NMO-IgG causes primary retinal damage without optic nerve injury

doi:10.21203/rs.3.rs-5013083/v1

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Research Article

NMO-IgG causes primary retinal damage without optic nerve injury

https://doi.org/10.21203/rs.3.rs-5013083/v1

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Background

Neuromyelitis Optica (NMO) is a neuroimmune disorder primarily driven by autoantibodies against aquaporin 4 (AQP4), known as NMO-IgG. Although the mechanisms underlying NMO-IgG-induced retinopathy are not fully understood, the high expression of AQP4 in retinal Müller cells suggests a direct interaction that may trigger inflammatory processes in the retina. Previous studies indicate that microglia play a critical role in mediating immune responses, leading to neuronal dysfunction.

Methods

NMO-IgG obtained from clinical patients was administered via intravitreal injection to female C57BL/6 mice. Techniques such as optical coherence tomography (OCT), electroretinography (ERG), real-time fluorescence quantitative PCR (RT-qPCR), and immunofluorescence analyses were used to assess retinal changes. The potential for reversing retinopathy was explored by depleting microglial cells using the CSF1 receptor inhibitor PLX3397. Additionally, a Transwell co-culture system of MIO-M1 (Müller cells) and BV2 (microglia) cells was established to study their interactions.

Results

Intravitreal injection of purified NMO-IgG in mouse models led to its deposition in the retina and downregulation of AQP4 in Müller cells. Vascular leakage was observed, alongside retinal dysfunction characterized by thinning of the retinal nerve fiber layer (RNFL) and loss of retinal ganglion cells (RGCs), consistent with ERG findings. By day 7, C3 expression was upregulated in Müller cells, followed by microglial activation. Significant morphological changes in microglia were noted, with increased expression of iNOS and C1q, indicating substantial activation. Ablating microglia significantly mitigated NMO-IgG-induced injury to RGCs. In vitro, NMO-IgG-treated MIO-M1 cells secreted higher levels of C3, enhancing the activation and migration of BV2 cells compared to controls.

Conclusions

The retinal dysfunction observed in NMO may primarily be linked to the activation of Müller cells by NMO-IgG, leading to increased C3 secretion, which in turn activates microglia. Therapeutic strategies targeting Müller cell-microglia interactions in NMO-IgG-induced retinopathy could be promising in addressing the underlying retinal pathology in this condition.

NMO

Retinopathy

Müller cells

Microglia

C1q

Neuromyelitis Optica Spectrum Disorder (NMOSD) is an autoimmune inflammatory demyelinating condition that predominantly affects the optic nerve and spinal cord within the central nervous system (CNS)[1–3]. Patients with NMOSD-associated optic neuritis (ON) frequently exhibit thinning of the ganglion cell and inner plexiform layer (GCIPL) and the retinal nerve fiber layer (RNFL), indicating neuronal and axonal degeneration[4]. Interestingly, research suggests that subclinical visual function and structural abnormalities may be present even in the unaffected eyes of NMOSD patients, which indicates that primary retinal injury might precede the onset of optic neuritis in NMO[5]. This primary retinal pathology could potentially lead to significant vision and visual field impairments independent of optic nerve damage.

More than 70% of NMO patients are seropositive for an autoantibody (NMO-IgG or AQP4-IgG) that targets the water channel protein aquaporin-4 (AQP4), a key pathogenic factor in NMOSD[6]. The binding of this autoantibody to AQP4 can trigger the internalization of AQP4, leading to astrocyte dysfunction and subsequent secondary demyelination and axonal loss[7]. Early animal models of NMO suggested that NMO-IgG mediates complement-dependent cytotoxicity (CDC) against astrocytes, which plays a critical role in the disease's pathogenesis[8]. However, in vitro studies have shown that NMO-IgG induces rapid AQP4 internalization, which reduces complement-dependent cytotoxicity[9]. Additionally, recent translational research revealed that in over 50% of NMO patients, NMO-IgG primarily induces AQP4 internalization rather than CDC[10].

Clinical neuropathological observations have highlighted prominent microglial activation in the early stages of NMO, underscoring the significance of microglia in NMO pathogenesis[11]. A novel animal model has uncovered an unexpected astrocyte-microglia interaction that drives the progression of NMO[12, 13]. However, whether microglial activation contributes to NMO-associated retinopathy remains an open question.

