Regulatable Complement Inhibition of the Alternative Pathway Mitigates Wet Age-Related Macular Degeneration Pathology in a Mouse Model

Abstract

Purpose: Risk for developing age-related macular degeneration (AMD) is linked to an overactive complement system. In the mouse model of laser-induced choroidal neovascularization (CNV), elevated levels of complement effector molecules, including complement C3, have been identified, and the alternative pathway (AP) is required for pathology. The main soluble AP regular is complement factor H (fH). We have previously shown that AP inhibition via subretinal AAV-mediated delivery of CR2-fH using a constitutive promoter is efficacious in reducing CNV. Here we ask whether the C3 promoter (pC3) effectively drives CR2-fH bioavailability for gene therapy.

Methods: Truncated pC3 was used to generate plasmids pC3-mCherry/CR2-fH followed by production of corresponding AAV5 vectors. pC3 activation was determined in transiently transfected ARPE-19 cells stimulated with H₂O₂ or normal human serum (+/− antioxidant or humanized CR2-fH, respectively). CNV was analyzed in C57BL/6J mice treated subretinally with AAV5-pC3-mCherry/CR2-fH using imaging (optical coherence tomography [OCT] and fundus imaging), functional (electroretinography [ERG]), and molecular (protein expression) readouts.

Results: Modulation of pC3 in vitro is complement and oxidative stress dependent, as shown by mCherry fluorescence. AAV5-pC3-CR2-fH were identified as safe and effective using OCT and ERG. CR2-fH expression significantly reduced CNV compared to mCherry and was correlated with reduced levels of C3dg/C3d in the retinal pigment epithelium/choroid fraction.

Conclusions: We conclude that complement-dependent regulation of AP inhibition ameliorates AMD pathology as effectively as using a constitutive promoter.

Translational Relevance: The goal of anticomplement therapy is to restore homeostatic levels of complement activation, which might be more easily achievable using a self-regulating system.

Introduction

Age-related macular degeneration (AMD) is a slowly progressing blinding disease that has two forms, dry AMD, which can advance to geographic atrophy (GA), and neovascular, or wet AMD, that is characterized by abnormal blood vessel growth underneath the macula at the posterior pole. Wet AMD accounts for ∼10% to 15% of AMD cases, and dry AMD accounts for 85% to 90% of cases. AMD affects approximately 15 million people in the United States alone and is expected to double by the year 2050.¹ AMD is a complex disease that has risk factors such as aging, smoking, genetic inheritance, race, sex, cardiovascular disease,² and polymorphisms in complement genes such as complement component 2,³ factor B,³ complement component 9,⁴ factor I, complement component C3 (C3),⁴ and complement factor H (fH),Y402H.⁵ The complement system is part of the immune system responsible for eliminating invading pathogen (non-self cells) and is thus activated early at sites of injury or infection, while self-cells are protected based on their expression of negative regulators. Complement can be activated by three pathways, lectin pathway, classical pathway, and alternative pathway (AP), which share a central protein, C3. Interestingly, complement dysregulation is believed to be associated with AMD pathobiology.⁶ Complement dysregulation refers to the central complement protein, C3, undergoing excessive cleavage by C3 convertases, which leads to a buildup of downstream biological effectors. Effector molecules such as anaphylatoxins, opsonins, and the membrane attack complex (MAC) can bind to self-tissues, initiating cellular damage responses, lysis, or immune cell–mediated phagocytosis.⁷ Notably, control of the amplification loop of the AP is crucial, as it is typically responsible for 80% to 90% of C3 cleavage products.⁸ Importantly, the AP is inhibited by circulating complement protein fH. Taken together, genetic and molecular studies have identified the complement system as an important component in AMD.⁶ 

Currently, the only treatment readily available for early and intermediate AMD are AREDS and AREDS-2, supplemental vitamins, and minerals, often prescribed to patients to impede dry AMD pathology based on the two NEI Age-Related Eye Disease Studies (AREDS and AREDS-2). To date, one treatment is also available for the treatment of GA. Apellis (Waltham, MA, USA) has received US Food and Drug Administration (FDA) approval for the pegylated compstatin peptide (pegcetacoplan; SYFOVRE),⁹ and Iveric (Parsippany, NJ, USA) has submitted their New Drug Application (NDA) for a pegylated anti-C5 aptamer (Avacincaptad Pegol). Both approaches lead to a slow reduction in GA growth over the study period¹⁰ but currently with no discernable benefit for the vision of the patient. Similarly, patients with neovascular AMD are currently being treated with intravitreal injections of anti–vascular endothelial growth factor (VEGF)–based therapeutics such as Lucentis (ranibizumab; Genentech), Avastin (bevacizumab; Genentech, South San Francisco, CA, USA), and Eylea (aflibercept; Regeneron, Tarrytown, NY, USA) every 4 to 12 weeks to mitigate vascular growth.¹¹ There is evidence that complement polymorphisms play a role in not only dry AMD but neovascular AMD as well, specifically the single-nucleotide polymorphism in fH protein (Y402H).¹² We have previously shown that complement activation drives VEGF_A production and secretion from human ARPE-19 cells, which could be prevented by AP inhibition.¹³ Likewise, in a mouse model of wet AMD, AP inhibition mitigated complement activation, VEGF_A bioavailability, and choroidal neovascularization (CNV) lesion size¹⁴ and fibrosis.¹⁵ Taken together, there is evidence for a significant role for overactive complement in neovascular AMD pathogenesis. 

