Retinoic Acid Receptor-related Orphan Receptor α Drives Glucose Reprogramming and Mitochondrial Rescue Mitigate Subarachnoid Hemorrhage-Induced Early Brain Injury

Abstract

Aims:

Subarachnoid hemorrhage (SAH) is a devastating cerebrovascular event characterized by early brain injury (EBI) within 72 h that is driven by oxidative stress, mitochondrial dysfunction, and metabolic collapse. The retinoic acid receptor-related orphan receptor alpha (RORα) is a nuclear receptor implicated in metabolic and inflammatory regulation, but it has not been studied in SAH. We aimed to determine whether RORα confers neuroprotection after SAH and to elucidate its underlying mechanisms.

Methods and Results:

We used mouse SAH models and primary cortical neurons to assess the RORα expression, functional outcomes, and metabolic changes. The RORα expression was markedly reduced post-SAH. Genetic knockdown or deficiency (staggerer mice) exacerbated neuronal apoptosis, neuroinflammation, and behavioral deficits. Conversely, pharmacological activation with SR1078 significantly improved neurological scores, preserved neuronal morphology, and reduced oxidative stress. RORα overexpression or SR1078 treatment enhanced neuronal viability in vitro under hemoglobin-induced stress. Transcriptomic and epigenomic profiling revealed that RORα directly regulated glucose-6-phosphate dehydrogenase and α subunit of peroxisome proliferator activated receptor-γ coactivator-1. This promoted pentose phosphate pathway flux and mitochondrial biogenesis. A metabolic flux analysis confirmed increased nicotinamide adenine dinucleotide phosphate hydrogen and glutathione synthesis, reduced reactive oxygen species accumulation, and an improved oxygen consumption rate and spare respiratory capacity. All of these results indicated a shift toward oxidative phosphorylation and enhanced bioenergetics.

Innovation and Conclusion:

We are the first to demonstrate that RORα activation reprogrammed neuronal glucose metabolism and strengthened antioxidant defenses to mitigate SAH-induced EBI. The targeting of RORα could represent a promising therapeutic strategy for stroke-related metabolic failure and oxidative stress. Future work should explore the translational potential in clinical settings. Antioxid. Redox Signal. 44, 275–291.

Graphical Abstract

Keywords

subarachnoid hemorrhage nuclear receptors oxidative stress metabolic remodeling pentose phosphate pathway neuroprotection

Introduction

Spontaneous subarachnoid hemorrhage (SAH) is currently the third most prevalent type of stroke, and ruptured intracranial aneurysms account for 50%–85% of its primary causes (Gu et al., 2025). Pathophysiological mechanisms after the hemorrhage include blood–brain barrier disruption, cerebral edema formation, microvascular dysfunction, inflammatory response activation, and cell apoptosis (Kusaka et al., 2004). These secondary brain injuries occur within 72 h and are referred to as early brain injury (EBI). These injuries are critical factors that lead to neurological dysfunction, delayed cerebral ischemia, and other adverse outcomes (such as late mortality and disability) in surviving patients (Ahn et al., 2018; de Oliveira Manoel et al., 2016). There do exist current clinical treatments that target the primary insult, but there are no neuroprotective strategies aimed at EBI that have yielded satisfactory outcomes. Glucose metabolic disorders are major EBI contributing factors that result in impaired anabolism and ATP depletion. However, the production levels of antioxidants, such as glutathione, taurine, nicotinamide adenine dinucleotide phosphate hydrogen (NADPH), and NAD+, are significantly reduced, further exacerbating cellular vulnerability. The rescue of cerebral metabolic function using early interventions represents a promising therapeutic strategy (Cueto et al., 2024).

Innovation

This study was the first to demonstrate that RORα activation reprogramed neuronal metabolism to mitigate SAH-induced injury. We demonstrated that RORα effectively safeguarded neuronal survival and function following SAH. Mechanistically, RORα enhanced transcriptional activity to stimulate neuronal glucose uptake and the PPP, thereby bolstering reducing equivalents to counteract oxidative stress. Concurrently, RORα redirected glucose flux toward OXPHOS, and rewiring augments basal and maximal respiratory capacity. RORα-mediated neuroprotection may extend to other forms of cerebrovascular injury, and it could potentially offer a generalizable therapeutic strategy for early brain injury following stroke.

Nuclear receptors (NRs) belong to the superfamily of ligand-regulated transcription factors. Approximately half of the NRs have well-defined natural ligands, whereas the remaining receptors are classified as orphan NRs due to the absence of characteristic ligands (Hummasti and Tontonoz, 2008). The retinoic acid receptor-related orphan receptor α (RORα) is a profound metabolic regulator, as evidenced in recent studies. RORα exerts protective effects in multiple organs by modulating cellular autophagy and maintaining mitochondrial metabolism under pathological conditions such as myocardial ischemia and renal ischemia–reperfusion injury (He et al., 2016; Zhao et al., 2017). Mechanistically, these regulatory effects depend on the ability of RORα to act as a gene transcription repressor or activator by recruiting co-repressors and coactivators. Several studies have indicated that RORα alleviates neuroinflammation (Yue et al., 2020), but whether RORα plays a protective role in brain protection, particularly in early brain injury, and the underlying mechanisms require further investigations.

We utilize staggerer mice, RORα-overexpressing neurons, and various behavioral and molecular biology experiments to clarify the RORα expression after SAH and its impact on prognoses. In addition, we explore the RORα translational potential as a therapeutic target through the in vitro and in vivo applications of the agonist SR1078. Chromatin immunoprecipitation sequencing (ChIP-Seq) and transcriptome sequencing (RNA-Seq) were then used in vitro to explore the RORα protective mechanism. We then demonstrated that RORα transcriptionally activated glucose-6-phosphate dehydrogenase (G6PD) and the α subunit of peroxisome proliferator activated receptor-γ coactivator-1 (PGC1-α) to enhance glucose uptake and subsequently activate pentose phosphate pathway (PPP) flux, promoting mitochondrial biogenesis. RORα activation effectively improved cell survival and neurological function. These results indicated that RORα is a promising target for neuroprotection. We provided novel insights in this study for RORα theoretical research for SAH treatment.

Results

Expression and localization of RORα after SAH

First, we induced SAH in wild-type (WT) C57BL/6J mice. We then assessed the mRNA and protein levels in the cerebral cortexes to characterize RORα expressions in the brains under normal and pathological conditions. The Western blot and quantitative real-time polymerase chain reaction (RT-PCR) analyses revealed significantly decreased RORα expressions 24 h post-SAH conditions compared with the sham-operated controls (Fig. 1A,B). Immunofluorescence staining demonstrated that RORα was primarily localized in neuronal nuclei and secondarily in astrocytes, with negligible colocalization in microglia (Fig. 1C,D, Supplementary Fig. S1A–C).

FIG. 1.

Expression and cellular localization of RORα in the cerebral cortex post-SAH. (A) Western blot analysis of RORα protein levels in the cerebral cortex; β-actin was used as loading control (n = 5). (B) Relative mRNA expression of RORα measured by qRT-PCR 24 h after SAH. Data are shown as the mean ± SEM (n = 6). (C, D) Immunofluorescence staining and analysis showing nuclear localization of RORα in NeuN-positive neurons. Scale bar = 20 μm (n = 6). Sham, sham surgery mice; RORα, retinoic acid receptor-related orphan receptor alpha; SAH, subarachnoid hemorrhage; qRT-PCR, real-time quantitative polymerase chain reaction; SEM, standard error of mean. **p < 0.01, ***p < 0.001.

