The glial cell lines U87 (cat. no. CL-0238), U251 (cat. no. CL-0237) and HMC3 (cat. no. CL-0620; all from Procell Life Science & Technology Co., Ltd.) were cultured in complete MEM (HyClone; Cytiva) supplemented with 10% fetal bovine serum (FBS), 1% non-essential amino acids, sodium pyruvate and penicillin-streptomycin. The cells were maintained in a humidified incubator at 37°C with 5% CO₂. When the cells reached ~85% confluence, they were passaged using trypsinization. All cell lines were authenticated using STR profiling.

To investigate the involvement of the proteasome degradation pathway in the downregulation of ATP7A and ATP7B, U87 and U251 glioma cells were treated with the proteasome inhibitor MG132. Specifically, cells were incubated with MG132 (cat. no. HY-13259; MedChemExpress) at a final concentration of 10 μM for 24 h.

Protein extraction was performed from cells or tissue samples using RIPA lysis buffer (Beijing Solarbio Science & Technology Co., Ltd.). The lysates were centrifuged at 10,000 × g for 15 min to remove cellular debris, and the supernatant containing the total protein was collected. Protein concentrations were quantified using a BCA assay. Equal amounts of protein samples (15 μg/lane) were then subjected to 10% SDS-PAGE. Following electrophoresis, the proteins were transferred onto a PVDF membrane (MilliporeSigma). The membrane was then blocked with 5% non-fat milk or BSA (cat. no. A8010; Beijing Solarbio Science & Technology Co., Ltd.) in TBST (TBS with 0.1% Tween-20) for 1 h at room temperature. The membrane was incubated overnight at 4°C with primary antibodies specific to the target proteins SDF2 (cat. no. PK47764), SDF2L1 (cat. no. PK17377), ERdj3 (cat. no. PA2683), GRP78 (cat. no. T55166), ATP7A (cat. no. PA7106), ATP7B (cat. no. TA0410), dihydrolipoamide S-acetyltransferase (DLAT; cat. no. T58125) and GAPDH (cat. no. M20006; 1:1,000; all purchased from Abmart Pharmaceutical Technology Co., Ltd.). Lipoic acid (cat. no. 437695) was purchased from MilliporeSigma. After washing with TBST to remove unbound primary antibodies, the membrane was incubated with HRP-conjugated secondary antibodies (cat. no. ZB-2301 for rabbit; cat. no. ZB-2305 for mouse) Beijing Solarbio Science & Technology Co., Ltd.) for 1 h at room temperature. Following several washes with TBST, the bound antibodies were visualized using an enhanced chemiluminescence (ECL) detection system (Beijing Solarbio Science & Technology Co., Ltd.). The intensity of each protein band was quantified using ImageJ analysis software (version 10; Media Cybernetics, Inc.). GAPDH was used as an internal control.

Total RNA was extracted from cells using TRIzol reagent, and cDNA was synthesized following the instructions provided with the PrimeScript™ RT Master Mix (Perfect Real Time) (Takara Bio, Inc.). The reverse transcription conditions were set at 37°C for 15 min, followed by 85°C for 5 sec and then 4°C indefinitely. For qPCR, a 20-μl reaction mixture consisting of 10 μl of Applied Biosystems™ SYBR Green Master Mix (Thermo Fisher Scientific, Inc.), 1 μl each of forward and reverse primers, 2 μl of cDNA, and 6 μl of ddH₂O was prepared. The PCR protocol included initial denaturation at 95°C for 30 sec, followed by 40 cycles of amplification with denaturation at 95°C for 5 sec, annealing at 60°C for 30 sec, and extension at 72°C for 30 sec. qPCR was conducted on a LightCycler 480 system (Roche Diagnostics) to quantify gene expression. The primers used in the present study were listed as follows: SDF2 forward, 5′-GGAGCTTGGCATCATGGACT-3′ and reverse, 5′-GGAGCTTCATAGCGTCTCCA-3′; CHOP forward, 5′-ATGAACGGCTCAAGCAGGAA-3′ and reverse, 5′-GGGAAAGGTGGGTAGTGTGG; GRP78 forward, 5′-TCAGGCCAAGCCCAATACAG-3′ and reverse, 5′-TCCACGGTAGTGAGAGCCTT-3′; XBP1s forward, 5′-TGCTGAGTCCGCAGCAGGTG-3′ and reverse, 5′-GCTGGCAGGCTCTGGGGAAG-3′; XBP1u forward, 5′-ACGGGACCCCTAAAGTTCTG-3′ and reverse, 5′-TGCACGTAGTCTGAGTGCTG-3′; and GAPDH forward, 5′-CCAGCAAGAGCACAAGAGGA-3′ and reverse, 5′-GGGGAGATTCAGTGTGGTGG-3′. The 2^−ΔΔCq method was used for quantification assay (16).