In this study, we developed a new mouse model of NMO-IgG-associated retinopathy. Our results demonstrate that NMO-IgG induces retinal ganglion cell (RGC) injury, leading to abnormalities in electroretinography (ERG) and optical coherence tomography (OCT). Microglial activation is an early hallmark in this model, and depletion of microglia significantly rescues RGC damage induced by NMO-IgG. Furthermore, in vitro studies revealed that NMO-IgG stimulates Müller cells to produce complement C3, which attracts microglia to release complement C1q, thereby inducing neuronal dysfunction. Our findings suggest that microglia may serve as a promising therapeutic target for primary retinopathy in NMOSD.

Patient Samples and Human IgG Purification

Blood samples were collected from four healthy controls (HCs) and four patients diagnosed with Neuromyelitis Optica Spectrum Disorder (NMOSD) at the Ophthalmology Department of the Chinese People’s Liberation Army (PLA) General Hospital. The selected patients were seropositive for AQP4-IgG, as confirmed by a cell-based indirect immunofluorescence assay, and were diagnosed according to the 2015 International Consensus Diagnostic Criteria. Exclusion criteria included the presence of other immune disorders, infections, or cancer. Patient demographics, including age, sex, and disease duration, are summarized in Table 1. None of the healthy controls had a history of disease or infection, and none had received any treatment in the preceding two months. Informed consent was obtained from all participants.

Table 1

Demographic and clinical characteristics of NMOSD patients and healthy controls.
	Age (years)	Sex (M/F)	AQP4-IgG titer		Disease duration (Months)
	Age (years)	Sex (M/F)	CSF	Plasma	Disease duration (Months)
Patient 1	52	F	1:3.2	1:1000	11
Patient 2	31	F	1:1	1:1000	20
Patient 3	50	F	-	1:1000	3
Patient 4	45	F	1:3.2	1:1000	9
Control 1	42	M	-	-	-
Control 2	42	F	-	-	-
Control 3	31	F	-	-	-
Control 4	40	F	-	-	-

Serum samples were heat-inactivated at 56°C for 30 minutes to deactivate complement proteins. Total IgG was purified from 4 mL of pooled serum from NMOSD patients and healthy controls using a protein-A column (GE Healthcare Bio-sciences, USA) and concentrated with Amicon Ultra-4 centrifugal filters (Merck Millipore, Germany). The purified IgG was diluted to 4 mL with phosphate-buffered saline (PBS) and sterilized using a 0.2 µm filter.

Animals and Intravitreal Injection

Female C57BL/6 mice aged 6–8 weeks and weighing 20–25 g were obtained from SPF Biotechnology (Beijing, China) and housed under controlled conditions with a 12-hour dark/light cycle, with free access to food and water. All experimental protocols adhered to the Statement for the Use of Animals in Ophthalmic and Vision Research and were approved by the Institutional Animal Care and Research Advisory Committee of the PLA General Hospital (Approval number: 2021-X17-08).

Mice were anesthetized via intraperitoneal injection of sodium pentobarbital. Intravitreal injections of 2 µL purified NMO-IgG were administered to the right eyes of the mice, while the left eyes served as controls and received equal volumes of purified HC-IgG. To prevent leakage, the needle was kept in place for an additional minute post-injection. Retinal circulation was confirmed by assessing the color of the retinal vessels, and antibiotic ointment was applied to prevent infection.

Pharmacological Ablation of Microglia

To deplete retinal microglia, mice were fed a diet containing the colony-stimulating factor 1 receptor (CSF1R) inhibitor PLX3397 (600 mg/kg, Moldiets, China) starting 7 days before the intravitreal injection and continuing until sacrifice. Control groups were fed an AIN76A chow diet, which had the same composition as the experimental diet but lacked PLX3397. The effectiveness of microglial ablation was evaluated by immunostaining retinal tissue sections.

Evans Blue Assay

A 2% (w/v) solution of Evans Blue dye (Sigma-Aldrich, St. Louis, Missouri, USA) was injected via the tail vein, followed by a 2-hour circulation period. After circulation, eyeballs were collected and immersed in 4% paraformaldehyde (PFA) for 2 hours. The retinas were carefully dissected and shaped into a four-leaf clover configuration. Blood-retinal barrier (BRB) integrity was examined using a microscope (Olympus, Japan) with an excitation wavelength of 594 nm.

Immunofluorescence Staining

After anesthesia, mice were perfused with saline and 4% PFA. Eyeballs were then fixed with 4% PFA for 1 hour at room temperature (RT). For flat mounts, retinas were dissected and rinsed with PBS containing 0.1% Tween (PBST). For frozen sections, eyeballs were dehydrated in sucrose at increasing concentrations (10%, 20%, and 30%) after fixation. Following the removal of corneas and lenses, the eyeball cups were embedded in O.C.T. and frozen at − 80°C. Cross-sections (12 µm) were obtained using a Thermo cryostat (CRYOSTAR NX50, Thermo, USA) and mounted onto microscope slides.