RORα deficiency exacerbated neurological deficits

We also established SAH models using staggerer (sg/sg) mice with natural RORα deficiencies to investigate the functional relevance of RORα. The sg/sg mice showed significantly lower scores in the Garcia, foot fault, and rotarod tests during the acute (24 h) and recovery phases compared with the WT controls. These results indicated more severe neurological impairment (Supplementary Fig. S2A–C, Fig. 2A).

FIG. 2.

Neurological function is impaired in RORα-deficient mice. (A) A timeline illustrating behavioral testing is presented. (B) Confirmation of RORα knockdown(shRORα) via Western blot analysis. Scramble shRNA (SC) was used as control (n = 6). (C) The modified Garcia Score test was used to assess the neurobehavioral impairment in RORα knockdown mice (shRORα vs. SC). (D) Foot fault test (shRORα vs. SC). The percentage of incorrect steps made by the left forelimb out of all steps taken by the mice was calculated. (E) Rotarod performance (shRORα vs. SC). The latency to fall from a rotating drum after placement of the mice was measured. (F–I) Morris water maze evaluation of spatial learning and memory (shRORα vs. SC). Data shown as mean ± SEM (n = 6); ns: p > 0.05, *p < 0.05, **p < 0.01.

Due to these cerebellar developmental defects in sg/sg mice, we further generated neuron-specific RORα knockdown mice via intravenous delivery of lentiviral shRNA (Fig. 3A). The knockdown efficiency was confirmed using Western blot (Fig. 2B). The RORα-silenced mice exhibited significantly worse motor performance and spatial learning/memory deficits in the Morris water maze tests compared with those of the control mice (Fig. 2C–I, Supplementary Fig. S2E–G). This result supported a neuron-intrinsic role of RORα in mitigating SAH-induced neurobehavioral impairments.

FIG. 3.

RORα activation by SR1078 improves functional outcomes. (A) A schematic of RORα knockdown, drug administration, and behavioral testing is presented. (B–G) Behavioral performance of Garcia, foot fault, and rotarod tests showed that SR1078 treatment improved the motor performance (B–D), but the knockdown of RORα in neurons eliminated the effect (E–G) (SC-SAH-SR1078, SAH mice injected with scramble RNA and SR1078; sh-SAH-SR1078, SAH mice injected with RORα-shRNA and SR1078; sh-SAH-DMSO, SAH mice injected with RORα-shRNA and DMSO; SC-SAH-DMSO, SAH mice injected with scramble RNA and DMSO). (H–K) Water maze test showed that SR1078 treatment improved the spatial learning and memory SC mice but not in RORα-deficient mice (n = 6). Data shown as mean ± SEM; ns: p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001. SC, scramble shRNA.

RORα activation improved neurological outcomes

SR1078 is a common synthetic RORα/γ agonist. The RORγ expression in the brain cortical tissue or primary neurons of the mice was shown to be nearly indetectable (Supplementary Fig. S3A,B); hence, SR1078 was used to test the therapeutic effect of targeting RORα. All mice received daily intraperitoneal injections (10 mg/kg) from 2 days pre-SAH until day 14 (Fig. 3A). The SR1078-treated mice showed significant improvements in Garcia scores, foot fault rates, and rotarod performances compared with the scores of the vehicle-treated controls (Fig. 3B–D). However, when RORα was knocked down, the SR1078 therapeutic efficacy was eliminated (Fig. 3E–G). A central nervous system permeability RORγ inhibitor, GSK805, was then combined with SR1078 to further eliminate the influence of RORγ. The results showed that GSK805 did not affect the behavior of SAH mice, regardless of whether SR1078 was used in combination or not (Supplementary Fig. S3C–E). A Morris water maze analysis further revealed that SR1078 enhanced spatial learning and memory performance in WT mice but not RORα knockdown mice (Fig. 3H–K). These data indicated that the SR1078 neuroprotective effect is RORα dependent.

SR1078 treatment ameliorated neuropathy

Nissl staining was conducted 24 h post-SAH, and it showed pronounced neuronal atrophy and cytoplasmic disintegration in the RORα-deficient and SAH mice. However, SR1078 partially preserved the cortical neuron integrity (Fig. 4A,B). Fluoro-jade C (FJC) staining corroborated the increased neuronal degeneration in the sg/sg and SAH groups, and this degeneration was significantly reduced by the SR1078 treatment (Fig. 4C,D).

FIG. 4.

RORα activation ameliorates neuronal death and inflammation. (A) Nissl staining was used to assess neuronal morphology in the cortex, scale bar = 20 μm. (B) The statistical analysis of abnormal Nissl bodies in the cortical region was conducted, n = 6. (C) FJC staining was used to show the degenerated neurons, Scale bar = 50 μm. (D) The quantification of FJC-positive cells (n = 6). (E–G) mRNA levels of IL-1β, IL-6, and TNF-α in cortical tissue measured by qRT-PCR (n = 6). WT-SAH, SAH wild-type mice; SR1078, SAH wild-type mice with SR1078 injection; WT-Sham, wild-type mice with sham surgery; sh-SAH, SAH mice injected with RORα-shRNA; Sg/Sg-SAH, SAH Staggerer mice. Staggerer (sg/sg) mice are a genetic model with a mutation in the RORα gene, resulting in decreased and dysfunctional RORα expression in all tissues; FJC, Fluoro-Jade C staining; IL-6, interleukin 6; TNF-α, tumor necrosis factor-alpha; IL-1β, interleukin 1 beta. Data shown as mean ± SEM; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

The qRT-PCR showed significantly elevated inflammatory cytokine levels (IL-1β, IL-6, TNF-α) in the cortexes of the sg/sg mice and reduced levels in the SR1078-treated mice (Fig. 4E–G). These results suggested that RORα deficiency exacerbated neuroinflammation and neuronal loss, whereas its activation exerted protective effects.

RORα overexpression enhanced neuronal transcriptional activity and metabolism

The downstream regulatory effects of RORα were explored using primary mouse neurons transduced with a RORα-overexpressing lentivirus or treated with SR1078 following oxyhemoglobin (OxyHb) stimulation (Fig. 5A). The RORα overexpression was confirmed using Western blot and qRT-PCR (Fig. 5B, Supplementary Fig. S4A). Consequently, improved neuronal viabilities with decreased lactate dehydrogenase (LDH) release were observed in the RORα-overexpressing cells compared with those of the OxyHb-exposed controls (Fig. 5C,D). We then treated the RORα-knockdown cells with SR1078 and found that the RORα knockdown nullified the SR1078 therapeutic effect (Supplementary Fig. S4B, C). This step further clarified the pharmacological activation of RORα.

FIG. 5.