Human tumor and adjacent non-tumor tissue samples were obtained from patients with glioma during surgical procedures, with informed consent obtained from all participants. A total of 10 patients were enrolled in the present study, including 4 females and 6 males. The median age was 66 years (range, 53-67 years). Ethical approval for sample collection and study was granted by the institutional review board of the First Affiliated Hospital of Xi'an Jiaotong University (approval no. 2024SF-FAHX-212; Xi'an, China).

Additionally, the GEPIA database (http://gepia.cancer-pku.cn/) was used to analyze SDF2 mRNA expression levels across various tissue types and pathological conditions. Statistical significance was assessed using GEPIA's integrated analytical tools, providing insights into the potential role of SDF2 in disease pathology.

Cells from each group were collected, and the levels of MDA, reduced GSH and ATP were measured using the appropriate assay kits in strict accordance with the manufacturers' protocols. The lipid peroxidation level was assessed using an MDA Assay kit (cat. no. S0131S), ATP levels were quantified using an ATP Assay kit (cat. no. S0026), and GSH and oxidized glutathione (GSSG) levels were measured using a GSH and GSSG Assay kit (cat. no. S0053) (all from Beyotime Institute of Biotechnology).

The procedure was conducted using a BeyoClick EdU Cell Proliferation Kit (DAB method) (cat. no. C0085S; Beyotime Institute of Biotechnology). Briefly, cells were incubated in medium containing 10 μM EdU for 2 h. After EdU labeling, the medium was removed, and 1 ml of 4% paraformaldehyde (PFA; cat. no. P0099; Beyotime Institute of Biotechnology) was added to each well before incubation at room temperature for 15 min. The fixative was then removed, and the cells were washed three times with 1 ml of wash solution per well, with each wash lasting 5 min. Following the wash steps, 1 ml of PBS containing 0.3% Triton X-100 was added to each well, and the cells were incubated at room temperature for 15 min to permeabilize the cells. Next, 0.5 ml of Click reaction solution was added to each well, and the cells were incubated in the dark at room temperature for 30 min. The Click reaction solution was then aspirated, and the cells were washed three times with wash solution, with each wash lasting 5 min. Finally, the cells were mounted with antifade mounting medium containing DAPI (a final concentration of 1 μg/ml) and observed under a fluorescence microscope.

The bottom membrane of the Transwell insert (Corning, Inc.) was coated with Matrigel matrix (Corning, Inc.) at 37°C for 1 h. U87 and U251 glioma cells, which were cultured to ~85% confluence, were harvested using trypsinization and centrifuged at 1,000 × g for 15 min. The cell density was adjusted to 5×10³ cells/100 μl, and the cells were resuspended in serum-free medium to create a single-cell suspension. Transwell inserts (8 μm pore size) were placed in a 24-well plate, 600 μl of serum-containing medium was added to the lower chamber to act as a chemoattractant, and 200 μl of the treated cell suspension was added to the upper chamber. After 24 h of incubation, the inserts were removed, and the remaining medium was aspirated. The inserts were washed with PBS and fixed with 4% PFA for 30 min at room temperature, followed by staining with 0.1% crystal violet for 15 min. The cells that had migrated through the membrane were visualized and images were captured under an inverted light microscope (Keyence Corporation). The number of invasive cells was quantified using ImageJ software.

Cells in the logarithmic growth phase were harvested by trypsinization, collected in centrifuge tubes, and resuspended in PBS. After the cells were counted, 5×10⁵ cells were collected and washed with PBS. The cells were then resuspended in binding buffer from an Annexin V-PE/7-AAD Apoptosis Detection Kit (cat. no. E-CK-A216; Elabscience Biotechnology, Inc.) and incubated in the dark at room temperature for 20 min. Following incubation, the cells were washed, resuspended and analyzed for apoptosis by flow cytometry (CytoFLEX; Beckman Coulter, Inc.). The data were analyzed using FlowJo 10 (FlowJo LLC). The experiment was performed in triplicate for statistical reliability.