Tissues were blocked with PBS containing 5% bovine serum albumin and 0.1% Triton X-100 (blocking buffer) and incubated with primary antibodies overnight at 4°C. After washing with PBST, tissues were incubated with secondary antibodies for 1 hour at RT, followed by counterstaining with DAPI. Images were captured using the Operetta CLS High-Content Analysis System (PerkinElmer, Waltham, MA, USA) or a confocal imaging system (Olympus FV100, Olympus, Japan).

The antibodies used included mouse anti-AQP4 (ab9512, Abcam), rabbit anti-AQP4 (A5971, Sigma), mouse anti-GFAP (MAB360, Sigma), rabbit anti-IBA-1 (ab178846, Abcam), rat anti-IBA-1 (ab283346, Abcam), rabbit anti-GS (PA5-28940, Thermo Scientific), rat anti-C3 (ab11862, Abcam), rabbit anti-RBPMS (PA5-31231, Invitrogen), mouse anti-C1q (ab182452, Abcam), rat anti-C1q (ab11861, Abcam), mouse anti-iNOS (sc-7271, Santa Cruz), rat anti-CD31 (ab56299, Abcam), rabbit anti-MBP (ab40390, Abcam), and goat anti-human IgG Alexa Fluor 488 (A11013, Invitrogen).

Images were analyzed using Fiji software (National Institutes of Health, Bethesda, MD, USA). The average retinal ganglion cell (RGC) count was calculated using two fields from each retinal quadrant in flat mounts. AQP4 signals were quantified by measuring immunofluorescence intensity at vascular walls, and signal intensity along lines was determined using the ImageJ plot profile. Colocalization was quantified using Pearson’s correlation coefficient via the JACoP plugin in ImageJ.

Hematoxylin and Eosin (H&E) Staining

Mice were euthanized by cervical dislocation, and retinas were promptly isolated, fixed in 4% PFA, embedded in paraffin, and sectioned at a thickness of 4 µm. The paraffin-embedded eye sections were stained with hematoxylin and eosin (H&E) for histological examination. Images were captured using a light microscope (Olympus, Japan).

RT-qPCR Assessment

Total RNA was extracted from retinas and cultured cells using TRIzol reagent (Vazyme, Nanjing, China) following the manufacturer’s protocol. Complementary DNA (cDNA) was synthesized using the PrimeScript RT Reagent Kit (RR047A, Takara, Tokyo, Japan) for reverse transcription of mRNA. Quantitative real-time PCR (RT-qPCR) was performed with UltraSYBR Mixture (CW0957M, CW Biotech Co., Beijing, China) on the CFX96 RT-PCR system (CFX96 Optics Module; Bio-Rad, USA).

The sequences of mouse primers used were as follows:

GAPDH: Forward: 5′-AGGTCGGTGTGAACGGATTTG-3′; Reverse: 5′-TGTAGACCATGTAGTTGAGGTCA-3′
C3: Forward: 5′-CCAGCTCCCCATTAGCTCTG-3′; Reverse: 5′-GCACTTGCCTCTTTAGGAAGTC-3′
C1q: Forward: 5′-AAAGGCAATCCAGGCAATATCA-3′; Reverse: 5′-TGGTTCTGGTATGGACTCTCC-3′
iNOS: Forward: 5′-GTTCTCAGCCCAACAATACAAGA-3′; Reverse: 5′-GTGGACGGGTCGATGTCAC-3′
Arg-1: Forward: 5′-CCTGAAGGAACTGAAAGGAAAG-3′; Reverse: 5′-TTGGCAGATATGCAGGGAGT-3′

The sequences of human primers used were as follows:

GAPDH: Forward: 5′-GGAGCGAGATCCCTCCAAAAT-3′; Reverse: 5′-GGCTGTTGTCATACTTCTCATGG-3′
AQP4: Forward: 5′-TCAGCATCGCCAAGTCTGTC-3′; Reverse: 5′-CTGGGAGGTGTGACCAGATAG-3′
C3: Forward: 5′-GGCTGTCTTCTCTCAAGCA-3′; Reverse: 5′-GGGAATCTCACACATCACTCT-3′

GAPDH was used as a reference gene for normalization. Gene expression levels were calculated using the 2^−△△CT method.

Optical Coherence Tomography (OCT)

Retinal thickness was assessed using a retinal imaging system (isOCT 4D-ISOCT, Optoprobe, Burnaby, Canada). Mice were anesthetized, and their eyes were dilated with 0.5% tropicamide eye drops. A single rectangular scan consisting of 800 A-scans by 800 B-scans over a 2.5 × 2.5 mm area centered on the optic nerve head (ONH) was performed on each eye. Retinal thickness was measured at 0.30 mm from the ONH. Images were acquired and retinal thickness measurements were calculated using software (version 2.0) from OptoProbe Research Ltd.