RORα promotes neuronal transcriptional activity and metabolic reprogramming. (A) Experimental setup for primary neuron transfection or treatment. (B) Confirmation of RORα overexpression in neurons by Western blot analysis (n = 3). (C) CCK-8 cell viability assay (n = 3). (D) LDH release assay in neuron coculture medium (n = 3). (E) ChIP-seq peak enrichment near transcription start sites. (F) Peak annotation showed the main peaks were in promoter areas (G) Functional KEGG analysis of RORα-activated genes. CCK-8, Cell Counting Kit-8; LDH, Lactate Dehydrogenase; ChIP, Chromatin Immunoprecipitation; KEGG, Kyoto Encyclopedia of Genes and Genomes; TSSs, transcription start sites. Data shown as mean ± SEM; **p < 0.01, ***p < 0.001.

The ChIP-seq using enhancer/promoter markers (H3K27ac, H3K4me3) revealed enrichment at the transcription start sites (Fig. 5E), and the peak annotation showed that the primary peaks were in promoter areas (1 kb) (Fig. 5F, Supplementary Fig. S4D). This result was consistent with enhanced transcriptional activity under RORα overexpression. Gene Ontology and the Kyoto Encyclopedia of Genes and Genomes (KEGG) analyses identified that the enrichment primarily occurred in metabolic pathways (Fig. 5G; Supplementary Fig. S4E–G). These results indicated that RORα regulated neuronal metabolism through transcriptional activation.

RORα promoted neuronal glucose metabolism

The RNA-seq of the RORα-overexpressing neurons identified 1001 differentially expressed genes (DEGs) (390 upregulated and 611 downregulated) under OxyHb stimulation. The upregulated genes included glycolytic and PPP and tricarboxylic acid (TCA) cycle-related genes such as hexokinase 1 (HK1) and G6PD (Fig. 6A). The KEGG enrichment analysis indicated upregulation of metabolic and protein biosynthesis pathways and downregulation of inflammatory and calcium signaling pathways (Fig. 6B,C).

FIG. 6.

RORα modulates glucose metabolic pathways under stress. (A) The heat map of differentially expressed genes under RORα overexpression. (B, C) KEGG enrichment of up- and downregulated pathways. (D) qRT-PCR validation of glucose metabolism genes (n = 3). Data shown as mean ± SEM; ^#p ≥ 0.05, *p < 0.05, **p < 0.01, ***p < 0.001.

The DEGs were categorized into three functional groups and examined using qRT-PCR to further validate the effect of RORα on neuronal glucose metabolism. The results demonstrated that the expressions of several key metabolic genes, such as G6PD, glucose transporter (GLUT)4, and HK1, were upregulated in the RORα-overexpressing neurons (Fig. 6D). These data suggested that RORα modulated carbon metabolism and enhanced neuronal glucose utilization under stress.

RORα enhanced the neuronal antioxidant capacity

GLUT3 and GLUT4 are primarily subtypes that are expressed in neurons (Koepsell, 2020). First of all, Western blotting revealed the selective upregulation of GLUT4 (but not GLUT3) in the RORα-overexpressing neurons (Fig. 7A, Supplementary Fig. S5A). This was accompanied by enhanced glucose uptake (Fig. 7B, Supplementary Fig. S5B) and HK1 expression (Fig. 7A, Supplementary Fig. S5A). The NADPH and glutathione glycosylase system (GSH) levels were significantly elevated following RORα activation, and this result indicated enhanced PPP flux and antioxidant capacity (Fig. 7C,D, Supplementary Fig. S5C,D). As a result, the reactive oxygen species (ROS) levels were significantly reduced in neurons with RORα activation (Fig. 7E,F, Supplementary Fig. S5E,F), indicating improved redox homeostasis. A metabolic flux analysis was then conducted to further elucidate the glucose metabolism differences induced by RORα overexpression. It was observed that the RORα-overexpressing cells channeled more glucose into the PPP. This resulted in increased GSH production. Regarding ATP/ADP/AMP labeling, a significant increase in the five-carbon labeling of ribose from ribose-5-phosphate was detected in the drug-treated group. This indicated that the RORα overexpression enhanced nucleic acid synthesis and energy production via the oxidative PPP (Fig. 7G). A TCA cycle comparison between the two groups revealed that the RORα-overexpressing cells exhibited a significant increase in the labeled flux from glucose to TCA cycle metabolites. In addition, there was a marked elevation in the total abundance of metabolites within the TCA pathway (Fig. 7H). These findings suggested that RORα overexpression substantially enhanced glucose conversion into the TCA cycle.

FIG. 7.

RORα enhances antioxidant capacity via PPP activation. (A) Western blot of Glut3, 4 and HK1 expression under RORα overexpression. (B) Glucose uptake assay. (C) NADPH levels. (D) Intracellular GSH content. (E, F) ROS levels analyzed by flow cytometry. (G) ¹³C-Flux ratio of PPP metabolites from the labeled U-¹³C–Glucose. (H) ¹³C-Flux ratio of TCA metabolites from the labeled ¹³C–Glucose. Glut3,4, glucose transporter3, 4; HK1, hexokinase1; NADPH, Nicotinamide Adenine Dinucleotide Phosphate Hydrogen; GSH, glutathione glycosylase system; GSSG, Oxidized Glutathione; ROS, reactive oxygen species. Data shown as mean ± SEM (n = 3); ns: p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

RORα improved mitochondrial bioenergetics

The neurons that overexpressed RORα enhanced the protein levels of mitochondrial complexes I–V (NDUFB8, SDHB, UQCRC2, MTCO1, and ATP5A) (Fig. 8A, Supplementary Fig. S6A). A seahorse analysis revealed that OxyHb reduced basal respiration and ATP production, and these were restored by RORα activation (Fig. 8C–G, Supplementary Fig. S6C–G). RORα also improved the maximal respiration and spare capacity, indicating increased mitochondrial resilience.

FIG. 8.

RORα maintains mitochondrial function and respiratory capacity. (A) Western blot analyzed the expression of mitochondrial complexes I–V under RORα overexpression. (B) Representative ECAR trace of neurons successively exposed to 10 mM glucose, 6 μM oligomycin A, and 100 mM 2-DG. (C) Representative OCR trace of RORα overexpression cells successively exposed to 6 μM oligomycin A, 4 μM FCCP, 2 μM antimycin A, and 2 μM rotenone. (D–G) Bioenergetics profiling: ATP production, maximal respiration capacity (MRC), and spare respiratory capacity (SRC). (H, I) Mitochondrial membrane potential assessed by JC-1 fluorescence. ECAR, extracellular acidification; OCR, oxygen consumption rate; Oligo, oligomycin A; 2-DG, 2-Deoxy-D-glucose; FCCP, Carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone; Rota, rotenone; AA, antimycin A; JC-1, 5,5′,6,6′-Tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide. Data shown as mean ± SEM; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

In addition, RORα reduced extracellular acidification (ECAR). This result suggested a shift from glycolysis to oxidative phosphorylation (OXPHOS) (Fig. 8B, Supplementary Fig. S6B). Flow cytometry confirmed that the mitochondrial membrane potential (MMP) was preserved by RORα, and it counteracted OxyHb-induced depolarization (Fig. 8H,I, Supplementary Fig. S6H,I). Collectively, RORα enhanced mitochondrial function and antioxidant defenses, thereby protecting neurons from ischemic stress.