Intracellular ROS levels were measured by flow cytometry using a fluorescent probe-based ROS detection kit (Wuhan Servicebio Technology Co., Ltd.). Briefly, the cells were washed twice with PBS, centrifuged at 800 × g for 5 min to remove the supernatant at room temperature, and collected. DCFH-DA working solution was then added to each sample at a density of 1×10⁵ cells/ml, and the cells were incubated at 37°C in a CO₂ incubator protected from light for 30 min. After incubation, the cells were centrifuged at 800 × g for 5 min to remove the DCFH-DA solution, followed by washing with PBS 3 times to ensure the removal of excess probe. The cells were finally resuspended in PBS and immediately analyzed by flow cytometry for ROS detection.

For transfection, cells were seeded in 6-well plates at a density of 4×10⁵ cells per well 24 h prior to transfection to achieve optimal confluency. Small interfering RNA (siRNA, 5′-AAAUAGAACAUUUUGAAGGUG-3′) targeting the gene of interest and a non-targeting control siRNA (NC, 5′-UUCUCCGAACGUGUCACGUTT-3′) were obtained from Shanghai GenePharma Co., Ltd. Transfection was performed using HiPerFect Transfection Reagent (cat. no. 301705; Qiagen, Inc.) following the manufacturer's protocol. Briefly, 150 ng of siRNA (final concentration of 10 nM) was diluted in 100 μl of serum-free DMEM. Subsequently, 6 μl of HiPerFect Transfection Reagent was added to the diluted siRNA solution, mixed gently, and incubated at room temperature for 10 min to allow complex formation. The siRNA-lipid complexes were then added dropwise to each well containing cells in 2 ml of complete growth medium. Cells were assessed 48 h post-transfection.

U87 and U251 cells were adjusted to a density of 1,000 cells/μl and seeded into a 96-well plate. The cells were incubated at 37°C in a 5% CO₂ incubator for 24 h. After incubation, the medium was removed, and the wells were washed with PBS. Fresh medium (100 μl) was added to each well, followed by 10 μl of CCK-8 solution (Beijing Solarbio Science & Technology Co., Ltd.). The plate was then incubated for an additional 1 h. Absorbance was measured at 450 nm using a microplate reader to assess cell viability. The experiments included the following groups: Ad-NC and Ad-shSDF2. To investigate the mechanisms underlying the reduction in cell viability induced by Ad-shSDF2, several inhibitors were added to the Ad-shSDF2 group: z-VAD (20 μM), Nec-1 (10 μM), VX-765 (10 μM), Fer-1 (1 μM), and TTM (10 μM). Cell viability was calculated based on absorbance (A450) values, allowing for the assessment of each inhibitor's effect on Ad-shSDF2-induced cell viability reduction and identification of the specific cell death pathways involved.

The intracellular copper ion content was quantified using a Copper (Cu²⁺) Colorimetric Assay kit (Elabscience Biotechnology, Inc.). For the assay, 15 μl of each of the eight copper standards at different concentrations, as well as 15 μl of each test sample, were added to individual wells. Then, 230 μl of chromogenic working solution was added to each well. The plates were covered with sealing film and incubated at 37°C for 5 min. The optical density (OD) of each well was measured at 580 nm using a microplate reader.

The intracellular copper ion distribution was visualized using the fluorescent probe Coppersensor 1 (MedChemExpress). Cells were incubated with the probe at a final concentration of 5 μM in each well, protected from light, at 37°C for 30 min. After incubation, the cells were observed under a fluorescence microscope to assess the localization and intensity of copper ion fluorescence.

Mitochondrial ROS levels were detected using the specific fluorescent probe MitoTracker Red CMXRos (Beyotime Institute of Biotechnology). Once the cells reached the desired density, the culture medium was removed and replaced with MitoTracker Red CMXRos working solution at a final concentration of 1 μM. The cells were incubated at 37°C for 30 min. After incubation, the working solution was removed, and prewarmed fresh culture medium was added. Fluorescence was observed using a fluorescence microscope to assess the mitochondrial ROS levels.