Electroretinogram (ERG) and Flash Visual Evoked Potential (f-VEP)

Mouse flash visual evoked potentials (f-VEP) and electroretinograms (ERG) were recorded using an OPTO-III visual electrophysiology instrument (Optoprobe Science, Burnaby, BC, Canada) to assess visual function. Mice were kept in darkness for over 12 hours before the procedure, which was conducted under dim red light.

For ERG recording, a loop electrode was placed on the corneal surface of the eye being tested, while needle electrodes were inserted under the skin of the cheek and tail. For detecting flash VEPs, a needle electrode was inserted under the skin between the ears, replacing the loop electrode. Three stable waveforms were recorded for each eye (normal and injured) in each animal.

Cell Culture

MIO-M1 cells, an immortalized retinal Müller glial cell line, were obtained from Biopike Technology Company Ltd (Minnesota, USA) and cultured in Dulbecco's Modified Eagle Medium (DMEM) containing 4.5 g/L glucose, supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin/streptomycin, and 1.0 mM glutamine.

BV2 microglia were obtained from Procell Life Science & Technology Co. Ltd (Wuhan, China) and cultured in DMEM supplemented with 10% FBS and 100 U/mL penicillin/streptomycin. Both cell lines were maintained in a humidified incubator with 5% CO2 at 37°C. The culture medium was refreshed every 2 to 3 days until the cells reached confluency.

Small-Interfering RNA (siRNA) Knockdown of AQP4

Three siRNA sequences targeting human AQP4 were designed and synthesized by Hanheng Biological Co. Ltd (Shanghai, China). Non-targeting siRNA was used as a control (NC). The siRNA sequence with the highest silencing efficiency was selected for subsequent experiments (siAQP4 sequence: GGUUGAGUUGAUAAUCACATT).

Transfection was carried out using the RNAiMAX transfection reagent (No.13778075, Invitrogen) according to the manufacturer's instructions. MIO-M1 cells were seeded into a 6-well plate, and RNAiMAX reagent, along with 5 pmol siRNA, was diluted separately in Opti-MEM and then added to each well. Treatments were conducted 24 hours post-transfection.

Transwell Co-culture of Retinal Müller Cells and Microglia

To investigate the effects of activated Müller cells on retinal microglia, MIO-M1 cells (retinal Müller glial cells) and BV2 cells (microglia) were co-cultured using a Transwell system. MIO-M1 cells were seeded onto the wells of a 24-well plate, while BV2 cells were seeded onto the Transwell permeable support membrane inserts with an 8 µm pore size (Corning, USA), which were placed above the MIO-M1 cells.

MIO-M1 cells were exposed to either control IgG or patients' purified human-IgG (diluted 1/50 in culture media). After treatment, these cells were co-cultured with BV2 microglia for 24 hours. Following the co-culture period, the cells were washed with PBS. MIO-M1 and BV2 cells were then harvested for further mRNA analysis and immunofluorescent staining.

To assess cell migration, the BV2 cells on the lower surface of the Transwell filters were stained with 5% crystal violet, allowing visualization and analysis of the BV2 cells that had migrated through the membrane pores towards the MIO-M1 cells.

Cell Viability Assay (CCK-8)

The primary retinal ganglion cells were acquired from Procella Co., Ltd (Catalog No. CP-M122) for the C1q neurotoxicity assay. The cell viability was assessed using the Cell Counting Kit-8 (CCK-8, Dojindo Molecular Technologies, Japan) according to the manufacturer's instructions. Briefly, cells were seeded in 96-well plates at a density of 10⁶ cells per well and allowed to adhere overnight at 37°C in a humidified containing C3 treated BV2 culture medium (without or with 100µg/ml C1q neutralizing antibody ANX005) for 3 hours 10 µL of CCK-8 solution was added to each well, and the plates were incubated for 4 hours at 37°C. The absorbance at 450 nm was then measured using a SpectraMax® i3x Multi-Mode Microplate Reader (Molecular Devices). The absorbance values were directly proportional to the number of living cells, and cell viability was calculated as a percentage of the control group.

Statistical Analysis

Quantitative data were normalized to the control group and presented as a percentage of the control (%). The statistical significance of differences between groups was evaluated using an independent sample t-test when comparing two groups. For comparisons involving more than two groups, one-way ANOVA was employed. If the data were not normally distributed, the Mann-Whitney rank-sum test was used instead. Statistical analyses and the generation of graphs were performed using GraphPad Prism 8.0 software (GraphPad Software Inc., San Diego, CA, USA). Statistical significance was indicated as follows: *P < 0.05; **P < 0.01; ***P < 0.001.

