Background
The retina consists of multiple orders of neurons that collaboratively conduct light-sensing and vision-forming functions. Photoreceptors are the first-order retinal neurons carrying out specialized light-sensing and phototransduction functions, thereby contributing in an indispensable manner to the initiation of the visual processes [
1]. Regardless of the etiologies, genetic or environmental, the progressive loss of photoreceptors by cell death is held responsible for irreversible vision impairment or blindness in patients with photoreceptor degenerative disorders, to name a few, dry age-related macular degeneration, Stargardt disease and retinitis pigmentosa [
2]. Developing mechanisms-based pharmacological agents with photoreceptor protective effects is required given that drug treatments that protect against photoreceptor degeneration are currently available in the clinical settings [
3].
The general consensus of the mechanisms underpinning photoreceptor cell death irrespective of the etiologies centers on the heightened level of oxidative stress [
2]. Oxidative stress, by exerting a broad impact on multiple molecular targets and cellular processes, triggers a cascade of harmful events that invariably lead to photoreceptor cell death. With respect to photoreceptor degeneration, mitochondrial dysfunction plays a protagonist role in oxidative stress as well as oxidative stress-induced cell death, emerging as an important target of photoreceptor protective therapies [
4‐
6]. Therefore, pharmacological agents with antioxidant and mitochondrial protective capacities are under close examination for their therapeutic potentials in attenuating photoreceptor degeneration. Natural products, rich in antioxidants, serve as a value resource for this purpose [
7].
Our previous study has demonstrated that
Cuscuta chinensis Lam., an herbal medicinal that is conventionally used for the treatment of vision impairment in traditional Chinese medicine, is effective at protecting against oxidative stress-mediated photoreceptor degeneration [
8]. Hyperoside, a flavonol glycoside and the signature chemical constituent of
Cuscuta chinensis Lam, is the most abundant flavonoids present in
Cuscuta chinensis Lam [
8]. Studies performed in experimental models of neurogenerative disorders, for instance, Alzheimer’s disease, have demonstrated that hyperoside protects against neuronal loss in part through suppressing oxidative stress and maintaining the functional integrity of mitochondria [
9]. However, whether hyperoside has a direct impact on oxidative stress-induced mitochondrial dysfunction and photoreceptor cell death remains unknown.
Therefore, we set out to test the hypothesis that hyperoside may protect against photoreceptor degeneration by suppressing oxidative stress-induced mitochondrial impairment and photoreceptor cell death. Meanwhile, it is also worth noting that perturbation of the homeostasis in the retinal microenvironment occurs not only as a consequence of photoreceptor degeneration, but also plays an active role in further exacerbating photoreceptor loss and promoting the functional deterioration of the retina. The most typical changes characterizing a pathologically imbalanced retinal microenvironment include inflammatory activation of the retinal resident immune cells, microglia, as well as the gliotic responses of Müller glia, a type of macroglia uniquely present in the retina [
10,
11]. Thus, to better understand the pharmacological implications of hyperoside in protecting against photoreceptor degeneration, we also assessed the impact of hyperoside on photoreceptor degeneration-associated reactive phenotypes in microglia and Müller cells in the retina.
Methods
Reagents
Hyperoside (purity > 98%, Lot. No. P14A11F121347) was ordered from Shanghai Yuanye Biotechnology Co., Ltd (China). Sodium nitroprusside (SNP), hydrocortisone 21-hemisuccinate, progesterone and putrescine were purchased from Sigma-Aldrich (USA). Dulbecco’s modified Eagle’s medium (DMEM) and penicillin/streptomycin were ordered from Thermo Fisher Scientific (USA). Fetal bovine serum (FBS) was purchased from Nobimpex (German).
Cell culture and treatments
661W cone photoreceptor cells, originally obtained from Dr. Muayyad R. AI-Ubaidi [
12], were cultured at 37 °C with 5% CO
2 in DMEM supplemented with 10% FBS, 1% penicillin/streptomycin, 40 ng/mL hydrocortisone 21-hemisuccinate, 40 ng/mL progesterone and 100 μmol/L putrescine. For the experiments involving SNP stimulation, 661W cells were treated with hyperoside at the indicated concentrations or vehicle for 1 h, followed by SNP incubation at 300 μM for 3 or 4 h. Preliminary studies were carried out to help determine the doses and the time points of the indicated treatments.