A major barrier for developing new therapies in AMD is a lack of understanding the role of complement dysregulations. Regulating the complement activation at the optimal level is currently being attempted with numerous different drug approaches in several diseases. One such disease is paroxysmal nocturnal hemoglobinuria (PNH), a hematologic disease caused by a dysregulation of MAC due to a lack membrane-bound complement regulators CD55 and CD59 that leads to cytolytic pore formation and hemolysis of red blood cells. Eculizumab, an anti-C5 antibody, was a game-changing therapeutic approved by the FDA for the treatment of PNH and the first treatment of its kind to provide clinical validation that blocking the complement cascade can foster life-altering results.¹⁶ Most of the anticomplement approaches for complement inhibition in AMD involve blocking specific proteins of the complement cascade using antibodies, peptides, or aptamers. However, as the complement system is a surveillance system, with only a fraction of the complement components being actively engaged in complement activation, these inhibitors will bind to all their target complement proteins. This means inhibitors will be wasted on complement proteins that are not actively engaged in complement activation and tissue damage. 

Thus, when designing a complement inhibitor, one consideration is how to design an inhibitor that only targets complement proteins that are actively engaged in pathophysiologic complement activation. Importantly, membrane-bound as well as fluid phase complement activation can occur to opsonize self-tissue. Endogenously, cells are protected by expressing several membrane-bound complement inhibitors such as CD59 glycoprotein, membrane cofactor protein (CD46), and CD55. Additionally, there are fluid phase inhibitors like C4b binding protein, vitronectin, and complement protein fH. Factor H is an abundant complement inhibitory protein responsible for controlling the fluid phase and membrane-bound tickover of the alternative pathway by binding C3b and is made up of 20 short consensus repeats (SCRs). Membrane-bound fH-mediated inhibition in ocular tissues requires targeting to glycosaminoglycans and other polyanions via SCRs 6–8, with the Y402H present in SCR7 reducing binding.¹⁷ Our strategy is to target complement protein fH to pathologic tissues where it is needed to reduce AP activation but to do so in a SCR 6–8 independent manner. Specifically, targeting is provided using the complement receptor 2 (CR2) domain that recognizes C3-based opsonins (iC3b, C3d, and C3dg),¹⁸ and AP inhibition is provided using only the inhibitory domain of fH (CR2-fH).¹⁹ Thus, CR2-fH comprises SCR 1–4 of CR2 and SCR 1–5 of fH.^18,19 This inhibitor has been extensively tested in cell-based models of retinal pigment epithelium (RPE) damage^20,21 as well as in vivo, in mouse models of wet¹⁴ and dry AMD²² and using systemic delivery,²³ cell encapsulation technology,²⁴ and gene therapy.²⁵ 

Previously, we have tested adeno-associated viral vector serotype 5 (AAV5) for delivery of CR2-fH.²⁵ AAV5 was chosen for its low immunogenicity and nonpathogenicity as well as its higher packaging capacity (8.9 kb) and long-lasting therapeutic gene expression.^26,27 In the mouse eye, we delivered the CR2-fH construct with an RPE cell-specific promoter, vitelliform macular dystrophy 2 (VMD2),²⁵ using transcorneal subretinal delivery to target the RPE. Using this vector, we were able to show that CR2-fH secreted by the RPE was sufficient to reduce CNV back to similar levels afforded by systemic delivery in a mouse model of wet AMD.²³ 

To develop therapeutic complement inhibition, complement activation needs to return to homeostatic levels. While overactivation leads to disease and may induce effector cells to harm host tissue,²⁸ too much complement inhibition may be detrimental for microbial recognition, cellular damage, and removal of debris via phagocytosis. To control for levels of CR2-fH expression of the gene cassette (i.e., expression limited to times of complement activation), we propose to drive gene expression using the C3 promoter (pC3) described by Cho et al.²⁹ This decision was based in part on our previous in vitro data using ARPE-19 cells.²⁰ In this study, we showed that C3 messenger RNA (mRNA), C3 protein secretion, and C3a production were increased in the presence of a cellular stressor (cigarette smoke exposure) known to generate oxidative stress and activate complement, as well as reduced to control levels by pretreatment with CR2-fH, C3a receptor antagonist, or antioxidant N-acetyl cysteine.^20,30 Similarly, it has been shown that ARPE-19 cells in the presence of H₂O₂ significantly upregulated C3 mRNA and protein production.³¹ Finally and most important, we have shown in the mouse CNV model that C3 mRNA production is decreased in the presence of CR2-fH.¹⁴ Based on these results, we hypothesized that the stress-inducible C3 promoter would be beneficial in driving the CR2-fH therapeutic protein cassette, pC3-CR2-fH. 

Here we investigate the efficacy and safety of a subretinally injected AAV vector containing a C3-inducible promoter regulating CR2-fH expression in the murine model of wet AMD. In this study, we expand upon our previous work by allowing a homeostatic relationship between pathology and complement activation. 

Materials and Methods

C3 Promoter Plasmids and Adeno-Associated Vectors

The C3 promoter (pC3) (−1005 to +251) was generously provided by the University of Texas MD Anderson Cancer Center under a Material Transfer Agreement and is described in Cho et al.²⁹ This C3 promoter fragment has previously been shown to drive expression of luciferase under stress conditions in malignant ovarian epithelial cells.²⁹ The fusion protein CR2-fH composition, used as previously described,²³ is made up of SCRs encoding the 4 N-terminal region of mouse CR2 (mature protein residues 1–257; RefSeq: M35684), linking the CR2 domain via a (G₄S)₂ linker to the 5 N-terminal SCRs of mouse fH (mature protein residues 1–303; RefSeq: NM009888). The University of Florida's Ocular Gene Therapy Core generated plasmids pC3-mCherry and pC3-CR2-fH by cloning the C3 promoter into a pTR backbone driving mCherry and CR2-fH expression, respectively, as described previously.¹⁴ These plasmids were used to generate serotype 5 adeno-associated viral vector constructs, AAV5-pC3-mCherry and AAV5-pC3-CR2-fH (Supplementary Fig. S1). 