RORα improved the gene expressions of G6PD and PGC1-α through the recruitment of promoters

It has been reported that RORα improves cell metabolism through the transcriptional regulation of G6PD, PGC1-α, or GA-binding protein (GABP) (Kim et al., 2017; Kim et al., 2024; Wang et al., 2024). In addition, estrogen-related receptor (ERR)γ is another potential target in metabolism affected by RORα (Han et al., 2016). PGC1-α-mitochondrial transcription factor A (TFAM) is a core regulatory axis in mitochondrial biogenesis(Dominy and Puigserver, 2013). The mRNA expressions of G6PD, PGC1-α, GABP, TFAM, and ERRγ were detected to clarify the direct RORα regulatory targets in neurons. The results showed that G6PD, PGC1-α, and TFAM instead of ERRγ or GABP were elevated by RORα activation (Fig. 9A). Subsequently, a Chip assay was used to explore the deep mechanism. We found that RORα was recruited to the promoters of the G6PD and PGC1-α genes (Fig. 9B). These results suggested the existence of an RORα-G6PD/PGC1-α-TFAM regulatory axis. To further clarify the effect of G6PD and PGC1-α in the RORα regulatory axis, G6PD and PGC1-α were knocked down, respectively, in RORα overexpressing cells. A Cell Counting Kit-8 (CCK8) assay and an LDH release assay showed that the G6PD or PGC1-α knockdown partially alleviated the therapeutic effect caused by RORα overexpression (Fig. 9C,D). Taken together, these results demonstrated that RORα was recruited to the promoters of the G6PD and PGC1-α genes to strengthen the transcription ability of RORα.

FIG. 9.

RORα recruits the promoters of G6PD and PGC1-α. (A) QRT-PCR showing expression of metabolic regulating genes. (B) RORα was recruited to G6PD and PGC1-α gene promoter in neuron cells through Chip assay. (C) Respectively knocked down the expression of G6PD and PGC1α in RORα-OV cells, cell viability was assessed by CCK-8 assay. (D) LDH release in the neuron coculture medium was measured. Data shown as mean ± SEM (n = 3); ns: p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. G6PD, glucose-6-phosphate dehydrogenase; PGC1-α, α subunit of peroxisome proliferator activated receptor-γ coactivator-1.

Discussion

Retinoic acid-related orphan receptors are increasingly recognized for their role in regulating homeostatic and pathological processes. RORα, in particular, influences cellular metabolism, immune responses, and neurodevelopment. We demonstrated in this study that RORα expression was significantly reduced following subarachnoid hemorrhage, and its deficiency exacerbated neurological deficits and worsened long-term prognoses. SR1078 is a synthetic agonist of RORα and RORγ, and it has been used to validate RORα-mediated transcriptional activation in various tissues (Kallen et al., 2004; Kallen et al., 2002). In our study, SR1078 administration improved neurological function and reduced neuronal degeneration in WT mice but failed to confer benefits in sg/sg or RORα knockdown mice. These results confirmed that SR1078 exerted its neuroprotective effects via RORα activation. Importantly, SR1078’s inability to rescue the function in the RORα-deficient models provided strong causal evidence that RORα is indispensable for its therapeutic action.

An increasing amount of evidence has indicated that mitochondrial dysfunction occurs following SAH, and this has been found to be associated with EBI and negative neurological outcomes (Chou et al., 2017). PGC-1α is a key regulator of mitochondrial biogenesis and promotes TFAM expression through multiple pathways that involve nuclear respiratory factor 1 (NRF1) and GABP (also known as the nuclear respiratory factor 2) (Dominy and Puigserver, 2013). Resveratrol enhances mitochondrial biogenesis via the PGC-1α pathway, thereby reducing oxidative stress and mitochondrial-dependent apoptosis induced by mitochondrial dysfunction after SAH (Zhou et al., 2021). Edaravone is a nonselective antioxidant that has demonstrated significant efficacy in rat SAH models (Gao et al., 2009). In addition, a randomized controlled trial that included 91 patients revealed that edaravone reduced the incidence of neurological deficits and poor outcomes associated with carotid artery syndrome following SAH (Munakata et al., 2009). We found that the neuroprotective effects of RORα were multifaceted. It drove a shift in intracellular glucose metabolism through the transcriptional activation of PGC-1α and G6PD and thus promoted PPP flux and mitochondrial biogenesis. These activities comprehensively improved the cellular redox balance and respiratory capacity.

ERRs are another class of profound mitochondrial metabolism modulators. Previous studies have shown that ERRα/β expression remains unaltered after SAH, whereas ERRγ exhibits dynamic temporal changes post-SAH, suggesting its therapeutic potential (Guo et al., 2023). Notably, RORα has been reported to antagonize ERRγ function in the liver (Han et al., 2016). Although our results confirmed that the ERRγ expression was unaffected by RORα overexpression, the issue of whether sex differences influence the therapeutic efficacy of RORα warrants further investigation.

Kelch-like ECH-associated protein 1/Nrf2 signaling is an essential cellular defense mechanism that protects against ROS. Nuclear Nrf2 binds to antioxidant response elements in the HO-1, MnSOD, and NFR-1 gene promoters, thus amplifying the initial signal and driving the transcription of downstream vitagenes (Calabrese et al., 2010). Previous studies have shown that RORα has a role in antioxidant defenses across various tissues. In neurons, RORα1 overexpression upregulates antioxidant enzymes such as peroxiredoxin 6 and glutathione peroxidase (GPx) in neurons, leading to a reduction in ROS-induced tissue damage (Boukhtouche et al., 2006). RORα activation in the liver has been shown to reduce oxidative damage in nonalcoholic steatohepatitis by inducing superoxide dismutase 2 and GPx1(Han et al., 2014; Ou et al., 2013). These results indicate that the mechanism by which RORα regulates redox homeostasis differs from that of NRF2, although they share some common downstream targets.

RORα contains natural ligands, and these are very crucial metabolic substances in the body. However, they are unsuitable for direct use as medications (Kim et al., 2023). Synthetic agonists, including neoruscogenin, nobiletin, and SR1078, can affect multiple organ systems (Helleboid et al., 2014; Kim et al., 2023; Tanaka et al., 2004). Neoruscogenin can increase the expression of liver RORα target genes, while nobiletin has been studied for its impact on the myocardium, skin system, adipose tissue, liver, nervous system, and malignant tumors. Nobiletin can reduce inflammation and cytokine expression in the nervous systems of patients with neurodegenerative diseases, including those with negatively affected cortexes, such as in Alzheimer’s (Wirianto et al., 2022). In addition, gene therapy targeting that which targets RORα has been explored to treat retinal degeneration (Akula et al., 2024), and CRISPR-based strategies have been proposed to modulate ROR pathways with greater precision (Sun et al., 2016). In our study, RORα activation via SR1078 was shown to provide robust neuroprotection against SAH-induced early brain injury. It is worth noting that although RORα functions are gradually being discovered, current therapeutic strategies that target RORα have not yet entered clinical trials. RORα has widespread expressions and functional roles in multiple organs, and future studies should thoroughly evaluate its in vivo safety and pharmacokinetic profiles to comprehensively assess the translational feasibility in clinical settings (Guntermann et al., 2017).