U87 and U251 cells were seeded in six-well plates at a density of 5×10⁵ cells per well and incubated at 37°C with 5% CO₂ until adherence. After 24 h, the cells were fixed with 500 μl of 4% PFA for 30 min, followed by permeabilization with 500 μl of 0.1% Triton X-100 for 30 min. The cells were then blocked with IF blocking buffer for 1 h. After the blocking solution was removed, a mixture of primary antibodies against TOM20 and DLAT (Abcam) was added (50 μl per well), and the cells were incubated overnight at 4°C. The next day, the primary antibodies were removed, and the cells were incubated with the appropriate secondary antibodies (500 μl per well) at room temperature, protected from light, for 1 h. DAPI staining solution (a final concentration of 1 μg/ml) was then applied for 5 min to stain the nuclei. Images were captured and stored using an inverted fluorescence microscope.

For these experiments, a co-IP kit (cat. no. P0401; GENESEED; https://www.geneseed.com.cn/web/lxwm.html) was used. Briefly, protein extracts were prepared using RIPA lysis buffer (Beijing Solarbio Science & Technology Co., Ltd.). A specific antibody against GRP78 was added to the protein samples to allow binding with the target protein, and the samples were incubated at 4°C for several hours or overnight. 30 μl of protein A or G magnetic beads (50% concentration) were then added to bind the antibody-protein complexes, which were collected using magnetic separation. The beads were washed multiple times with lysis buffer to remove non-specifically bound proteins. The immunoprecipitated proteins were then eluted from the beads by heating or by adding an elution buffer. Finally, the eluted proteins were separated by SDS-PAGE and analyzed by western blotting to detect co-immunoprecipitated proteins. By normalizing the Co-IP results to GAPDH levels, any potential discrepancies in protein loading were accounted for and it was ensured that the observed decrease in ATP7A and ATP7B co-precipitation was specifically attributable to the reduced availability of GRP78 for interaction, rather than other experimental variations.

Adenoviral vectors were constructed by Vigene Biosciences for the overexpression and knockdown of the SDF2 or GRP78 proteins. Adenoviral vectors carrying negative control genes (Ad-NC, UUCUCCGAACGUGUCACGUTT) or shRNA sequences (Ad-sh-SDF2, UCUUUUUGCCCACUGAUAGGU) targeting SDF2 were designed to infect cells. Adenoviral vectors were added at a concentration of 10⁹ PFU/ml, with 20 μl viral suspension per well in six-well plates. After incubation at 37°C for 24 h, the viral-containing medium was removed, cells were washed with PBS, and fresh medium was added.

A total of 15 SPF-grade BALB/c nude male mice (6-8 weeks old, 18-22 g, Cyagen Biosciences; https://www.lascn.net/SupplyDemand/Site/Index.aspx?id=30) were randomly assigned into three experimental groups (Ad-NC, Ad-SDF2, and Ad-SDF2 + Ad-GRP78), with five mice per group, using a computer-generated randomization schedule. This sample size was selected based on precedent studies and accepted practices in glioma xenograft models, where comparable group sizes have been sufficient to detect statistically significant effects (17-19). Investigators responsible for data collection and outcome assessment were blinded to group allocation to minimize potential bias. All animals were acclimatized to the laboratory environment for one week prior to experimentation and housed under standard specific pathogen-free (SPF) conditions [22±2°C, 55±10% humidity, 12/12-h light/dark cycle (7:00-19:00), with ad libitum access to diet and sterile water]. Logarithmically growing U251 cells were collected from each treatment group. Cell pellets were resuspended in 200 μl of ice-cold PBS to maintain viability and kept on ice until injection. Each mouse was subcutaneously injected with 1×10⁶ cells into the inguinal region. Tumor implantation was designated as day 0. Tumor growth and general health were monitored every two days. Tumor volume (mm³) was measured using calipers and calculated according to the formula: volume=1/2 × longest diameter (mm) × [shortest diameter (mm)]². At the experimental endpoint, mice were euthanized by cervical dislocation. Subcutaneous tumors were excised, weighed, and processed for further analysis. The maximum tumor diameter observed was 8.2 mm, corresponding to a volume of 106.64 mm³. All animal procedures were approved by the Institutional Animal Care and Use Committee of the Animal Experimentation Center, Xi'an Jiaotong University (approval no. XJTUAE2023-1944; Xi'an, China).