Cell Culture Experiments

Plasmid Transfections

Human ARPE-19 (ATCC CRL-2302; American Type Culture Collection, Manassas, VA, USA) cells were cultured in Dulbecco's modified Eagle's medium (DMEM) cell culture media (Gibco/Thermo Fisher Scientific, Waltham, MA, USA) containing high-glucose DMEM with D-glucose (4.5 g/L), L-glutamine, and sodium pyruvate (110 mg/L). Penicillin and streptomycin were added to media (1×) during cell growth but removed from transfection media. Cell media contained 10% fetal bovine serum. Cells (passage <10) were expanded in T75 cell culture flasks at 37°C, in the presence of 5% CO₂. Confluent cells were trypsinized with 0.05% trypsin (Gibco), and equal numbers of cells were seeded on 12-well plates (Costar/Thermo Fisher Scientific, Waltham, MA, USA), reaching 70% confluence within 24 hours. Cells were transfected using 100 µL of a master mix in Opti-MEM (Gibco) media, containing a transfection control plasmid (cytomegalovirus promoter driving green fluorescent protein expression, pCMV-GFP; Addgene, Watertown, MA, USA) and plasmid pTR-pC3-mCherry in equal molar concentrations with lipofectamine (1:2 ratio). Cells were exposed to plasmids for 8 hours, followed by media exchange to allow for 48-hour cellular recovery. 

Treatments

To provide evidence that pC3 drives mCherry expression under stress conditions when compared to control conditions, transfected ARPE-19 cells were washed with serum-free media (SFM) (Gibco) and left untreated or stressed with 0.1 mM H₂O₂ (Sigma-Aldrich, St Louis, MO, USA) or H₂O₂ in the presence of 10% complement-sufficient normal human serum (NHS; Complement Technologies, Tyler, TX, USA) for 4 hours. An equal number of cells were pretreated with antioxidant NAC (Sigma-Aldrich) or the AP inhibitor TT30 (humanized version of CR2-fH; generously provided by Yi Wang, Alexion Therapeutics, Boston, MA, USA) for 1 hour prior to stressor treatment. Following treatment, media were removed, and cells were washed with SFM three times and left in SFM for 36 hours to allow for expression of the reporter construct. After 36 hours, cells were stained with DAPI for 10 minutes, washed with SFM three times, and fixed in 4% paraformaldehyde for 20 minutes. Fixed cells were washed in phosphate-buffered saline solution and photographed using the TRITC channel (Olympus 1 × 73 Research Inverted Microscope equipped with cellSens imaging software). Images were analyzed on ImageJ software (Wayne Rasband, National Institutes of Health, Bethesda, MD, USA) using the threshold plugin to determine the gray value of fluorescent cells in each image. 

Viral Vector Injection

All animal experiments were performed in accordance with the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research and approved by the University Animal Care and Use Committee. C57BL/6J male and female mice (Jackson Laboratory, Bar Harbor, ME, USA) were generated from a colony established at the Medical University of South Carolina (MUSC) to ensure mice were all exposed to the same microenvironment. Subretinal injections were performed using the transcorneal subretinal injection route as described in Qi et al.³² Mice 8 to 10 weeks of age were injected under dissection microscope observation at 14× magnification according to published protocols^25,33,34 after anesthesia with xylazine (NDC 46066-750-02; Pivetal, Liberty, MO, USA) and ketamine (NDC 0143-9509-10; Hikima Pharmaceuticals, Berkeley Heights, NJ, USA) at 20 and 80 mg/kg, respectively. Pupils were dilated with 2.5% phenylephrine hydrochloride (NDC 17478-201-15; Akorn, Lake Forest, IL, USA) and 1% atropine sulfate (NDC 17478-215-15; Akorn), and the ocular surface was anesthetized with 0.5% proparacaine hydrochloride (NDC 61314-016-01; Alcon Laboratories, Fort Worth, TX, USA) and lubricated with 2.5% Goniotaire (NDC 59390-182-13; Altaire Pharmaceuticals, Aquebogue, NY, USA). As previously described,²⁵ a 30-gauge needle was used to puncture the limbus of mouse eyes to create a guide hole, through which a Hamilton blunt-tip needle on a Hamilton 2.5-µL syringe (Hamilton Co., Reno, NV, USA) was inserted. The Hamilton syringe was stabilized using a micromanipulator (M-125; Narishige International USA, Amityville, NY, USA),³⁴ allowing for a slow and steady injection of viral vectors (1 µL of 2.41 × 10¹¹ viral genomes [vg]/mL over 30 seconds’ duration). Injection of 1 µL leads to 60% to 90% retinal detachment. Postinjection antibiotic ointment neomycin polymyxin B sulfate and dexamethasone ointment (NDC 61314-631-36; Sandoz, Princeton, NJ, USA) was applied to prevent infection. In a previous study, driving expression of the AP inhibitor CR2-fH from an RPE-specific promoter (VMD2) at a concentration of 3 × 10¹¹ vg/mL was found to be both safe and efficacious.²⁵ A dose-finding study for AAV5-pC3-CR2-fH across 3 log units revealed that the same concentration of 2.41 × 10¹¹ vg/mL is appropriate and used throughout the study. 

Full-Field Electroretinography

Full-field electroretinography (ffERG) described previously³⁵ was performed at 1 month after subretinal injection.²⁵ Mice were dark adapted overnight and anesthetized, and pupils were dilated as described above. Celluvisc lubricant eye gel (Allergan, Irvine, CA, USA) was applied to allow electrical conductivity between corneal loop electrodes and the cornea. Needle electrodes were inserted in the upper snout and tail region to provide a reference and ground, respectively. GenTeal tear lubricating drops (Alcon Laboratories) were applied after testing to keep mouse eyes moist. The UTAS E-4000 System (LKC Technologies, Gaithersburg, MD, USA) was utilized to examine neuroretina and RPE function. Using this system, a-wave and b-wave amplitudes were recorded in response to 10-µs white light intensity flashes of −30 to 0 dB of attenuation (maximum light intensity in the dome of 2.48 photopic cd*s/m²) to examine photoreceptor and inner retina function.^36,37 C-wave amplitudes were recorded in response to a single 100-cd*s m² flash. All amplitudes were measured from baseline to the maximum peak. 