Real-time assessment of cell death
To assess cell death, the cell-impermeant dsDNA-binding YOYO-1 iodide dye (Thermo Fisher Scientific, USA) was applied, followed by real-time quantification of YOYO-1-stained cells by Incucyte live cell analysis platform. Briefly, 4 h after the indicated treatments, 661W cells were incubated in 0.1 μM YOYO-1 iodide staining solution at 37 °C for 10 h. Four phase-contrast and fluorescent images per well were automatically captured at 1-h intervals. The integrated object counting algorithm was used to isolate the fluorescent nuclear signal from background. Specifically, images were segmented in order to identify individual objects, counted, and reported on a per-area basis for each time point. The confluence of green fluorescent signals was then measured by the Incucyte Live-Cell Analysis System (Sartorius, German).
Assessment of mitochondrial membrane potential (MMP)
MMP was detected using the Mitochondrial Membrane Potential Assay kit (Beyotime, China). Briefly, 3 h after the indicated treatments, 661W cells were incubated in 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) detection solution at 37 °C for 20 min. The mitochondrial probe JC-1 is a lipophilic cationic dye that exhibits green fluorescence in the monomer forms. JC-1 dye can accumulate in the intact mitochondria and form the red fluorescent complex called J-aggregates that serve as a monitor of MMP [
13]. The red fluorescence of J-aggregates and the green fluorescence of JC-1 monomers were acquired using a fluorescence microscope (DMI6000, Leica, Germany). Quantification of immunofluorescence was performed by ImageJ.
Measurement of mitochondrial permeability transition pore (MPTP) opening
The opening of mPTP was detected using the Mitochondrial Permeability Transition Pore Assay kit (Beyotime, China). In brief, 3 h after the indicated treatments, cell-permeable calcein acetoxymethyl ester and CoCl2, the quencher of calcein fluorescence, were applied at 37 °C for 30 min to selectively label mitochondria. The green fluorescent signal of calcein was then visualized using a fluorescence microscope (DMI6000, Leica, Germany) and quantified by ImageJ.
Measurement of mitochondrial superoxide production
The mitochondrial superoxide production was detected using the MitoSOX Red mitochondrial superoxide indicator (Thermo Fisher Scientific, USA). In brief, 3 h after the indicated treatments, 661W cells were incubated in the solution containing MitoSOX Red mitochondrial superoxide indicator at 37 °C for 30 min. The red fluorescent signal indicative of mitochondrial superoxide was acquired using a fluorescence microscope (DMI6000, Leica, Germany) and quantification was performed by ImageJ.
Animals and treatments
Six-week-old female Balb/c mice were obtained from Shanghai Laboratory Animal Research Center and maintained in a laboratory with a 12/12 h light–dark cycle and a controlled temperature set at 20 ± 2 °C. The mice were dark-adapted for 24 h before the experimental light exposure (Compact Fluorescence Lamp, 45 W, Chaoya Lighting, Shanghai, China) delivered at 15,000 lx for 30 min. Hyperoside was dissolved in 0.5% sodium carboxymethyl cellulose solution and administered intraperitoneally 30 min before the light exposure at the indicated dose(s). Balb/c mice unexposed to the experimental light (normal controls) and the light-exposed mice without hyperoside treatment were treated with the vehicle in the same fashion. The volume of the intraperitoneal injection was controlled at 100 μL per mouse. Twenty-four mice from each experimental group were analyzed for the indicated imaging, electrophysiological, histopathological and molecular biological assessments at the specified time point(s). In total, 112 mice were utilized in the current study. The laboratory animal handling protocol was reviewed and approved by the Institutional Animal Care and Use Committee of Yueyang Hospital of Integrated Traditional Chinese and Western Medicine, Shanghai University of Traditional Chinese Medicine (YYLAC-2020-079-2 and YYLAC-2023-199-1) and carried out in accordance with the recommendations of the NIH Guide for the Care and Use of Laboratory Animals and the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research.