Intravitreal NMO-IgG Induces AQP4 Loss in the Retina but Not in the Optic Nerve

The experimental timeline is outlined in Fig. 1a. Following the intravitreal injection of NMO-IgG, no hemorrhages or signs of intraocular inflammation were detected under light microscopy. Three days post-injection, the binding of patient-derived NMO-IgG to retinal cells was examined. Immunofluorescence in retinal flat mounts, using a secondary antibody against human-IgG (green), revealed NMO-IgG deposition across the entire retina, particularly along blood vessels (Fig. 1b). This pattern of NMO-IgG distribution aligns with previous studies[14, 15].

Further investigation using double immunofluorescent staining for AQP4 (red) and GS (green, a Müller cell marker) on cryosections of retinas treated with control-IgG showed that AQP4 is specifically expressed in Müller cells[16]. However, in retinas treated with NMO-IgG, a significant reduction in AQP4 expression was observed in Müller cells, without evidence of cell death (Figs. 1c and 1d).

In retinal flat mounts, double staining for AQP4 (red) and CD31 (green, an endothelial cell marker) confirmed that AQP4 was localized around blood vessels. The NMO-IgG treatment resulted in a noticeable loss of AQP4 in the perivascular region, as indicated by the line intensity profile of AQP4 and CD31 fluorescence (Figs. 1e and 1f).

Interestingly, despite the significant AQP4 loss in the retina, no such loss was observed in the optic nerves (Fig. 1g). This suggests that intravitreal injection of NMO-IgG does not penetrate the optic nerves, thereby not affecting AQP4 expression in this region.

Intravitreal NMO-IgG Produces Retinal Vascular Leakage and Alterations in Retinal Thickness

Given the significant deposition of NMO-IgG in the perivascular zone, we examined retinal blood vessel permeability in the presence or absence of NMO-IgG (Fig. 2a). Evans Blue dye was injected into the tail vein, and retinal flat mounts were examined on days 7 and 14 after human IgG administration. HC-IgG did not induce any blood vessel hyperpermeability on either day 7 or day 14. However, NMO-IgG caused significant leakage of retinal vessels on day 7, which partially recovered by day 14 but remained detectable (Fig. 2a).

AQP4 loss and blood leakage indicated abnormalities in water transport throughout the retina. We then investigated the effect of NMO-IgG on the thickness of the retinal layers. OCT images showed that, on day 7 after NMO-IgG injection, the total retinal thickness was substantially increased, particularly in the RNFL, IPL, and OPL (Fig. 2b). H&E staining of retinal sections also revealed similar results in the IPL and OPL (Figs. 2j-k). This altered thickness gradually reverted, with a slight but significant decrease in RNFL thickness by day 21. Notably, we did not observe papillary edema in any of the retinas throughout the study (Fig. 2l).

Intravitreal NMO-IgG Contributes to RGC Loss and Visual Dysfunction

The swelling of retinal layers indicated potential RGC injury in the NMO-IgG-treated group. To assess RGC loss, we performed immunofluorescent staining for the RGC-specific marker RBPMS on whole-mount retinas. There was no difference in RGC density between the NMO-IgG group and the control IgG group on day 7 after IgG injection, suggesting that NMO-IgG did not induce RGC injury at the early stage (Figs. 3a-b). However, by day 21, we observed a significant decrease in RGC density in the NMO-IgG group, leading to secondary neuronal injury (Figs. 3a-b).

We then recorded ERG and VEP waveforms at different time points to evaluate retinal function. On day 7, dark-adapted (scotopic) ERG showed a small decrease in b-wave amplitude and preservation of a-wave amplitude in the NMO-IgG group compared to the control, indicating minor optic nerve dysfunction. By day 21, both a-wave and b-wave amplitudes in the light-adapted (photopic) and dark-adapted ERG were significantly decreased, coinciding with severe RGC injury observed in staining. F-VEP results also showed a significant decrease in wave amplitude on day 21, confirming NMO-IgG-dependent retinopathy initiated in Müller cells and subsequently spreading to RGCs (Fig. 3c).

Microglial Inflammation Contributes to NMO-IgG-Induced Retinopathy

The swelling of retinal layers and dysfunction of RGCs indicated potential inflammation in the retina after NMO-IgG injection. Given that microglia are the primary immune cells in the retina, we hypothesized their critical role in NMO-IgG-induced retinopathy. Iba1 immunostaining revealed significant microgliosis following NMO-IgG injection. Furthermore, mRNA analysis showed a significant increase in the M1 microglial polarization marker iNOS on day 7, suggesting an inflammatory role for microglia (Figs. 4a-e).