In conclusion, our findings identified RORα as a critical neuroprotective factor following subarachnoid hemorrhage. RORα promoted neuronal survival by reprogramming glucose metabolism, enhancing mitochondrial bioenergetics, and suppressing oxidative stress. The synthetic agonist SR1078, conferred neuroprotection in a RORα-dependent manner, and this result supported its potential as a therapeutic development candidate. Further research is required to assess whether RORα-mediated pathways offer a universal protection mechanism across diverse stroke and brain injury models. An exploration of combinatorial therapeutic strategies that target RORα and downstream metabolic pathways may yield novel approaches to mitigate early brain injury and improve long-term neurological outcomes.

Methods

Animals

Male WT C57BL/6J mice (6–8 weeks old, 22–28 g) were obtained from the Shanghai Laboratory Animal Center (Chinese Academy of Sciences). RORα-deficient staggerer (sg/sg) mice were kindly provided by Professor Jun Bu (Renji Hospital, Shanghai Jiao Tong University). All animals were housed in a specific pathogen-free environment under a 12-h light/12-h dark cycle, with free access to autoclaved food and sterile water. Cages were maintained under standardized hygienic conditions. All animal procedures conformed to the ARRIVE guidelines and the ethical guidelines of the Animal Welfare and Ethics Committee of Shanghai Rat & Mouse Biotech Co., Ltd. (Approval No. 2022110329).

Establishment of mouse SAH model

The SAH model was induced using an endovascular perforation method as previously described (Xiao et al., 2021). Mice were anesthetized with 4% isoflurane and maintained on a rodent ventilator (Matrx, Midmark). A filament (702234PK5Re, Doccol) was inserted via the left external carotid artery into the internal carotid artery and advanced to perforate the bifurcation of the middle cerebral artery and anterior cerebral artery. Sham-operated mice underwent the same procedure without vessel perforation. SAH severity was assessed 24 h postinduction using a validated grading scale and hemoglobin content quantification. Mice with SAH grades ≤7 were excluded.

Drug administration

SR1078 was prepared at 25 mg/mL by dissolving 1 mg powder in 40 µL DMSO and then diluted 1:10 in corn oil to yield a final concentration of 2.5 mg/mL. Mice received 10 mg/kg SR1078 intraperitoneally. Vehicle controls received an equivalent volume of DMSO per corn oil.

Neurological scoring

Neurobehavioral testing was conducted to evaluate neurological function before surgery and on days 1, 3, 7, 14, and 25 following SAH induction. A battery of tests was used, including the modified Garcia score, foot fault test, rotarod test, and Morris water maze.

The modified Garcia test was used to assess general neurological function. It comprised seven domains: spontaneous activity, symmetry of limb movement, forelimb outstretching, climbing, body proprioception, vibrissae response, and lateral turning. Each domain was scored from 0 to 3, with a maximum composite score of 21.

The foot fault test was applied to evaluate forelimb coordination and placement. Mice were placed on a metal grid (1.5 cm × 1.5 cm openings) and allowed to walk freely for 1 min. Any misplacement or slipping of the left forepaw into the grid openings was recorded as a foot fault. The foot fault percentage was calculated relative to the total number of steps.

Motor coordination and balance were evaluated using the rotarod test. Mice were placed on an accelerating rod rotating from 5 to 40 rpm over 300 s. Each mouse underwent three trials per time point, and the average latency to fall was recorded.

Cognitive function was evaluated by the Morris water maze test between days 21 and 25 post-SAH. During the place navigation phase, mice were trained to locate a hidden platform submerged below the water surface. Escape latency (time to reach the platform) was recorded. In the spatial probe trial conducted 24 h later, the platform was removed, and the time spent in the target quadrant and several platform area crossings were recorded to assess spatial memory.

Immunofluorescence staining

Mice were perfused with PBS and 4% paraformaldehyde. Brains were postfixed, dehydrated in 30% sucrose, embedded in OCT, cryo-sectioned at 40–50 µm, and stored at 4°C. Sections were incubated with primary antibodies and corresponding fluorescent secondary antibodies. Images were acquired from the left temporal cortex using a Leica SP8 confocal microscope. Antibodies: anti-RORα (1:200, ab278108, abcam), anti-Iba1 (1:1000, 019–19741, Wako), Donkey anti-rabbit IgG Secondary Antibody, Alexa Fluor™ 594 (1:1000, A-21207 Invitrogen), Goat anti-Rabbit IgG Secondary Antibody, Alexa Fluor™ 488 (1:1000, A-11008 Invitrogen), Goat anti-Mouse IgG Secondary Antibody, Alexa Fluor™ 488 (1:1000, A-11001, Invitrogen), Donkey anti-Mouse Adsorbed Secondary Antibody, Alexa Fluor™ 647 (1:1000, A-31571, Invitrogen), and NeuN Monoclonal Antibody (1:1000, MA5-33103 Invitrogen).

Nissl and FJC staining

Nissl staining assessed neuronal morphology. Slices were stained at 37°C for 30 min, differentiated, and mounted. FJC staining identified degenerating neurons: sections were oxidized with potassium permanganate, stained with FJC solution, dehydrated, and imaged. FJC-positive cells were quantified using ImageJ.

Western blot

Cortical tissues were homogenized in lysis buffer with protease inhibitors. Protein concentration was quantified (BCA assay), separated via SDS-PAGE, transferred to PVDF membranes, blocked in 5% BSA, and incubated with primary and HRP-conjugated secondary antibodies. Bands were visualized using ECL reagents and quantified with ImageJ. Antibodies: anti-RORα (1:1000, ab278108, abcam), anti-GLUT4 (1:1000, ab313775, abcam), anti-Hexokinase 1 (1:800, ab150423, abcam), anti-NDUFB8 (1:1000, ab192878, abcam), anti-SDHB, (1:500, ab175225, abcam), anti-UQCRC2 (1:1000, ab203832, abcam), anti-MTCO1 (1:800, ab14705, abcam), anti-ATP5A (1:1000, ab14748, abcam), and anti-pNRF2 (1:5000, ab76026, abcam).

qRT-PCR

RNA was extracted with TRIzol, and reverse transcription was performed using a SYBR Green PCR kit (TaKaRa). qRT-PCR was conducted using Applied Biosystems equipment. The primer sequences are shown in Table 1.

Table 1.