Choroidal Neovascularization

Here we used the mouse model of laser-induced choroidal neovascularization. This model has been confirmed to produce new vessels using multiple in vivo and ex vivo techniques by Campa et al.³⁸ and be dependent on VEGF^38,39 and complement activity.¹⁴ One month after subretinal injection, a Coherent ophthalmic laser (Coherent, Santa Clara, CA, USA) was used to photocoagulate (100-µm spot size, 532 nm, 100 mW, and 0.1-second duration) Bruch's membrane (BrM) at four equidistant locations around the optic nerve, as described in Annamalai et al.²⁴ Inclusion/exclusion criteria for successful lesion formation, such as bubble formation at the time of laser pulse administration and lack of vessel rupture, were adopted.⁴⁰ Five days following laser-induced CNV, lesions in mouse eyes were imaged by optical coherence tomography (OCT). Mice were sacrificed on day 6 post-CNV lesions to collect retina and RPE/choroid/sclera complexes (referred to as RPE/choroid) tissues. 

OCT Imaging

OCT was used to quantify retinal layer thickness and analyze CNV lesion sizes on day 5 after laser treatment.^15,25,35 Mice were anesthetized and pupils were dilated as described under ffERG. Mouse eyes were kept hydrated using GenTeal lubricating eye drops. The Bioptigen Spectral Domain Ophthalmic Imaging System (SD-OCT; Bioptigen, Durham, NC, USA) was used with a mouse retina lens with a 50° field of view. For retinal layer thickness analysis, images were taken centered on the optic nerve, with volumetric images set at 1.8 × 1.8 mm. The three-dimensional volume high-resolution images consist of 1000 A scans per B scan, collecting 11 B-scans averaging 5 frames per scan. Bioptigen InVivoVue Diver 3.4.4 Software⁴¹ was used on each image to automatically quantify separate retinal layer thicknesses throughout the OCT image, with each analysis generating 1000 × 11 × 5 = 55,000 A-lines × 9 boundaries to give 495,000 output points. Diver is designed to identify boundaries along the depth of each A-line and to exclude data from the optic nerve head area or B-scans where boundary identification fails. 

CNV lesions were analyzed in en face OCT images comprising 1000 A- and 100 B-scans, drawing a midline through the center of the hyperreflective spots representing RPE BrM rupture, with the axial interval positioned at the RPE/choroid complex level.⁴² Horizontal calipers set at 100 µm were placed at the site of each hyperreflective lesion. Spots were then bordered, and the area was measured using ImageJ software. The size of the pixels 1.8 × 1.8 mm was used to calculate lesion sizes (µm²). As Giani et al.⁴² have shown a “strong correlation between CNV volume and cross-sectional area measured on SD-OCT” on day 7 after induction of the lesions (correlation coefficient r = 0.936, P < 0.001), SD-OCT lesion volumes were not reconstructed. Finally, for each lesion, B-scans were analyzed for the presence of small, medium, or large fluid domes or the absence thereof as described.⁴² Specifically, a B-scan cross section at the center of each lesion’s OCT image was assigned a severity fluid dome score that ranged from 0 (absent), 1 (small), 2 (medium), to 3 (large) and an average severity score per mouse established. 

Fundus Photography

En face fundus images were taken of subretinally injected mouse eyes with the Micron III imaging system (Phoenix Research Labs USA, Pleasanton, CA, USA) to ensure retinal reattachment 1 month after retinal detachment. Before imaging, mice were anesthetized, and pupils were dilated as described in the ffERG section. 

Western Blot

Mouse eyecups (RPE/choroid/sclera complexes) were collected for protein extraction from control animals (no CNV, no vector injection) and CNV animals 1 month after vector injections (AAV5-pC3-mCherry or AAV5-pC3-CR2-fH) and day 6 after laser-induced CNV. Whole tissues were sonicated and protein extracted by solubilization in radioimmunoprecipitation assay buffer (Thermo Fisher Scientific, Waltham, MA, USA) containing a protease inhibitor cocktail (Sigma-Aldrich). Tissue lysates were centrifuged (20,000 × g for 30 minutes at 4°C) and supernatants collected. For Western blot analysis, 25 µL total protein sample per lysate was added to Laemmli buffer (Bio-Rad Laboratories, Hercules, CA, USA) and boiled at 95°C. Samples were separated by electrophoresis on 4% to 20% criterion TGX Precast Gels (Bio-Rad Laboratories), and proteins were transferred to a P 0.45 polyvinylidene fluoride (PVDF) membrane (Amersham Biosciences, Slough, Buckinghamshire, UK) using an Amersham Biosciences semi-dry transfer apparatus. PVDF membranes were blocked for 2 hours at room temperature with 5% nonfat milk in TBST buffer. After blocking, membranes were incubated with mouse primary antibody against C3d (clone 11), described in Thurman et al.⁴³ Secondary antibody (Santa Cruz Biotechnologies, Dallas, TX, USA) was diluted (1:3000) in 5% nonfat milk and membranes exposed overnight. The next day, membranes were washed three times with TBST buffer and then incubated with anti-mouse IgG horseradish peroxidase–linked secondary antibody (Cell Signaling Technology, Danvers, MA, USA) for 1.15 hours. Membranes were washed again three times with TBST buffer followed by a 5-minute incubation with Clarity Western ECL Substrate Reagent (ECL; Bio-Rad Laboratories) for chemiluminescent detection. Blots were stripped with stripping buffer (Thermo Fisher Scientific) for 20 minutes and reprobed with a rabbit monoclonal antibody against (human GAPDH; Cell Signaling Technology) for normalization purposes. A second chemiluminescent detection was performed, and all protein bands were captured by film. Protein bands were scanned, and their densities were quantified using ImageJ software. 