To confirm the contribution of M1 microglia to neuronal injury in the NMO-IgG-treated retina, we performed microglia depletion using the CSF1 receptor antagonist PLX3397 (Fig. 4f). Iba1 immunostaining demonstrated that microglia were completely ablated after 7 days of feeding with PLX3397 chow (Fig. 4g). Interestingly, microglial ablation significantly reduced NMO-IgG-induced RGC loss, suggesting that microglial suppression may provide protection against retinal ganglion cell loss following NMO-IgG transfer (Figs. 4h-i).

Müller Cell-Derived Complement C3 Activates Microglia Following NMO-IgG-Induced AQP4 Internalization

In previous studies, we reported that NMO-IgG induces AQP4 internalization and upregulates complement C3 expression in astrocytes[12, 13]. To further investigate this process in the retina, we examined complement C3 expression in an NMO-IgG intravitreal injection model. RT-qPCR analysis over a 21-day period revealed that C3 expression peaked on day 7 post-injection and then gradually decreased (Fig. 5a). Consistently, immunofluorescence staining showed clear co-localization of GS and C3 on day 7 after NMO-IgG injection, confirming that AQP4 internalization in Müller cells led to increased complement C3 production (Figs. 5b-c).

Given that complement C3-activated microglia induce evolving NMO lesions in the spinal cord, we hypothesized that retinal microglia might be activated by Müller cell-derived C3. To test this, we treated the Müller cell line MIO-M1 with NMO-IgG and examined AQP4 expression. Consistent with the in vivo model, AQP4 staining quantification showed a dramatic decrease 24 hours after NMO-IgG treatment (Figs. 5d-e, upper panel). Complement C3 protein expression was significantly elevated in MIO-M1 cells treated with NMO-IgG compared to the PBS and HC-IgG groups (Fig. 5e, lower panel).

Additionally, NMO-IgG-treated MIO-M1 cells induced robust BV2 cell migration compared to control IgG (Figs. 5g-i). This NMO-IgG-dependent Müller cell-microglia communication also increased iNOS expression in BV2 cells, indicating an inflammatory response (Fig. 5f). To confirm that Müller cell-microglia communication was AQP4-IgG-specific, we transfected AQP4 siRNA into MIO-M1 cells to downregulate AQP4 expression. The AQP4 siRNA effectively reduced AQP4 expression in MIO-M1 cells (Fig. 5h), which also alleviated NMO-IgG-induced microglia migration in MIO-M1/BV2 co-cultures (Figs. 5i-j).

C1q Derived from Activated Microglia Is Neurotoxic to RGCs in NMO-IgG-Induced Primary Retinopathy

The complement pathway initiator C1q is primarily released by activated microglia. In the retina treated with NMO-IgG, C1q mRNA levels significantly increased by day 7 (Fig. 6a). We also observed C1q protein within microglia located in the NMO-IgG-induced retinal lesion (Figs. 6b-c). Remarkably, the depletion of microglia using PLX3397 completely inhibited NMO-IgG-induced C1q expression (Fig. 6d). However, this depletion did not affect the expression of complement C3, which is produced by Müller cells (Figs. 6e-f).

To further investigate the role of C1q in retinal neurotoxicity, we explored whether C1q alone could induce neurotoxicity in a cell culture model. When primary cultured mouse RGC cells were treated with 100 µg/ml of mouse C1q, significant RGC cell damage was observed, as measured by the CCK8 assay (Fig. 6g), indicating C1q-dependent cytotoxicity. Additionally, the C1q-neutralizing antibody ANX005 reduced C1q-induced RGC cell injury in a dose-dependent manner (Fig. 6h).

Clinical evidence suggests that AQP4-IgG seropositive NMOSD patients without a clinical history of optic neuritis (ON) exhibit lower macular RNFL and GCIPL thickness and faster rates of peripapillary RNFL (pRNFL) thinning compared to healthy controls[17]. Foveal thickness is also reduced in NMOSD patients who have not experienced visual symptoms[18]. However, another cohort study revealed that while macular outer plexiform layer (OPL) and outer nuclear layer (ONL) loss was not specific to AQP4-IgG seropositive NMOSD, reductions in the retinal nerve fiber layer (RNFL) and ganglion cell-inner plexiform layer (GCIPL) were still observed[19].