Quantitative Real Time PCR Primer Sequences

Gene	Forward	Reverse
RORα	5′-CACGAACACAAGGTACTCCGATCTC-3′	5′-CACGAACACAAGGTACTCCGATCTC-3′
IL-1β	5′-AACTGTTCCTGAACTCAACT-3′	5′-ATCTTTTGGGGTCCGTCAACT-3′
TNF-α	5′-CCTCACACTCAGATCATCTTCT-3′	5′-GCTACGACGTGGGCTACAG-3′
IL-6	5′-AGTCCTTCCTACCCCAATTTCC-3′	5′-TTGGTCCTTAGCCACTCCTTC-3′
PCX	5′-ACTTGCCAAACATGCTGCAG-3′	5′-CGTCCCCCGTGTATGAGATG-3′
IDH1	5′-AAGGTTATGGCTCCCTTGGC-3′	5′-TAGTGACGTGTGACAGTGCC-3′
GLUD1	5′-GTGGAAGACCTGAAGACCCG-3′	5′-CGGTAGCCTTCGATGACCTC-3′
ACO1	5′-AACAGAGGCCTAACACCACG-3′	5′-TCCTGAGGGGAGATGGACAG-3′
FH1	5′-AGAGTTTGCGCAGGTCATCA-3′	5′-GCAGCGAGCTCGTAGATTCT-3′
OGDC	5′-CCTGCTGAAAGAGGCACAGA-3′	5′-TGGTGGTACGAAGTCTTGGC-3′
CS	5′-TGGTCCCAGGATACGGTCAT-3′	5′-TTGTACAGCTGAGCCACCAG-3′
DLST	5′-AGCTGCTAAAGGTCTGCGTT-3′	5′-GTCAGCATTGCACACGTGTT-3′
MDH2	5′-GACCTGTTCAACACCAACGC-3′	5′-TAGGGTTGTACACACCGTGC-3′
LDHA	5′-CAAGGAGCAGTGGAAGGAGG-3′	5′-CCAAGTCTGCCACAGAGAGG-3′
PGK1	5′-TGGCCTCTGGTATACCTGCT-3′	5′-GAGTGACTTGGTTCCCCTGG-3′
PKM	5′-GGGAGATGTGGTCATTGTGC-3′	5′-AAGGTACAGGCACTACACGC-3′
PFK1	5′-TGCGTAGAGAGGAGTTTCAGC-3′	5′-CTGTGCTCTGAGTAGTCCTGAC-3′
ENO1	5′-CATGGGGAAGGGTGTCTCAC-3′	5′-GTGCCGTCCATCTCGATCAT-3′
GLUT4	5′-CCAACAGCTCTCAGGCATCA-3′	5′-CACCGAGACCAACGTGAAGA-3′
GLUT3	5′-GTGACTGTGCTGGAGCTCTT-3′	5′-ATCCCAGAGAGCTGCTGAGA-3′
GLUT1	5′-CACTGTGGTGTCGCTGTTTG-3′	5′-AAAGATGGCCACGATGCTCA-3′
HK1	5′-CGGCTCAGAAAAGGGGGATT-3′	5′-TCTTCTCGTGGTTCACCTGC-3′
GPX4	5′-AAAGTCCTAGGAAACGCCCG-3′	5′-CTGTTGCAGTACTGGGGAGG-3′
GPX1	5′-GGACACCAGGAGAATGGCAA-3′	5′-GTAAAGAGCGGGTGAGCCTT-3′
PRPS1	5′-GCAGAGCCAGCTGTCCTAAA-3′	5′-ACCATGCGGTCCACTTCATT-3′
TALDO1	5′-CCACTGCAACATGACACTGC-3′	5′-GTGACACTCTTGACCCCAGG-3′
GLYCTK	5′-ATGCTGCAGAGGACACTGTC-3′	5′-GTAGAGGTTTTCCCGCAGCT-3′
PGD	5′-GGCAAGCCGTGGATGATTTC-3′	5′-CCTTCCTCACCACCACTGAC-3′
G6PD	5′-GTGCTGAGGTTTGCCAACAG-3′	5′-GGTTCCCCACATATTGGCCA-3′
b-actin	5′-CTACCTCATGAAGATCCTGACC-3′	5′-CACAGCTTCTCTTTGATGTCAC-3′

G6PD, glucose-6-phosphate dehydrogenase; GLUT, glucose transporter; HK1, hexokinase 1; IL, interleukin; PCR, polymerase chain reaction; RORα, retinoic acid receptor-related orphan receptor alpha; TNF-α, tumor necrosis factor-alpha.

AAV injection

AAV9 vectors carrying RORα-shRNA under the neuron-specific hSyn promoter (GeneChem) were injected via the tail vein. Protein knockdown was confirmed by Western blot after 2 weeks.

Primary neuron culture and In vitro SAH model

Cortical neurons were harvested from E14.5 embryos, dissociated in papain, and plated at 1 × 10⁶ cells/well in the Neurobasal medium. Glial proliferation was inhibited with cytosine arabinoside. Neurons were transduced with lentiviral vectors expressing RORα or control on DIV2. On DIV10, cells were exposed to 10 µM OxyHb to simulate hemorrhagic insult. For drug studies, 5 µM SR1078 or 0.5% DMSO was added post-OxyHb exposure for 12 h.

Cell viability assay

Neurons (5000/well) were plated in 96-well plates. Viability was assessed with CCK-8 reagent (BestBio) and read at 450 nm (Tecan, Infinite M200). An LDH-Cytotoxicity Colorimetric Assay Kit (Biovision) was used to analyze LDH release in the medium.

ChIP-Seq and RNA-Seq

ChIP was conducted using the Simple ChIP Plus Sonication Chromatin IP Kit Cell Signaling Technology. Libraries were prepared using NEBNext Ultra kits and sequenced (Illumina platform, 2 × 150 bp). RNA-seq was performed after TRIzol extraction, quality-checked (Bioanalyzer, NanoDrop), and libraries were constructed using poly(A) enrichment and size selection. Sequencing was conducted on the Illumina NovaSeq 6000. The dataset generated from this study is available from the National Genomics Data Center (https://www.ngdc.cncb.ac.cn/) with accession numbers PRJCA047313, PRJCA047940.

Bioinformatics analysis

ChIP-seq reads were processed using FASTQC and Bowtie2, with peak calling via MACS2 and annotation using HOMER and ChIPseeker. RNA-seq data were analyzed with fast, HISAT2, StringTie, edgeR, and differential expression defined by FC >2 or <0.5, p < 0.05.

ChIP-qPCR

ChIP was conducted using ChIP Plus Sonication Chromatin IP Kit (Cell Signaling Technology). Cells were cross-linked by 37% methanol and terminated by glycine solution (10×) at room temperature. After washing with PBS twice, cells were collected and were sufficiently lysed with SDS lysis buffer. The ultrasonic (VCX750, Sonics, USA) was performed to cut DNA fragments from 200 to 800 bp. The primary RORα antibody (1 μg; ab278108, abcam; Abcam, USA) was added to immune complex. After a series of centrifugation and washing, the precipitation was collected to perform the q-PCR assay. The primer sequences are shown in Table 2.

Table 2.

Chromatin Immunoprecipitation–Quantitative PCR Primer Sequences

Gene	Forward	Reverse
PGC-1α	5′-CGGAGCTGCTAACTAACAATGG-3′	5′-GATGTGTATCACTGCACCACAC-3′
G6PD	5′-AAACGGTCGTACACTTCG GG-3′	5′-GGTAGTGGTCGTTGCGGTAG-3′

Glucose uptake and metabolic assays

Glucose uptake was measured using a colorimetric assay (BioVision) in 96-well plates. Oxygen consumption rate and ECAR were assessed with Seahorse XF96 analyzers (Agilent) using standard Mito and Glycolysis Stress Test Kits (Li et al., 2018).

D-glucose-¹³C₆ metabolic flux assay and data analysis

The neuron cells (10⁶) were incubated with DMEM containing 2 g/L D-glucose-¹³C₆ for 3 h and then washed with PBS and rapidly quenched using a precooled solution of methanol:acetonitrile:water (2:2:1). The resulting solution was immediately frozen and stored for subsequent metabolomic flux analysis. Metabolomic flux analysis was performed by Shanghai BioTree Biomedical Technology Co., Ltd.