Optical coherence tomography (OCT)
Image-guided OCT (OCT 2 with Micron IV, Phoenix Research labs, USA) was adopted to image the retina 7 d after the experimental light exposure. In brief, anesthesia was induced by intraperitoneal injection of ketamine hydrochloride (82.5 mg/kg bw) and xylazine (8.25 mg/kg bw), followed by dilation of pupils using 1% tropicamide (Santen Pharmaceutical, Japan) prior to OCT imaging. Five full-retinal scans were acquired and automatically averaged using Phoenix Reveal OCT software (Phoenix Research labs, USA). The averaged scans were presented and subject to morphological evaluation of the retina. The thickness of the outer nuclear layer (ONL) was measured with Insight Image Segmentation Software for the Phoenix OCT and Retinal Imaging System (Version 2.0.5490, Voxeleron LLC, USA).
Electroretinography (ERG)
Seven days after the experimental light exposure, the mice were dark-adapted for 24 h and subjected to ERG analysis under the safe light (5 lx) as previously described [
14]. Briefly, prior to ERG procedures, intraperitoneal injection of ketamine hydrochloride (82.5 mg/kg bw) and xylazine (8.25 mg/kg bw) was performed to induce anesthesia. Pupils were then dilated using 1% tropicamide and the eyes were kept moisturized using 0.5% hypromellose solution. Once the mice were sedated, the reference and ground electrodes were inserted subcutaneously in the head in the midline between the ears and in the tail toward the base of the tail, respectively. The ERG responses were then recorded and analyzed by LabScribe software using Ganzfeld (ERG 2, Phoenix Research Labs, USA). Flashes of green light (504 nm) were delivered at the intensity of -2 (0.5 ms duration and 5 s inter-stimulus-interval), − 0.8 (1 ms duration and 5 s inter-stimulus-interval), 0.4 (1 ms duration and 10 s inter-stimulus-interval), 1.6 (1 ms duration and 20 s inter-stimulus-interval) and 3.1 (1 ms duration and 60 s inter-stimulus-interval) log cd·s·m
−2.
Histological examination and immunohistochemistry (IHC)
The enucleated eyes were fixed in 4% paraformaldehyde for 24 h before further processing and paraffin sectioning. Paraffin sections (4 μm in thickness) were stained with hematoxylin and eosin (HE) or subjected to IHC examination using primary antibodies including mouse anti-rhodopsin (1:1000) (Novus, USA), rabbit anti-opsin (1:100) (Red/Green, M-opsin) (Millipore, USA), rabbit anti-opsin (1:100) (Blue, S-opsin) (Millipore, USA) and rabbit anti-glial fibrillary acidic protein (GFAP) (1:500) (DAKO, USA) as well as Cy3-conjugated sheep anti-rabbit (1:1000) or sheep anti-mouse secondary antibodies (1:1000) (Sigma-Aldrich, USA). In addition, eye cups free of the cornea and lens were made and fixed in 4% paraformaldehyde for 2 h at room temperature and processed for cryosectioning. Cryosections (12 µm in thickness) were subjected to IHC examination using primary antibodies including rabbit anti-ionized calcium binding adaptor molecule 1 (Iba-1) (1:500) (Wako Chemicals, USA) and the secondary antibody Cy3-conjugated sheep anti-rabbit (1:1000). Counterstaining of 4-6-diamidino-2-phenylindole (DAPI) (Sigma Aldrich, USA) was performed for nuclei visualization. Microscopic imaging was performed by a light microscope (DM2000, Leica, Germany) or a fluorescent microscope (DM6000B, Leica, Germany). The value of gain and exposure time were maintained the same for the microscopic imaging of the fluorescent signals.
Transmission electron microscope (TEM)
The superior part of the eye cup was dissected and fixed in 2.5% glutaraldehyde at 4 °C overnight. After washing off glutaraldehyde, the specimens were post-fixed in 1% osmic acid, dehydrated in ascending concentrations of ethanol, stained with 3% uranyl acetate and embedded in Epon 812 embedding fluid. Ultra-thin sections (70 nm in thickness) were then made and stained with lead citrate. Images were recorded digitally by a H-7650 transmission electron microscope (HITACHI, Japan).
Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL)
Enucleated eyes were fixed in 4% paraformaldehyde and processed for paraffin sectioning. Paraffin sections (4 μm in thickness) were then subjected to TUNEL assay (DeadEnd™ Fluorometric TUNEL System, Promega) following the manufacturer’s protocols. TUNEL positivity was observed and recorded using a fluorescent microscope (DM6000B, Leica, Germany). The value of gain and exposure time were maintained the same during the microscopic imaging process. ImageJ was used to quantify the TUNEL positivity.
RNA sequencing
Total RNA was extracted from dissected retinas using mirVana miRNA Isolation Kit (Thermo Fisher Scientific, USA). RNA purity and quantification were evaluated by a NanoDrop 2000 spectrophotometer (Thermo Scientific, USA). Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA) was used to assess RNA integrity. TruSeq Stranded mRNA LT Sample Prep Kit (Illumina, San Diego, CA, USA) was adopted to construct the cDNA libraries. The libraries were sequenced on an Illumina HiSeq X Ten platform and 150 bp paired-end reads were generated. Raw data (raw reads) of fastq format were processed using Trimmomatic [
15] and the reads of low quality were removed to obtain the clean reads. Fragments per kilobase of transcript per million mapped reads (FPKM) of each gene was calculated using Cufflinks [
16]. The read counts of each gene were obtained by HTSeq count [
17]. Principal component analysis (PCA) was performed to evaluate the distribution and variation of the samples. The correlation heatmap was produced in R using the stats package. Differential expression analysis was performed using the DESeq (2012) R package. P value < 0.05 and fold change > 2 or fold change < 0.5 were set as the threshold for significantly differential expression. Hierarchical cluster analysis of differentially expressed genes (DEGs) was performed to demonstrate the expression pattern of genes in the indicated experimental groups and individual samples. Gene set enrichment analysis (GSEA) was performed for functional enrichment of the genes using Kyoto Encyclopedia of Genes and Genomes (KEGG) and Gene Ontology (GO) gene sets with ClusterProfiler R package and org.Mm.eg.db annotation package. The gene sets with significant enrichment were defined with absolute value of normalized enrichment score (NES) > 1 and false discovery rate (FDR) q-value < 0.05. Benjamini–Hochberg procedure was used for the correction of the related FDR q-value. The bubble plot of GSEA core-enriched signaling pathways was generated with ggplot2 in R package.
Real-time qPCR analysis
TRIzol reagent (Invitrogen, USA) was used for isolating RNA from the mouse retinas. PrimeScript RT Master Mix (TaKaRa, Japan) was then used for reverse transcription. The expression of
Abca4,
Aim2,
Axl,
Bbs9,
Casp4,
Casp8,
Ccl2,
Cd68,
Clec7a,
Cnga1,
Crx,
Gfap,
Glul,
Gnat1,
Guca1a,
Il1b,
Mlkl,
N2re3,
Naip2,
Nlrp3,
Nrl,
Nxnl1,
Opn1mw,
Opn1sw,
P2ry12,
Pde6b,
Prph2,
Rdh12,
Reep6,
Rho,
Ripk1,
Ripk3,
Rom1,
Rp1l1,
Rpgrip1,
Slc24a1,
Tmem119,
Tnf,
Tspo and
Ush2a was examined using SYBR Green I Master (Roche, USA) on a Roche Light Cycler 480 II (Roche, USA). The primer sequences were included in Table
1. The expression of the indicated genes was normalized to
18S rRNA. The fold change of gene expression was calculated based on 2
−[Ct (candidate gene)−Ct (18s rRNA)].