These clinical findings raise the question of whether primary retinopathy exists in NMOSD[20]. Clinically, it is challenging to rule out the presence of possible asymptomatic subclinical optic nerve lesions and trans-synaptic degeneration in the affected eyes. Mechanistically, AQP4 is polarized in the membrane domains of astrocytes and Müller cells, which face the vitreous body and blood vessels in the retina[21]. It is possible that NMO-IgG increases vascular permeability, promoting its transferability and allowing it to penetrate deep into the retina, cross the blood-retinal barrier, and bind to AQP4. Therefore, clinical evidence of AQP4-IgG in intraocular fluid (aqueous humor and vitreous liquid) remains necessary to understand the impaired retinal homeostatic mechanisms in NMOSD.

The passive transfer of AQP4-IgG into the central nervous system via intrathecal, intravenous, and intracranial injection has been shown to induce NMO-like pathology[8, 22–25]. While ON and myelitis are potential outcomes of these models, their impact on the retina is limited. A 2016 study by Felix et al. reported that intravitreal injection of rAb-53 in a rat model demonstrated that AQP4-IgG can cause primary RGC loss and thinning of the ganglion cell complex through a complement-independent pathway[15]. Zeka et al. found that AQP4 loss in Müller cells correlated with T cell infiltration and microglia activation/macrophage recruitment but not with antibody-dependent cellular cytotoxicity (ADCC) or complement-dependent cytotoxicity (CDC)[26, 27].

In our study, we developed a new intravitreal injection mouse model without exogenous complement to further explore retinopathy. A clinical study revealed that in NMO patients, the complement cascade mediated cell death has no difference during the disease onset and recovery phase[28]. In addition, human NMO-IgG has limited ability to activate the mouse complement system[29]. Therefore, we hypothesized that pathology and functional impairment are not induced by complement-mediated cytotoxicity (CDC). Our observations revealed significant AQP4 loss in NMO-IgG-treated retinas without evidence of Müller cell death. This finding contradicts the idea of Müller cell apoptosis and supports the notion of AQP4 internalization following IgG binding[21]. NMO-IgG-induced AQP4 internalization could potentially reduce complement deposition on the cell surface, thereby protecting Müller cells from CDC[10].

Microglia activation is another significant pathological feature observed in our experimental mice. Microglia activity was also reported in a rat model of AQP4-IgG induced optic neuritis model[30]. We previously demonstrated that in a mouse NMO spinal cord model, astrocytic C3 activates microglia via microglial C3a receptors[22, 31]. The C3-C3aR pathway, which mediates microglia-astrocyte interaction, has recently been described in demyelinating diseases and neurodegenerative disorders[32, 33]. The upregulation of C3, specifically associated with neurotoxicity in both the retina and spinal cord, drives microglia to release pro-inflammatory cytokines[34]. Consistently, in the retina, NMO-IgG induced complement C3 expression in Müller cells, leading to microglia-Müller cell interaction[33].

This microglia-Müller cell interaction coincided with significant retinal ganglion cell (RGC) loss and blood-retinal barrier (BRB) hyperpermeability following NMO-IgG treatment. The observed RNFL thinning and INL thickening in this model align with findings from our previous clinical study[5]. Specifically, our prior work showed that individuals positive for AQP4-IgG antibodies had thinner peripapillary RNFL and thicker INL compared to those negative for the antibodies[35, 36]. The fluctuation in inner retinal layer thickness in this model may be associated with the loss of perivascular AQP4 and subsequent vascular leakage. Previous studies have reported subclinical changes in parafoveal vessels, suggesting that the intermediate vascular plexus in the INL and the deep vascular plexus in the ONL could be involved at an early stage of NMOSD[35, 37].

Although C3-expressing astrocytes have been reported to be neurotoxic[38], our study suggests that microglia activation is crucial in NMO-IgG-induced retinopathy. Microglia ablation prevented NMO-IgG-induced neuronal damage, indicating that the interaction between activated microglia and Müller cells plays a central role in retinal injury. On the other hand, reactive microglia produce IL-1α, TNF, and C1q, which can sufficiently induce the A1 astrocyte phenotype[39, 40]. Astrocyte dysfunction results in downregulation of chemokines and growth factors which maintain the homeostasis of central nerves system[41, 42]. Notably, in our model, C3 expression in the non-ablated group was much higher than in the microglia-ablated group, suggesting a positive feedback loop between activated microglia and Müller cells.

Activated microglia upregulated complement C1q, the initial complement protein[43]. C1q has been reported to bind to mitochondria, block the respiratory chain, and induce neuronal oxidative stress[44]. Additionally, C1q can bind to endothelial cells, leading to BRB hyperpermeability[45]. Therefore, microglial C1q might induce RGC cell loss and retinal vessel leakage in this model (Fig. 7).