Statistics

Data are presented as mean ± SEM. Statistical analysis for single comparisons was performed using unpaired t-tests with mean value differences considered significant at *P ≤ 0.05, **P ≤ 0.01, and ***P ≤ 0.001. Repeated-measures analysis of variance was used for a- and b-wave analysis with Bonferroni corrections for multiple comparisons across all intensities. Analysis was via GraphPad Software (San Diego, CA, USA). 

Results

Proof of Concept That the C3 Promoter Is Induced by Oxidative and Complement Stress

The complement C3 promoter spans more than 1 kb, and several regulatory elements have been identified. Specifically, the promoter fragment (−1005 to +251), which has previously been shown to drive gene expression in malignant ovarian epithelial cells,²⁹ was obtained for these experiments. In an in vitro proof-of-concept study, we explored whether pC3 can be regulated in an oxidative/complement stress-inducible manner in ARPE-19 cells. ARPE-19 cells were cotransfected with two plasmids, one confirming transfection (pCMV driving GFP; data not shown) and one testing for C3 promoter activity (pC3 driving mCherry). Forty-eight hours after cells were transfected, cells were treated with stressors known to trigger oxidative stress and complement C3 activation in these cells.⁴⁴ H₂O₂ treatment, with or without the presence of NHS, significantly induced mCherry expression when compared to pC3-mCherry untreated controls (Figs. 1A, 1B, 1C). One-hour pretreatment with the antioxidant NAC or the alternative pathway of complement inhibitor TT30 (humanized CR2-fH)²³ significantly prevented the induction of pC3-driven mCherry fluorescence (Figs. 1D, 1E, respectively). Fluorescence intensity of images from three independent experiments was quantified (Fig. 1F), showing that stress conditions produced a significant increase in mCherry compared to control. This effect was reduced to control levels by pretreatment with NAC or TT30, respectively (Fig. 1F). These experiments provide evidence that the pC3 fragment is sufficient to drive gene expression in RPE cells and that the promoter activity can be regulated in a stress-dependent manner. 

Figure 1.

 

Proof-of-concept study, confirming C3 promoter activity in ARPE-19 cells under stress and protection conditions. The pTR plasmid described in Supplementary Figure S1 with the C3 promoter driving the mCherry reporter construct was used to transfect ARPE-19 cells, which after 48 hours were exposed to different treatments. Untreated (A) cells expressed baseline levels of mCherry, whereas 0.1 mM H₂O₂ (B) or 0.1 mM H₂O₂ plus 10% normal human serum (NHS) (C) significantly induced the C3 promoter activity, as shown by the increase in red fluorescence. On the other hand, 1-hour pretreatment of antioxidant N-acetyl cysteine³⁰ (D) or the complement alternative pathway inhibitor TT30 (humanized CR2-fH) (E) prevented the induction of C3 promoter activity. (F) mCherry fluorescence was analyzed for the five experimental conditions, using ImageJ. Scale bar: 100 µm. Data shown are average values (± SEM), with n = 3 biological and 3 technical replicates per condition, *P ≤ 0.05.

Evaluation of Retinal Thickness and Function 1 Month After Subretinal Injection of AAV5-pC3-CR2-fH

Experiments to analyze the effects of subretinal injections of AAV5-pC3-CR2-fH vector (see Supplementary Fig. S1 for map of the vector) on structure and function of the retina were performed 1 month after subretinal detachment, a time point previously used by us and others.^25,27,45,46 One month postinjection, we observed that the retina was reattached to the RPE layer as shown in representative fundus and OCT images of both AAV5-pC3-mCherry and AAV5-pC3-CR2-fH mouse eyes (Fig. 2A). OCT B-scan imaging, using InVivoVue Diver software for the analysis of retinal layer thickness, was used to compare retinal nerve fiber layer, inner plexiform layer, inner nuclear layer, outer plexiform layer, outer nuclear layer,⁴⁷ inner segments, outer segments, RPE, and total retinal thickness between control AAV5-pC3-mCherry and AAV5-pC3-CR2-fH–injected eyes across the central retina. Results indicate that there was no difference in any of the layers when comparing eyes with expression of CR2-fH to the control vector (Figs. 2B, 2C). Likewise, if a single location (350 µm nasal of the optic nerve) was manually compared, there was no difference between the two groups (P > 0.05; data not shown). 

Figure 2.

 

Optical coherence tomography analysis 1 month after AAV5 treatment confirms lack of cytotoxicity. (A) Optical coherence tomography B-scan imaging 1 month after the subretinal injection of AAV5-pC3-mCherry or AAV5-pC3-CR2-fH reveals complete reattachment of the retina to the RPE layer at the posterior pole and appearance of the retina that did not vary between the two treatments. Scale bar: 200 µm. In the presence of CR2-fH expression, all retinal layer thicknesses are maintained (B), including total retinal thickness (C), when compared to mCherry controls. INL, inner nuclear layer; IPL, inner plexiform layer; IS, inner segments; ONL, outer nuclear layer; OPL, outer plexiform layer; OS, outer segments; RNFL, retinal nerve fiber layer. Unpaired t-test for single comparisons was performed using GraphPad. Data are expressed as mean ± SEM, representing n = 10 mouse eyes per condition.

Visual electrophysiology tests 1 month postinjection, using ffERG, were used to measure the electrical activity of the retina and RPE tissues in response to a light stimulus. Electroretinography (ERG) a- and b-wave amplitudes at increasing light flash intensities from −30 to 0 dB were found not to differ between animals treated with C3-promoter driven expression of CR2-fH compared to mCherry controls (P = 0.54 and P = 0.13, respectively) (Fig. 3A). Likewise, c-wave amplitudes measured in response to a single maximal intensity light flash did not differ (P = 0.35) between cohorts (Fig. 3B) at the same dose. Taken together, our results conclude that retinal structure and function were not affected by the expression of CR2-fH when compared to mCherry control. 

