States) was used as the protein-loading control of total lysates. Finally, the PVDF membranes were visualized using a G-BOX image capture system (Syngene, Cambridge, MA, United States), and band intensities were quantified using ImageJ 1.0 software (NIH, Bethesda, MD, United States).

All data are presented as the mean ± standard deviation. The Shapiro-Wilk test was employed to assess data normality. Data were normally distributed. Differences between groups were analyzed using One-way analysis of variance followed by Tukey’s post hoc test or t-test, performed with SPSS 20.0 software (IBM, Almonk, NY, United States). Visualization was conducted using GraphPad Prism 8.0.0 software (GraphPad, San Diego, CA, United States). Differences were considered significant with a value of P < 0.05.

To establish a stable OGD cell model, we measured the PO₂ in the cell supernatant and conducted CCK-8 assay and TUNEL staining to determine the optimal OGD exposure time. Cells were exposed to OGD for six time periods: 15, 30, 60, 120, 240, and 480 min. As shown in Figure 1A, the PO₂ in the supernatant was significantly reduced to below 10 mmHg after 1 h of OGD exposure, confirming that the cells were in a hypoxic state. These results demonstrated that OGD (1% O2, serum-free, and no sugar) exposure for more than 1 h successfully created a hypoxic environment in the culture medium. Compared with the Con group, LC astrocytes in the OGD group exhibited reduced survival rate and increased numbers of TUNEL-positive cells (Figures 1B–D, P < 0.05). Within 30 min of OGD, LC astrocyte viability decreased gradually, and after 1 h, cell viability decreased significantly, reaching 40%–60%, and by 8 h, cell viability decreased to 30% (Figure 1B). The apoptosis rate of LC astrocytes ranged from 40% to 50% at 1 h of OGD induction (Figures 1C,D). Therefore, we used the 1 h of OGD treatment in subsequent experiments, as it significantly reduced cell survival (53.00% ± 6.56%) and enhanced apoptosis (41.67% ± 8.50%).

Next, we tested four reoxygenation time points (12 h, 24 h, 3 days, and 7 days) following 1 h of OGD to determine whether the OGD/R model could activate LC astrocytes. Following reoxygenation, the area of LC astrocytes increased significantly, especially at 24 h (Figure 2A). Additionally, OGD/R resulted in a pronounced decrease in the length-to-width ratio of LC astrocytes at 12 h and 24 h, and a pronounced increase at 3 days and 7 days (Figure 2A). Immunostaining and Western blotting revealed that GFAP expression was significantly elevated in LC astrocytes at 12 h, 24 h, 3 days and 7 days after recovery, peaking at 24 h post-reoxygenation (Figures 2B–D, P < 0.05). Thus, OGD/R induced the activation of LC astrocytes, promoting morphological changes and enhancing GFAP expression.

To investigate whether reactive LC astrocytes play a protective role in blood vessel regulation, similar to cortical astrocytes (Wahis et al., 2021), we measured vasoactive arachidonic acid (AA) metabolites around the LC astrocytes. LC astrocytes and supernatants were collected after 1 h of OGD exposure and recovery for 12 h, 24 h, 3 days, and 7 d cPLA2 expression, which indicates AA release from cells, peaked after 24 h of recovery and slightly decreased after 7days (Figures 3A–C, P < 0.05). Notably, PGE₂ protein secretion in the supernatant was increased after 24 h of recovery (Figure 3D, P < 0.05). These indicates that LC astrocytes secrete vasoactive factors following OGD/R exposure.

To further investigate this, we established a LC astrocyte-VSMCs co-culture system. Immunofluorescent staining revealed that MYPT1 expression was higher in VSMCs co-cultured with reactive LC astrocytes compared to those cultured alone (Figures 4A,B, P < 0.05). Compared to the Con group, the cell surface area of VSMCs was significantly larger, and the cell length-to-width ratio was significantly smaller in the co-culture group (Figures 4C,D, P < 0.05), suggesting a relaxed VSMC morphology. Taken together, these findings reveal that reactive LC astrocytes regulate VSMC morphology following OGD/R exposure.

To further explore the mechanisms linking reactive LC astrocytes to vascular regulation, we hypothesized that the mTOR mediates their function. We first investigated whether mTOR was activated in LC astrocytes after 1 h of OGD/R. The results showed that the expression of mTOR and p-mTOR were significantly higher after recovery (Figures 5A–F, P < 0.05), demonstrating that reactivation of LC astrocytes following OGD/R was associated with activation of mTOR.

To further investigate the role of mTOR in reactive LC astrocytes, we pre-treated primary LC astrocyte cultures with rapamycin prior to OGD/R exposure. A similar effect was observed for GFAP protein expression at 24 h and 3 days (Figures 6A,B, P < 0.05). These results suggest that rapamycin attenuated the increased levels of GFAP in LC astrocytes following OGD/R. We then assessed the expression of cPLA2 to further investigate the role of the mTOR pathway. Both immunofluorescence and Western blot analysis demonstrated that rapamycin treatment suppressed the changes in cPLA2 expression observed in LC astrocyte cultures following OGD/R (Figures 6C,D, P < 0.05). Therefore, mTOR signal pathway mediates the function of LC astrocytes in regulating cPLA2 levels.