Table 1
Primer sequences for real-time qPCR analysis
Abca4 | CAGAAGATTCGCTTTGTAGTGGA | CCTTGTTGGGAAAATGGCATTC |
Aim2 | GTCACCAGTTCCTCAGTTGTG | CACCTCCATTGTCCCTGTTTTAT |
Axl | ATGGCCGACATTGCCAGTG | CGGTAGTAATCCCCGTTGTAGA |
Bbs9 | AGCCACCAATGTGGAACCTGGA | CTGTAGTGGAGGCTGCACATAG |
Casp4 | GTGGTGAAAGAGGAGCTTACAGC | GCACCAGGAATGTGCTGTCTGA |
Casp8 | CGGTGAAGAACTGCGTTTCC | ACGCCAGTCAGGATGCTAAG |
Ccl2 | AGCTGTAGTTTTTGTCACCAAGC | GTGCTGAAGACCTTAGGGCA |
Cd68 | GGCGGTGGAATACAATGTGTCC | AGCAGGTCAAGGTGAACAGCTG |
Clec7a | GACTTCAGCACTCAAGACATCC | TTGTGTCGCCAAAATGCTAGG |
Cnga1 | CGAGCCATTTGTGCTGCTTA | TCATGGTTAGTTTAATATCTGCGCT |
Crx | CCAATGTGGACCTGATGCACCA | GTACTGGGTCTTGGCAAACAGG |
Gfap | CCGAGTACTGAAGCCAAGGG | GCAGTTTGTAACCCCTCCCA |
Glul | GAGGAGAATGGTCTGAAGTGC | ACCGGCAGAAAAGTCGTTGA |
Gnat1 | CCCGACTACGATGGACCTAAC | TTGACGTTCTGTGTGTCGGT |
Guca1b | CTGGACATTGTGGAGGCGAT | GACAGCTGGCCGTCTCCATT |
Il1b | TGCCACCTTTTGACAGTGATG | AAGGTCCACGGGAAAGACAC |
Mlkl | CTGAGGGAACTGCTGGATAGAG | CGAGGAAACTGGAGCTGCTGAT |
N2re3 | GCCTTATCACCGCCGAAACTTG | CATGGATGCCATCCAGACTGCA |
Naip2 | AGCTTGGTGTCTGTTCTCTGT | GCGGAAAGTAGCTTTGGTGTAG |
Nlrp3 | ATTACCCGCCCGAGAAAGG | TCGCAGCAAAGATCCACACAG |
Nrl | CTCTTGGCTACTATTCAGGGAGC | GGTTCAACTCGCGCACAGACAT |
Nxnl1 | GGAACAACAGCGACCAGGAT | GTGAGCCGCACGAAGAAGT |
Opn1mw | AGCCCTTTGGCAATGTGAGA | AAGGCCAGTACCTGCTCCAA |
Opn1sw | TCATCTTCTGTTTCATCATTCCTCT | CTTTTGTGTCGTAGCAGACTCTT |
P2ry12 | ATGGATATGCCTGGTGTCAACA | AGCAATGGGAAGAGAACCTGG |
Pde6b | TGGAGAACCGTAAGGACATCGC | TCCTCACAGTCAGCAGGCTCTT |
Prph2 | GCAATCGCTACCTGGACTTCTC | GTGAGCTGGTACTGGATACAGG |
Rdh12 | ATCTTGGTACTGCTTACGTCCT | CACCAGCAAAGAACTTCCTGA |
Reep6 | CGGTTACGGGGCCTCTCTA | CCAGTAGGTTAGCCACACAGT |
Rho | CCTTTGTCATCTACATGTTCGTGGT | CTTCCTTCTCTGCCTTCTGAGTGGT |
Ripk1 | GACTGTGTACCCTTACCTCCGA | CACTGCGATCATTCTCGTCCTG |
Ripk3 | CCACACCAGCAAGGACATCT | GCCGAACTTGAGGCAGTAGT |
Rom1 | TGGGTCAGCAACCGTTACTTGG | GAGAGTTGGCTTTGCAGGCAAG |
Rp1l1 | TGTGACTGCGAGGAGTGAACGT | TCAAGGAAACCTGCCGCAGCTT |
Rpgrip1 | GACCACGAAGAGAAACTGGAGC | CAGAGTGCCATACGCGACATCT |
Slc24a1 | GTCAAGGTCTGAAGGTTTGGG | TCTTTGGTCGGAGTAACCGC |
Tmem119 | CCTACTCTGTGTCACTCCCG | CACGTACTGCCGGAAGAAATC |
Tnf | ACGTCGTAGCAAACCACCAA | GCAGCCTTGTCCCTTGAAGA |
Tspo | GAGCCTACTTTGTACGTGGCGA | GCTCTTTCCAGACTATGTAGGAG |
Ush2a | TGCTCAGTGACCCTGTTTCCGT | TTGTTGCGAGCTGGTGTAGACC |
18S rRNA | GAGGTTCGAAGACGATCAGA | TCGCTCCACCAACTAAGAAC |
Statistical analysis
Data were expressed as mean ± standard error of mean (SEM) or mean ± standard deviation (SD). Statistical analyses were performed by one-way or two-way ANOVA with the Tukey’s multiple comparisons test (GraphPad Prism 9, USA). Statistically significance was defined if P < 0.05.