Figure 3.

 

Functional assessment by electroretinography 1 month after AAV5 treatment confirms lack of cytotoxicity. (A) Retinal a- and b-waves (photoreceptors and bipolar cells) in response to light intensities from –30 to 0 dB white light were evaluated by ffERG 1 month after AAV5-pC3-mCherry or AAV5-pC3-CR2-fH injection. (B) C-wave amplitudes (RPE standing potential) measured by ffERG. Overall, retina and RPE tissue function were not affected by the presence of the AP inhibitor when compared to mCherry. n = 7 animals per condition. The a- and b-wave data were analyzed using repeated-measures analysis of variance with Bonferroni corrections for multiple comparisons across all intensities. Data are expressed as mean ± SEM.

C3-Promoter Drives CR2-fH Expression and Reduces Complement Activation In Vivo

One month after AAV injections, eyes were harvested and CR2-fH expression and complement inhibition were analyzed in the RPE/choroid complex of CNV mice injected with AAV5-pC3-CR2-fH when compared to AAV5-pC3-mCherry or uninjected, no CNV age-matched control mice. 

Quantitative reverse transcriptase polymerase chain reaction (RT-PCR) analysis, using primers spanning the CR2-fH linker, confirmed the presence of CR2-fH mRNA in the pC3-CR2-fH–injected eyes (Supplementary Fig. S2A), and dot blot analysis documented the presence of CR2-fH protein (Supplementary Fig. S2B). 

Complement activation was analyzed using Western blot analysis of RPE/choroid complexes, probing for the presence of the long-lasting opsonin C3d. We have previously reported a reduction in C3d in AAV5-VMD2-CR2-fH–treated mice when compared to AAV5-VMD2-mCherry.³³ The same antibody⁴³ was used to probe for C3d-specific bands in Western blots comparing naive eyes to CNV mice injected with AAV5-pC3-mCherry or AAV5-pC3-CR2-fH (Fig. 4A). C3d fold change levels were low in samples from control animals but significantly elevated in response to CNV lesions in AAV5-pC3-mCherry–treated animals (P = 0.03). In contrast, AAV5-pC3-CR2-fH vector injections significantly reduced complement activation or C3d deposition (P = 0.03) when compared to AAV5-pC3-mCherry. Taken together, pC3 mediated CR2-fH mRNA and protein expression and led to a reduction in complement activation products C3d/C3dg in the CNV mouse model. 

Figure 4.

 

AAV5-pC3-CR2-fH subretinal injections mitigate complement activation compared to controls. (A) Western blots of RPE/choroid homogenates probed for C3dg/C3d (37–34 kDa) followed by GAPDH (36 kDa) revealed a reduction in the amount of C3dg/C3d deposited in mouse CNV-lesioned eyes after AAV5-pC3-CR2-fH injection compared to no CNV lesions or CNV eyes treated with AAV5-pC3-mCherry. (B) Quantification of the amount of C3dg/C3d normalized to GAPDH and plotted as fold change over control. Data are expressed as mean ± SEM; n = 4–5 eyes per condition, *P ≤ 0.05.

C3-Promoter Driven CR2-fH Attenuates CNV Lesion Size and Fluid Leakage

Mice were injected subretinally with either the pC3-mCherry control vector or the pC3-CR2-fH experimental vector. Bleb formation or retinal detachment was confirmed by fundus imaging on the day of the injection, typically detaching ∼60% to 90% of the retina (data not shown). Fundus imaging confirmed that retinas reattach 1 month after subretinal injection, documenting little scar formation (Fig. 5A). En face OCT imaging of the CNV lesions (Fig. 5A) revealed that AAV5-pC3-CR2-fH at 2.41 × 10¹¹ vg/mL were found to be significantly efficacious at reducing CNV lesion sizes (P = 0.008) compared to AAV5-pC3-mCherry controls (Fig. 5B). The percent reduction in lesion size area (∼30%) is similar to that of our previous experiments using AAV5-VMD2-CR2-fH²⁵ or systemic injections of CR2-fH protein.²³ The lesions contain choroidal fibrovascular tissue (Fig. 5C), which we have shown previously to be immunopositive for the endothelial cell marker Isolectin B4.¹⁴ Likewise, signs of subretinal fluid accumulation were reduced in AAV5-VMD2-CR2-fH–treated animals. Subretinal fluid was present in ∼75% of all lesions, with many of the fluid domes in the large category in animals receiving AAV5-VMD2-mCherry. In contrast, in animals that received AAV5-VMD2-CR2-fH, fluid accumulations could only be identified in ∼25% of the CNV lesions, with most of the fluid domes in the small category (Figs. 5C, 5D). 

Figure 5.

 

Choroidal neovascularization lesions are mitigated by CR2-fH expression. Mice subretinally injected with 1 µL AAV5-pC3-mCherry or AAV5-pC3-CR2-fH at 2.41 × 10¹¹ (vg) were exposed to laser CNV 1 month after injection. (A) Fundus images document retinal reattachment (left); optical coherence tomography en face imaging was used to determine CNV lesion sizes (right). (B) Lesion sizes induced by laser photocoagulation were quantified using ImageJ and found to be significantly smaller in AAV5-pC3-CR2-fH–treated animals compared to AAV5-pC3-mCherry controls. (C) Representative B-scan images showing subretinal fluid accumulation (white arrows) in AAV5-pC3-mCherry and AAV5-pC3-CR2-fH–treated mice. (D) Quantification of lesion severity scores. Lesions with or without subretinal fluid area received a severity score ranging from absent to small, medium, and large. AAV5-pC3-CR2-fH–treated mice exhibited fewer fluid domes that were smaller in size when compared to mCherry controls. Scale bars: 200 µm. Data in (B) are expressed as mean ± SEM; n = 7 animals per condition, **P ≤ 0.01, in (D) as percentage of the overall number of lesions, ***P ≤ 0.001.