LC-MS method: Water soluble metabolite measurements were obtained by running samples on the Q Exactive PLUS hybrid quadrupole-orbitrap mass spectrometer (Thermo Scientific) coupled with hydrophilic interaction chromatography. An XBridge BEH Amide column (150 × 2.1 mm, 2.5 μM particle size, Waters, Milford, MA) was used. The gradient was solvent A (95%:5% H₂O: acetonitrile with 20 mM ammonium acetate, 20 mM ammonium hydroxide, pH 9.4) and solvent B (100% acetonitrile) 0 min, 90% B; 2 min, 90% B; 3 min, 75%; 7 min, 75% B; 8 min, 70%; 9 min, 70% B; 10 min, 50% B; 12 min, 50% B; 13 min, 25% B; 14 min, 25% B; 16 min, 0% B; 20.5 min, 0% B; 21 min, 90% B; and 25 min, 90% B. The flow rate was 150 μL/min with an injection volume of 5 μL and a column temperature of 25°C. The MS scans were in negative ion mode with a resolution of 140,000 at m/z 200. The automatic gain control target was 1 × 10⁶, and the scan range was m/z 75–1000. All data from isotope labeling experiments were analyzed by El-MAVEN with natural abundance correction. The dataset generated from this study is available from the National Genomics Data Center (https://www.ngdc.cncb.ac.cn/) with accession number PRJCA047941.

ROS and NADPH quantification

ROS levels were measured with a fluorescence-based kit (Abcam, ab186029) and analyzed by flow cytometry (CytoFLEX LX, FlowJo v10.8.1). NADPH was quantified using a commercial kit (Boxbio) at 450 nm.

Mitochondrial membrane potential assay

Mitochondrial membrane potential was assessed using JC-1 dye (Abcam, ab113850), followed by flow cytometry. The emission wavelengths were 529 nm (monomer) and 590 nm (aggregate).

Statistical analyses

Data are expressed as mean ± standard error of mean (SEM). Statistical analyses were conducted using GraphPad Prism 10.1.2 and R Studio. Differences between groups were evaluated using Student’s t-test, one- or two-way ANOVA with LSD, or Dunnett’s T3 post hoc tests. Significance was set at p < 0.05. Sample sizes: in vivo (n ≥ 6), in vitro (≥3 independent replicates). Blinding was applied during data collection and analysis where feasible. Electronic laboratory notebook was not used.

Authors’ Contributions

X.H.Z. and J.L. designed the experiment and wrote the article. J.L. and J.Q.W. performed most of the experiments. S.J.G. generated the SAH model and primary neurons. F.C.Z. and Y.C.J. performed the statistical analysis. All the authors read and approved the final article.

Footnotes

Acknowledgment

The authors thank Professor Jun Pu and professor Yichao Zhao (Renji Hospital, Shanghai Jiao Tong University School of Medicine) for providing the sg/sg mice and LetPub () for its linguistic assistance during the preparation of this article.

Author Disclosure Statement

The authors declare no conflict of interest.

Funding Information

This work was supported by the National Natural Science Foundation of China (82371309 to X.Z. and 82071359 to Y.J.), the Fundamental Research Funds for the Central Universities (No. 24X010202059), and Shanghai Engineering Research Center of Peri-operative Organ Support and Function Preservation (20DZ2254200).

Institutional Review Board Statement

The animal study was approved by the Animal Welfare and Ethics Committee of Shanghai Rat & Mouse Biotech Co., Ltd.

Supplemental Material

Abbreviations

References

Ahn

, Savarraj

, Pervez

, et al. The subarachnoid hemorrhage early brain edema score predicts delayed cerebral ischemia and clinical outcomes. Neurosurgery, 2018; 83(1):137–145; doi: 10.1093/neuros/nyx364

Akula

, McNamee

, Love

, et al. Retinoic acid related orphan receptor α is a genetic modifier that rescues retinal degeneration in a mouse model of Stargardt disease and Dry AMD. Gene Ther, 2024; 31(7–8):413–421; doi: 10.1038/s41434-024-00455-z

Boukhtouche

, Vodjdani

, Jarvis

, et al. Human retinoic acid receptor-related orphan receptor alpha1 overexpression protects Neurones against oxidative stress-induced apoptosis. J Neurochem, 2006; 96(6):1778–1789; doi: 10.1111/j.1471-4159.2006.03708.x

Calabrese

, Cornelius

, Stella

, et al. Cellular stress responses, mitostress and carnitine insufficiencies as critical determinants in aging and neurodegenerative disorders: Role of hormesis and vitagenes. Neurochem Res, 2010; 35(12):1880–1915; doi: 10.1007/s11064-010-0307-z

Chou

, Lan

, Esposito

, et al. Extracellular mitochondria in cerebrospinal fluid and neurological recovery after subarachnoid hemorrhage. Stroke, 2017; 48(8):2231–2237; doi: 10.1161/strokeaha.117.017758

Cueto

, Shen

, Liu

, et al. SAH is a major metabolic sensor mediating worsening metabolic crosstalk in metabolic syndrome. Redox Biol, 2024; 73:103139; doi: 10.1016/j.redox.2024.103139

de Oliveira Manoel

, Goffi

, Marotta

, et al. The critical care management of poor-grade subarachnoid haemorrhage. Crit Care, 2016; 20(1):21; doi: 10.1186/s13054-016-1193-9

Dominy

, Puigserver

. Mitochondrial biogenesis through activation of nuclear signaling proteins. Cold Spring Harb Perspect Biol, 2013; 5(7):a015008; doi: 10.1101/cshperspect.a015008

Gao

, Ding

, Xu

, et al. Neuroprotective effects of Edaravone on early brain injury in rats after subarachnoid hemorrhage. Chin Med J (Engl), 2009; 122(16):1935–1940.

10.

, Zhou

, Zhang

, et al. Global, regional, and national burden of subarachnoid hemorrhage: Trends from 1990 to 2021 and 20-year forecasts. Stroke, 2025; 56(4):887–897; doi: 10.1161/strokeaha.124.048950

11.

Guntermann

, Piaia

, Hamel

, et al. Retinoic-acid-orphan-receptor-C inhibition suppresses Th17 cells and induces Thymic aberrations. JCI Insight, 2017; 2(5):e91127; doi: 10.1172/jci.insight.91127

12.

Guo

, Hu

, Huang

, et al. Role of estrogen-related receptor γ and PGC-1α/SIRT3 pathway in early brain injury after subarachnoid hemorrhage. Neurotherapeutics, 2023; 20(3):822–837; doi: 10.1007/s13311-022-01330-8

13.

Han

, Kim

, Na

, et al. RORα switches transcriptional mode of ERRγ that results in transcriptional repression of CYP2E1 under ethanol-exposure. Nucleic Acids Res, 2016; 44(3):1095–1104; doi: 10.1093/nar/gkv1034

14.

Han

, Kim

, et al. RORα decreases oxidative stress through the induction of SOD2 and GPx1 expression and thereby protects against nonalcoholic steatohepatitis in mice. Antioxid Redox Signal, 2014; 21(15):2083–2094; doi: 10.1089/ars.2013.5655

15.

, Zhao

, Xu

, et al. The nuclear melatonin receptor RORα is a novel endogenous defender against myocardial ischemia/reperfusion injury. J Pineal Res, 2016; 60(3):313–326; doi: 10.1111/jpi.12312

16.