Tumor tissue sections were subjected to Ki-67 IHC. In brief, the tissues (5 μm) were deparaffinized in xylene, rehydrated in a graded ethanol series, and treated with 3% hydrogen peroxide to inactivate endogenous peroxidase activity. Antigen retrieval was performed, followed by blocking with goat serum (cat. no. C0625; Beyotime Institute of Biotechnology). Ki-67 primary antibody (1:200; cat. no. AB2008; Beyotime Institute of Biotechnology) was applied, and the samples were incubated overnight at 4°C. The sections were then incubated with HRP-conjugated goat anti-rabbit IgG (cat. no. A0208; Beyotime Institute of Biotechnology) at 37°C for 30 min, followed by the application of DAB substrate for color development (30 sec). Hematoxylin counterstaining was performed for 1 min, and differentiation was achieved in 1% hydrochloric acid ethanol. The sections were then dehydrated through a graded ethanol series, cleared in xylene, and mounted with neutral resin. Images were captured using a light microscope. In each field, the number of Ki-67-positive tumor cell nuclei and the total number of tumor cell nuclei were manually counted. The proliferation index was then calculated as the percentage of Ki-67-positive cells among the total tumor cells in each field, and the average of these five fields was taken as the final proliferation index for the sample.

Statistical analyses were performed using GraphPad Prism software (version 10.0; Dotmatics). The data are presented as the mean ± standard deviations (SDs). For relative expression graphs, the control group was normalized to a fixed value (for example, 1 or 100%). For comparisons between two groups, an unpaired two-tailed Student's t-test was employed. For comparisons among multiple groups, one-way analysis of variance was conducted, followed by Tukey's post hoc test to assess pairwise differences between group means. P<0.05 was considered to indicate a statistically significant difference.

Bioinformatic predictions revealed an increase in SDF2 mRNA expression in glioma tissues compared with control tissues (Fig. 3A and B). Consistently, RT-qPCR analysis revealed significantly elevated SDF2 mRNA levels in tumor tissues relative to adjacent normal tissues (Fig. 3C). IF staining further demonstrated an increase in SDF2 fluorescence intensity in glioma tissues compared with control tissues (Fig. 3D). Western blot analysis confirmed higher SDF2 protein expression in glioma samples than in adjacent non-tumor tissues (Fig. 3E). These findings collectively suggest that SDF2 expression is upregulated in glioma, potentially implicating it in glioma pathogenesis.

U87 and U251 cells were transfected with Ad-SDF2 to overexpress SDF2. Following SDF2 overexpression, both the mRNA and protein levels of SDF2 were significantly elevated in U87, U251 and HMC3 cells (Fig. 4A-F). Additionally, cell viability assays demonstrated that, compared with the control, SDF2 overexpression enhanced the viability of U87 and U251 cells in a time-dependent manner (Fig. 4G and H), but the viability of HMC3 cells did not change (Fig. 4I). The EdU staining results further confirmed that SDF2 overexpression increased the proliferative capacity of U87 and U251 cells relative to that of the controls (Fig. 4J-M). Moreover, Transwell assays indicated that SDF2 overexpression promoted the migratory ability of U87 and U251 cells (Fig. 4N and O). These findings suggest that SDF2 plays a role in promoting the malignant proliferation and migration of glioma cells.

U87 and U251 cells were transfected with Ad-shSDF2 to achieve SDF2 knockdown. The mRNA and protein levels were decreased in U87 and U251 cells transfected with Ad-shSDF2 (Fig. 5A-D). Compared with the control, Ad-shSDF2 transfection resulted in a significant increase in the proportion of late apoptotic and necrotic cells in both the U87 and U251 lines (Fig. 5E and F). Additionally, ROS levels were significantly elevated following SDF2 knockdown in U87 and U251 cells, indicating increased oxidative stress (Fig. 5G and H). Interestingly, only the cuproptosis inhibitor TTM reversed the decrease in viability induced by Ad-shSDF2 in U87 and U251 cells (Fig. 5I and J). These results suggest that SDF2 knockdown induces oxidative stress, increases apoptosis, and may involve copper-dependent cell death pathways in glioma cells.

In both the U87 and U251 cell lines, SDF2 knockdown led to a significant increase in the intracellular copper ion concentration compared with that in the controls. This finding was confirmed through fluorescence staining (Fig. 6A) and further quantified using colorimetric assays (Fig. 6B and C). Additionally, the expression of the copper-exporting proteins ATP7A and ATP7B, as well as FDX and lipid-DLAT, was reduced in U87 and U251 cells following SDF2 knockdown, whereas DLAT protein levels remained unchanged (Fig. 6D and E). Furthermore, SDF2 knockdown resulted in decreased intracellular GSH levels (Fig. 6F) and increased ROS and MDA levels, indicating elevated oxidative stress and lipid peroxidation within the cells (Fig. 6G and H). These results suggest that SDF2 plays a role in maintaining copper homeostasis, antioxidant capacity, and oxidative stress regulation in glioma cells.