Discussion
In the present study, we demonstrate that hyperoside is pharmacologically active at protecting against oxidative stress-induced mitochondrial impairment and photoreceptor cell death in vitro. Most importantly, these cell-intrinsic photoreceptor protective activities of hyperoside are translatable in vivo as hyperoside attenuates photooxidative stress-induced photoreceptor degeneration on molecular, cellular, structural and functional levels. Moreover, hyperoside treatment counteracts neuroinflammatory responses and reactive alterations in microglia and Müller cells in the retina, providing additional evidence supporting the pharmacological significance of hyperoside in maintaining retina homeostasis under photooxidative stress conditions.
Our major findings here relate to the pharmacological implications of hyperoside in suppressing oxidative stress-mediated photoreceptor degeneration. Oxidative stress plays a pivotal role in triggering cell death, which is especially relevant in the case of photoreceptor cell death, the key cellular mechanism underlying photoreceptor degeneration [
20]. This is due to highly specialized function of photoreceptors for light sensing, a process naturally generating ROS; high density of mitochondria, the primary source of ROS, in the photoreceptor IS; enrichment of the polyunsaturated fatty acids in the photoreceptor OS, rendering photoreceptors enhanced susceptibility to oxidation damage; anatomical adjacency of photoreceptors to the choroidal vasculature, an environment with high oxygen flow. Mitochondrial impairment is at the center of oxidative stress in that mitochondria are not only the major source of ROS, they are also the victims of the deleterious impact of ROS. Mitochondrial impairment and oxidative stress are early changes associated with photoreceptor degeneration [
5]. Dysfunctional mitochondria further induce and aggravate oxidative stress [
21,
22]. This mitochondria-based vicious cycle of overproduction of ROS is especially relevant for the pathophysiology of photoreceptor degeneration given that as high energy-consuming cells, photoreceptors contain approximately 90% of retina’s mitochondria and rely heavily on mitochondria for their survival and functionality. The critical role of intact mitochondria for photoreceptor health is also supported by the findings that photoreceptor degeneration is one of the major phenotypes of mitochondrial disorders [
23,
24]. Therefore, protecting against mitochondrial impairment and oxidative stress serves as a viable route for the control of photoreceptor degeneration. Here we demonstrate that hyperoside suppresses oxidative stress-triggered mitochondrial impairment and photoreceptor cell death in vitro and in vivo. Hyperoside has been shown to directly counteract the deleterious impact of oxidative stress on the survival of PC12 neuronal cells [
25]. Meanwhile, hyperoside mitigates Aβ-induced mitochondrial dysfunction in HT22 neuronal cells. The work here further demonstrates that hyperoside attenuates oxidative stress-induced mitochondrial dysfunction and cell death in 661W photoreceptor cells. Most importantly, our in vivo findings provide direct support to the significant effects of hyperoside on preserving the integrity of mitochondria under photooxidative stress conditions. Furthermore, hyperoside protects against photooxidative stress-induced loss of photoreceptor integrity on molecular, morphological, structural and functional levels. Putting these findings together, it is possible that hyperoside may exert photoreceptor protection in part by alleviating oxidative stress-induced mitochondrial impairment in photoreceptors. However, the molecular targets mediating the suppressive effects of hyperoside on oxidative stress-induced photoreceptor mitochondrial impairment and cell death remain to be identified in the future studies.