Another pathological observation in our study is that NMO-IgG induced Müller cell dysfunction. As part of the retinal neurovascular unit, Müller cells are essential for maintaining retinal homeostasis and regulating the progression of retinal inflammation[14, 46]. The binding of autoantibodies to AQP4 mediates partial AQP4 internalization before complement cascade initiation, leading to reduced plasma membrane water permeability. This impairment in cell volume regulation results in adverse swelling under stressful conditions in the retina. Another study showed that in AQP4 ^-/- mice, a sustained thickening of the retina, primarily due to changes in the inner retinal layers, was observed during recovery from experimental autoimmune encephalomyelitis (EAE)[46–48]. Subsequently, this thickening dropped below its baseline level. Müller cell dysfunction in AQP4 ^-/- mice results in structural damage to the gliovascular unit of the retina and promotes scar formation during CNS inflammation[46]. Therefore, we propose that the thickness changes in the intravitreal NMO-IgG infusion model are mainly driven by Müller cells. (Fig. 7)

There are still several limitations to our experiments. First, the long-term effects of NMO-IgG in the retina remain unclear. Further evaluation and observation over an extended period, along with detailed pathological studies, are required. Additionally, our experimental results, such as the in vivo expression levels of C3, cannot definitively rule out the contribution of astrocytes. Furthermore, C3aR (complement C3a receptor) antagonists and C3aR knockout mice should be utilized in future research to further elucidate the mechanisms underlying Müller cell-microglia interaction. The pathogenesis of retinal abnormalities in NMOSD may be multifactorial and more complex than a simple pathogenic role of NMO-IgG, warranting further investigation.

In conclusion, NMO-IgG induces primary retinopathy through a complement-independent pathway, with microglia activation playing a critical role in NMO pathogenesis. Müller cell-derived C3 is an upstream factor in microglial C1q release, and restricting microglia activation could be a novel therapeutic target for NMO-IgG-induced retinopathy.

NMO	Neuromyelitis optica
AQP4	Aquaporin-4
GFAP	Glial fibrillary acidic protein
GS	Glutamine synthetase
Iba1	Ionized calcium-binding adaptor molecule-1
RBPMS	RNA-binding protein with multiple splicing
OCT	Optical coherence tomography
ERG	Electroretinography
f-VEP	Flash visual evoked potential
RT-qPCR	Real-time fluorescence quantitative PCR
CSF1R	Colony stimulating factor 1 receptor
Arg-1	Arginase-1
iNOS	Inducible nitric oxide synthase
PFA	Paraformaldehyde
H&E	Hematoxylin and eosin
RNFL	Retinal nerve fiber layer
SD-OCT	Spectral-domain optical coherence tomography
RGCs	Retinal ganglion cells
INL	Inner nuclear layer
IPL	Inner plexiform layer
ONL	Outer nuclear layer
OPL	Outer plexiform layer
DMEM	Dulbecco’s modifed eagle medium
FBS	Fetal bovine serum

Ethics approval and consent to participate

All experimental procedures described in this article were reviewed and approved by the Ethics Committee of Chinese PLA General Hospital.

Consent for publication

Not applicable.

Availability of data and materials

All data generated or analyzed during this study are included in this published article.

Competing interests

The authors declare that they have no competing interests.

Funding

This project was supported by National Natural Science Foundation of China (No.82101110).

Authors’ contributions

T.C and S.W conceived the study and coordinated the project. B.C and H.Z designed the experiments. B.C, H.S and M.S performed the experiment. Q.X and Y.L analyzed the data. Y.Y, Q.L and M.G reviewed the manuscript. All authors read and approved the final manuscript.

Acknowledgments

The schematics in this paper are constructed in BioRender.

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NMO-IgG causes primary retinal damage without optic nerve injury

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Version 1

Abstract

Background

Methods

Results

Conclusions

Figures

Background

Materials and methods

Cell Viability Assay (CCK-8)

Statistical Analysis

Results

Intravitreal NMO-IgG Induces AQP4 Loss in the Retina but Not in the Optic Nerve

Intravitreal NMO-IgG Produces Retinal Vascular Leakage and Alterations in Retinal Thickness

Intravitreal NMO-IgG Contributes to RGC Loss and Visual Dysfunction

Microglial Inflammation Contributes to NMO-IgG-Induced Retinopathy

Müller Cell-Derived Complement C3 Activates Microglia Following NMO-IgG-Induced AQP4 Internalization

C1q Derived from Activated Microglia Is Neurotoxic to RGCs in NMO-IgG-Induced Primary Retinopathy

Discussion

Abbreviations

Declarations

References

Additional Declarations

Status:

Version 1