Discussion

The purpose of this study was to evaluate the utilization of an AAV-mediated delivery of CR2-fH in a complement-regulatable construct driven by the C3 promoter. The results of the study are as follows: (1) we have provided proof of concept that pC3 driving the reporter mCherry is induced by cellular stressors, and pretreatment with a complement AP inhibitor TT30 (CR2-fH)²³ or antioxidant NAC prevented its induction. (2) Fundus and SD-OCT imaging 1 month after subretinal detachment confirmed reattachment of the retinas. We report no changes to individual retinal layers or total retinal thickness by AAV5-pC3-CR2-fH compared to AAV5-pC3-mCherry–injected eyes. (3) Additionally, ERG amplitudes (a-, b-, and c-waves) were not differentially affected by the treatment. (4) PCR and dot blot analysis confirmed expression of the CR2-fH construct at the mRNA and protein levels, respectively, in AAV5-pC3-CR2-fH–treated eyes. (5) Significant complement inhibition in the RPE/choroid of CNV mice was by AAV5-pC3-CR2-fH as measured by C3d/C3dg Western blot analysis. (6) Last, we report a significant reduction in CNV lesion sizes and subretinal fluid accumulation with AAV5-pC3-CR2-fH treatment. Overall, these results show that regulating the AP in a complement-dependent manner lessens complement activation and CNV in a mouse model. We demonstrate that AAV5-pC3-CR2-fH is not only efficacious but also safe based on retinal structure and function analysis. 

A C3-Regulatable Promoter

Previously, we have shown that CR2-fH, when expressed in the RPE under a VMD2 RPE-specific promoter and delivered via AAV5, reduces CNV and smoke-induced ocular pathology. However, as we indicated in the original manuscript, CR2-fH expression needs to be titrated to identify a dose that is not damaging to the retina.²⁵ This observation is not unexpected, as both knockouts of complement activators or inhibitors lead to retinal damage.^48,49 To circumvent some of the dosing issues, we investigate the use of the complement C3 promoter to drive expression of CR2-fH in a complement-dependent manner. We first provided evidence that the pC3 is activated in RPE cells by cellular stressors. Trakkides et al.³¹ have shown that ARPE-19 cells significantly increased C3 mRNA and protein production in the presence of H₂O₂, and we have shown that oxidative stress induced by cigarette smoke exposure in ARPE-19 cell monolayers led to C3 expression and secretion, which was prevented by complement inhibition or the addition of an antioxidant.²⁰ We used the same paradigm to demonstrate that pC3-driven mCherry expression in ARPE-19 cells can be activated by oxidative stress or complement activation and prevented by pretreatment with the antioxidant NAC or the AP inhibitor TT30 (CR2-fH).²³ Based on these results, an AAV5 viral vector was generated driving CR2-fH under the pC3. A safe concentration (1 µL of 2.41 × 10¹¹ vg/mL) of AAV5-pC3-CR2-fH was confirmed based on the analysis of mouse retinal and RPE structure and function 1 month postinjection. AAV5-pC3-CR2-fH was found to produce CR2-fH mRNA and protein, reduce complement activation, and reduce CNV in this short-term model. It would be of great interest to perform a long-term study to confirm that the pC3 mimics complement activation as well as deactivation. Additionally, future experiments are needed that confirm that after long-term AAV5-pC3-CR2-fH exposure, the tissues maintain homeostatic levels of complement opsonins and anaphylatoxins, which are essential for normal tissue function and wound repair in CNV.¹⁵ 

Gene therapy using a regulatable promoter has been reported by Biswal et al.,⁵⁰ who used a hypoxia-regulatable cassette in mouse models of wet AMD. That study's intent was to mitigating hypoxia-induced upregulation of inflammatory cytokine products and neovascularization. Specifically, they transduced the RPE using subretinal injections with an AAV2 vector containing an RPE cell-specific promoter (RPE65) construct, hypoxia-regulated elements, and endostatin,⁵¹ an antiangiogenic agent termed REG-RPE-ENDO. This hypoxia-inducible promoter drove the production of endostatin in a hypoxia-dependent manner. They concluded that this construct mitigated angiogenesis in mouse models of wet AMD by ∼80%.⁵⁰ Overall, pC3-CR2-fH and REG-RPE-ENDO are both approaches that regulate pathology on an as-needed basis. Both constructs appear to stabilize inflammation and immune response overactivation, as well as provide a homeostatic environment for tissue health. 

Safety Analysis: Retinal Thickness and Function After Reattachment

To confirm retinal reattachment to the posterior mouse eye, 1 month after detachment, fundus and SD-OCT imaging were performed. Here we report no changes in retinal layer thickness in the central retina when comparing AAV5-pC3-CR2-fH with AAV5-pC3-mCherry–injected eyes. Measurements were used to determine if retinal detachment, virus, or expression of CR2-fH caused loss of cells or affected layer thickness. Likewise, ERG analyses, determining photoreceptor, bipolar, and RPE cell function, demonstrated that light responses were similar under both conditions. It has been shown that changes in retinal anatomy and thickness correlate with a decrease in light sensitivity and visual acuity.^52,53 Additionally, retinal layer thickness has often been correlated with the health of mouse retinas.^41,54 Since we report preservation in retinal thickness, we suggest that the expression of CR2-fH is not toxic to the retina. However, structure–function analysis will need to be repeated to examine long-term effects of both the virus and the cargo. 