Aside from photoreceptors, it is worth noting that hyperoside treatment attenuates photoreceptor degeneration-associated neuroinflammatory responses and microglial activation in the retina. Retina is a complex neuronal tissue consisting of various types of neurons to execute the essential function of vision formation as well as non-neuronal cells that provide a supportive microenvironment underpinning the normal structural and functional homeostasis of the retina. Among the non-neuronal cellular constituents of the retina, glial cells, namely microglia, astrocytes and Müller cells, are equipped with important functions for maintaining a homeostatic microenvironment in the retina [
26]. The pathophysiological implications of microglia in photoreceptor degeneration have been increasingly acknowledged. Under normal conditions, the resident immune cells of the retina, microglia, play an important role in maintaining retinal homeostasis via immune surveillance. Disturbance in the retinal microenvironment leads to rapid activation of microglia. More than merely an immediate response to the insults to the retina, aberrantly activated microglia are key players in mediating neuroinflammatory responses that are toxic to the retinal neurons, thereby exacerbating photoreceptor degeneration [
10]. The results from our whole-genome gene expression profiling analyses demonstrate that LE retinas are characterized by significant upregulation of multiple molecular pathways associated with neuroinflammation and microglial activation. Most importantly, hyperoside treatment results in remarkable downregulation of the pathways implicated in neuroinflammation and microglial activation. In addition, microglia are normally located in the inner plexiform layer and outer plexiform layer in the retina. During photoreceptor degeneration, ectopic microglia are found in the outer nuclear layer and the subretinal space. Our results shown that hyperoside treatment also results in fewer microglia in the outer retina. Although it is likely that dampened activation of microglia is due to attenuated photoreceptor degeneration resulting from hyperoside treatment, a direct impact of hyperoside on microglial inflammatory activation is still possible. This possibility is supported by the findings that hyperoside suppresses lipopolysaccharide-induced inflammatory responses in microglia and attenuates the neurotoxic effects of activated microglia [
27,
28]. Therefore, future studies are necessary to elucidate the potential implications of hyperoside in suppressing photoreceptor degeneration-associated microglia activation.
In addition to the impact on microglia, the suppressive effects of hyperoside on the reactive gliotic pathologies of Müller cells are also worth noting. Müller cells are not only the predominant glia of the retina, they are also the only glia specifically found in the retina. Müller cells closely interact with the retinal neurons including photoreceptors to maintain a homeostatic environment by regulating the metabolism as well as the extracellular milieu essential for the survival and normal function of the retinal neurons. Nearly all known retinal disorders are associated with a reactive gliotic changes in Müller cells, which is characterized by aberrantly upregulated expression of Gfap [
11]. Although reactive gliosis of Müller cells is initiated with the intention to restore retinal homeostasis, it may accelerate the progression of retinal degeneration when Müller cells acquire malfunctional phenotypes defined in part by downregulation of the key enzyme glutamine synthetase encoded by
Glul. Glutamine synthetase is primarily responsible for catalyzing the conversion of glutamate, a neurotransmitter that mediates normal excitatory synaptic transmission, to glutamine, the precursor of glutamate, thereby playing an essential role in regulating the glutamate-glutamine cycle, the shuttle of glutamate and glutamine between neurons and Müller cells. Inhibition of the glutamine synthetase leads to disturbed glutamate–glutamine cycle, causing marked reduction in the function of the retinal neurons including photoreceptors [
29]. Injured photoreceptors release massive amounts of glutamate and excessive glutamate is neurotoxic. Downregulation of
Glul expression can therefore directly cause insufficient metabolism of glutamate released by damaged photoreceptors, further exacerbating photoreceptor cell death. On the other hand, increasing the expression of glutamine synthetase confers protection against retinal degeneration [
30]. Thus, gliotic Müller cells may fail to carry out their neuron-supportive functions and contribute to neuronal dysfunction and cell death [
31]. Our findings here demonstrate that LE retinas are characterized by upregulated expression of
Gfap and simultaneous downregulation of
Glul. Meanwhile, aberrant expression pattern of Gfap is notable throughout the course of photoreceptor degeneration. These observations indicate that reactive gliosis of Müller cells is not only a phenotype of disturbed retinal homeostasis, it is also actively involved in the progression of photoreceptor degeneration under photooxidative stress conditions. Hyperoside treatment results in decreased expression of
Gfap and increased expression of
Glul in the light-exposed retinas. Meanwhile, the expression pattern of Gfap in the HYP retinas remains comparable to the NLE retinas. These results further highlight the beneficial impact of hyperoside on the retinal homeostasis under photooxidative stress conditions. However, whether hyperoside exerts direct impact on the pathophysiological changes in Müller cells remains to be investigated.
Publisher's Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.