Helleboid

, Haug

, Lamottke

, et al. The identification of naturally occurring neoruscogenin as a bioavailable, potent, and high-affinity agonist of the nuclear receptor RORα (NR1F1). J Biomol Screen, 2014; 19(3):399–406; doi: 10.1177/1087057113497095

17.

Hummasti

, Tontonoz

. Adopting new orphans into the family of metabolic regulators. Mol Endocrinol, 2008; 22(8):1743–1753; doi: 10.1210/me.2007-0566

18.

Kallen

, Schlaeppi

, Bitsch

, et al. Crystal structure of the human RORalpha Ligand binding domain in complex with cholesterol sulfate at 2.2 A. J Biol Chem, 2004; 279(14):14033–14038; doi: 10.1074/jbc.M400302200

19.

Kallen

, Schlaeppi

, Bitsch

, et al. X-ray structure of the hRORalpha LBD at 1.63 A: Structural and functional data that cholesterol or a cholesterol derivative is the natural ligand of RORalpha. Structure, 2002; 10(12):1697–1707; doi: 10.1016/s0969-2126(02)00912-7

20.

Kim

, Mawatari

, Yoo

, et al. The circadian Nobiletin-ROR axis suppresses Adipogenic differentiation and IκBα/NF-κB signaling in adipocytes. Nutrients, 2023; 15(18):3919; doi: 10.3390/nu15183919

21.

Kim

, Han

, Na

, et al. Liver-specific deletion of RORα aggravates diet-induced nonalcoholic steatohepatitis by inducing mitochondrial dysfunction. Sci Rep, 2017; 7(1):16041; doi: 10.1038/s41598-017-16077-y

22.

Kim

, Lee

, Jeong

, et al. RORα–GABP–TFAM axis alleviates myosteatosis with fatty atrophy through reinforcement of mitochondrial capacity. J Cachexia Sarcopenia Muscle, 2024; 15(2):615–630; doi: 10.1002/jcsm.13432

23.

Koepsell

. Glucose transporters in brain in health and disease. Pflugers Arch, 2020; 472(9):1299–1343; doi: 10.1007/s00424-020-02441-x

24.

Kusaka

, Ishikawa

, Nanda

, et al. Signaling pathways for early brain injury after subarachnoid hemorrhage. J Cereb Blood Flow Metab, 2004; 24(8):916–925; doi: 10.1097/01.Wcb.0000125886.48838.7e

25.

, Liu

, et al. KPNA2 promotes metabolic reprogramming in glioblastomas by regulation of c-myc. J Exp Clin Cancer Res, 2018; 37(1):194; doi: 10.1186/s13046-018-0861-9

26.

Munakata

, Ohkuma

, Nakano

, et al. Effect of a free radical scavenger, edaravone, in the treatment of patients with aneurysmal subarachnoid hemorrhage. Neurosurgery, 2009; 64(3):423–428; discussion 428–429; doi: 10.1227/01.Neu.0000338067.83059.Eb

27.

, Shi

, Gilroy

, et al. Regulation of the human hydroxysteroid sulfotransferase (SULT2A1) by RORα and RORγ and its potential relevance to human liver diseases. Mol Endocrinol, 2013; 27(1):106–115; doi: 10.1210/me.2012-1145

28.

Sun

, Lutz

, Tao

. The CRISPR/Cas9 system for gene editing and its potential application in pain research. Transl Perioper Pain Med, 2016; 1(3):22–33.

29.

Tanaka

, Sato

, Akimoto

, et al. Prevention of UVB-induced photoinflammation and photoaging by a Polymethoxy flavonoid, Nobiletin, in human keratinocytes in vivo and in vitro. Biochem Pharmacol, 2004; 68(3):433–439; doi: 10.1016/j.bcp.2004.04.006

30.

Wang

, Zhang

, Wu

, et al. RORα inhibits gastric cancer proliferation through attenuating G6PD and PFKFB3 induced glycolytic activity. Cancer Cell Int, 2024; 24(1):12; doi: 10.1186/s12935-023-03201-4

31.

Wirianto

, Wang

, Kim

, et al. The clock modulator Nobiletin mitigates astrogliosis-associated neuroinflammation and disease hallmarks in an Alzheimer’s disease model. Faseb J, 2022; 36(3):e22186; doi: 10.1096/fj.202101633R

32.

Xiao

, Lv

, Hou

, et al. Sirtuin 5-Mediated lysine Desuccinylation protects mitochondrial metabolism following subarachnoid hemorrhage in mice. Stroke, 2021; 52(12):4043–4053; doi: 10.1161/strokeaha.121.034850

33.

Yue

, Zhao

, Chen

, et al. MicroRNA-7, synergizes with RORα, negatively controls the pathology of brain tissue inflammation. J Neuroinflammation, 2020; 17(1):28; doi: 10.1186/s12974-020-1710-2

34.

Zhao

, Xu

, Ding

, et al. Novel protective role of the circadian nuclear receptor retinoic acid-related orphan receptor-α in diabetic cardiomyopathy. J Pineal Res, 2017; 62(3):e12378; doi: 10.1111/jpi.12378

35.

Zhou

, Yang

, Shen

, et al. Resveratrol improves mitochondrial biogenesis function and activates PGC-1α pathway in a preclinical model of early brain injury following subarachnoid hemorrhage. Front Mol Biosci, 2021; 8:620683; doi: 10.3389/fmolb.2021.620683

Supplementary Material

Please find the following supplemental material available below.

For Open Access articles published under a Creative Commons License, all supplemental material carries the same license as the article it is associated with.

For non-Open Access articles published, all supplemental material carries a non-exclusive license, and permission requests for re-use of supplemental material or any part of supplemental material shall be sent directly to the copyright owner as specified in the copyright notice associated with the article.

0.00 MB

4.31 MB

Retinoic Acid Receptor-related Orphan Receptor α Drives Glucose Reprogramming and Mitochondrial Rescue Mitigate Subarachnoid Hemorrhage-Induced Early Brain Injury

Abstract

Aims:

Methods and Results:

Innovation and Conclusion:

Graphical Abstract

Keywords

Introduction

Innovation

Results

Expression and localization of RORα after SAH

RORα deficiency exacerbated neurological deficits

RORα activation improved neurological outcomes

SR1078 treatment ameliorated neuropathy

RORα overexpression enhanced neuronal transcriptional activity and metabolism

RORα promoted neuronal glucose metabolism

RORα enhanced the neuronal antioxidant capacity

RORα improved mitochondrial bioenergetics

RORα improved the gene expressions of G6PD and PGC1-α through the recruitment of promoters

Discussion

Methods

Animals

Establishment of mouse SAH model

Drug administration

Neurological scoring

Immunofluorescence staining

Nissl and FJC staining

Western blot

qRT-PCR

AAV injection

Primary neuron culture and In vitro SAH model

Cell viability assay

ChIP-Seq and RNA-Seq

Bioinformatics analysis

ChIP-qPCR

Glucose uptake and metabolic assays

D-glucose-13C6 metabolic flux assay and data analysis

ROS and NADPH quantification

Mitochondrial membrane potential assay

Statistical analyses

Authors’ Contributions

Footnotes

Acknowledgment

Author Disclosure Statement

Funding Information

Institutional Review Board Statement

Supplemental Material

Abbreviations

References

Supplementary Material

D-glucose-¹³C₆ metabolic flux assay and data analysis