The effects of SDF2 on key proteins in the ER stress pathway were investigated. Compared with Ad-NC, Ad-shSDF2 increased GRP78 mRNA levels and the ratio of XBP1s/XBP1u in U87 and U251 cells, whereas CHOP mRNA levels remained unchanged (Fig. 7A and C). Additionally, Ad-shSDF2 increased GRP78 protein expression in both cell lines (Fig. 7B and D). To confirm the involvement of the proteasome degradation pathway in ATP7A and ATP7B downregulation, U87 and U251 cells were co-incubated with the proteasome inhibitor MG132. Compared with the control treatment, SDF2 knockdown reduced ATP7A and ATP7B expression, whereas MG132 treatment increased ATP7A and ATP7B expression (Fig. 7E-H). Notably, pretreatment with MG132 significantly reversed the Ad-shSDF2-induced reduction in ATP7A and ATP7B levels (Fig. 7E and F). Furthermore, compared with Ad-NC, Ad-shSDF2 increased DLAT aggregation within mitochondria; however, MG132 pretreatment reduced DLAT aggregation (Fig. 7G and H). These findings indicate that SDF2 knockdown promotes ATP7A and ATP7B degradation through a GRP78-mediated ER-associated degradation pathway, with potential downstream effects on mitochondrial DLAT accumulation.

To investigate the role of GRP78 in this process, si-GRP78 was utilized to specifically knock down GRP78 expression. RT-qPCR analysis confirmed a significant reduction in GRP78 mRNA levels in U87 and U251 cells following si-GRP78 transfection (Fig. 8A and D). Western blot analysis further verified that si-GRP78 effectively suppressed GRP78 protein expression in both glioma cell lines (Fig. 8B and E). Importantly, GRP78 silencing also blocked the Ad-shSDF2-induced downregulation of ATP7A and ATP7B in U87 and U251 cells (Fig. 8B and E). Co-IP results demonstrated that knockdown of GRP78 led to a reduction in the amount of ATP7A and ATP7B co-precipitated with GRP78. This decrease was observed after normalizing the Co-IP signals to GAPDH levels in the corresponding Input samples (Fig. 8C and F). These findings suggest that GRP78 may play a role in regulating ATP7A and ATP7B stability in glioma cells. Additionally, GRP78 silencing partially reversed the Ad-shSDF2-induced increase in the intracellular Cu²⁺ and mito-ROS levels in U87 and U251 cells (Fig. 8G and H). These findings indicate that SDF2 suppresses ATP7A and ATP7B expression through a GRP78-dependent mechanism, impacting copper ion homeostasis and mitochondrial oxidative stress in glioma cells.

Compared with Ad-NC, Ad-SDF2 significantly increased tumor volume, and weight in a nude mouse model (Fig. 9A-C). IHC analysis revealed that Ad-SDF2 increased the Ki-67 positivity rate in tumors, whereas the co-expression of Ad-GRP78 reversed this increase, suggesting an impact on cell proliferation (Fig. 9D). Further analysis revealed that Ad-SDF2 increased the GSH level in tumor tissues, which was effectively reversed by Ad-GRP78 (Fig. 9E). Similarly, Ad-SDF2 overexpression led to decreased MDA levels, whereas co-expression of Ad-GRP78 partially restored MDA levels (Fig. 9F). Moreover, Ad-SDF2 increased ATP levels and reduced Cu²⁺ concentrations in tumor tissues, while co-expression with Ad-GRP78 restored both parameters (Fig. 9G and H). Additionally, Ad-SDF2 overexpression suppressed GRP78 expression while increasing the expression of copper transporters ATP7A and ATP7B. Co-expression of Ad-GRP78 significantly reversed these changes (Fig. 9I). These findings indicate that GRP78 overexpression can counteract the effects of SDF2-induced changes in tumor growth and metabolism, highlighting the regulatory role of GRP78 in modulating SDF2-mediated changes in glioma growth and oxidative stress.