Notably, one limitation in our study, as well as others that use subretinal injections, is that a retinal detachment causes neuroinflammation that impacts neuronal survival in the retina. Numerous studies have shown that detachment of the retina from the RPE results in activation and infiltration of macrophages that change the environmental cytokine and chemokine profile, leading to photoreceptor cell death.^55,56 Likewise, the AP of complement has been shown to be upregulated by retinal detachment, contributing to photoreceptor cell damage.⁵⁷ Thus, any subretinal injections must counteract not only the proinflammatory environment caused by the disease itself but also that caused by the delivery technique. In that context, long-term models (readout at 7 months postinjection) such as the smoke-induced ocular pathology model investigated by us²² might be more predictive of vector efficacy than the short-term model used here (readout at 1.5 months postinjection). Previous work in Schnabolk et al.²⁵ has shown, using ffERG analyses, that c-wave response amplitudes after subretinal injection compared to preinjection were significantly lower but not different between CR2-fH and control mCherry cohorts.²⁵ With that being said, it has been shown that a retinal detachment of ∼60% to 90% coverage in a mouse eye, which we report here, significantly reduces ERG b-wave amplitude responses compared to noninjected mouse eyes.³² 

In contrast to the animal studies in which damage and neuroinflammation are probably caused by the large retinal detachments,⁵⁵ much smaller subretinal injections are being performed on patients with AMD in clinical trials. These smaller injections are aimed at localizing treatments to areas of the retina that require protection from the advancing geographic atrophy area⁵⁸ or in retinitis pigmentosa, to target a healthy area of the retina.⁴⁷ It is of interest to note that in patients receiving RPE65 gene replacement, despite functional improvement, cone photoreceptors still degenerate, albeit at a slower pace,⁵⁹ potentially due to long-term effects of the subretinal injection even in small spots. 

AMD Therapy Targeting Complement

Increasing evidence suggests that complement dysregulation is involved in AMD pathogenesis. Numerous different AMD therapies are in development from novel fusion proteins to antibodies and even gene therapies targeting complement proteins or activation itself. When designing a complement inhibitor, it is important to keep in mind that complement factor H (Y402H)⁵ C3 (R102G and L314P)⁶⁰ and factor B (rs641153)⁶¹ polymorphisms and the ARMS2 (A69S)⁶² variant are all associated with increased complement activation, presumably contributing to both wet and dry AMD pathology. 

The current study is part of a long-term program, characterizing the CR2-fH fusion protein and its effects in mouse and cell-based models of AMD. CR2-fH was initially injected systemically as a fusion protein,¹⁴ followed by delivery of fH fusion proteins through cell encapsulation using both intravitreal²⁴ and systemic delivery,⁶³ leading to gene therapy efforts.²⁵ Here we extended this analysis to gene therapy using a regulatable promoter, documenting safety and efficacy in the mouse laser-induced CNV model. However, it will be important to confirm our results obtained by OCT to determine lesion size and fluid accumulation with additional tools such as OCT angiography, fluoresceine angiography, or histology as well as in additional models such as the smoke-induced ocular pathology model.³³ 

Currently, numerous therapeutic approaches targeting complement dysregulation are in development and in clinical trials for AMD and GA, targeting different components of the complement cascade, ranging from complement C3 and C5 inhibitors along the common terminal pathway to MAC. C3 inhibition, targeted by Apellis, is a recently FDA-approved intravitreal injection of pegcetacoplan, a pegylated peptide C3 inhibitor, that significantly mitigated GA progression (NCT04770545). Complement inhibition by pegcetacoplan therapy targets all local complement protein as it represents the central protein for all three pathways. It is expected that C3 inhibition would result in a local tissue environment with less complement-mediated inflammation but also fewer opsonins (C3b, iC3b, C3dg, and C3d) and anaphylatoxins required for clearance of debris or the polarization of monocytes and macrophages fundamental in wound repair.⁶⁴ Similarly, C5 inhibition is being targeted by intravitreal injections of a C5 inhibitor (avacincaptad pegol [Zimura]; Iveric) that has completed phase III clinical trials and reduced the growth rate of GA by the same margin as C3 inhibition.⁶⁵ Reducing C5 cleavage would reduce C5a production, as well as MAC formation, which should reduce inflammation and cell destruction. Nevertheless, it is important to note that C5a also has an important role in signaling RPE cells to produce essential and neuroprotective VEGF_A.¹³ Finally, in both the trials for pegcetacoplan and avacincaptad pegol, an increase in conversion rate to exudative AMD and CNV formation has been reported when compared to the controls. It has been speculated that complement inhibition might alter microglial cell polarization toward a proangiogenic phenotype, as has been shown for vascular development,⁶⁶ the addition of polyethylene glycol (PEG) providing a proangiogenic moiety, as documented for animal models of CNV,⁶⁷ or anti-PEG antibodies being boosted by the presence of the pegylated drug, as has been shown for the SARS-CoV-2 vaccine,⁶⁸ potentially triggering antibody-mediated complement activation in the RPE/BrM/CC, all counteracting the protective effects of the anticomplement activity. Thus, it will be of great interest to determine whether a nonpegylated anticomplement therapeutic that is based on providing homeostatic control over complement activation such as factor I (fI) overexpression (NCT03846193) will have similar side effects. Here we showed that CR2-fH reduced not only CNV lesion size but also fluid leakage. 

The eye is an ideal organ for lasting gene therapy as it is to some degree immune privileged and contains tissues (including the RPE) that do not turn over and can therefore express genetic material long term. Many AAV gene therapy vectors target the complement cascade in AMD pathology. AAV gene therapy vectors are extensively used and currently in clinical trials to treat ocular diseases,²⁶ including Leber congenital amaurosis.⁶⁹ Gene therapy efforts in AMD are focused primarily on complement inhibition.^8,70 Gyroscope's investigative fI gene replacement therapy (GT005) is a subretinal injection of the AAV2 vector that contains their therapeutic cassette of chicken β-actin promoter driving factor I (NCT03846193). The fI variants have been linked to the development of dry AMD and retinal photoreceptor loss due to their inability to inactivate or cleave C3b to iC3b.⁷¹ Factor I and cofactor fH together are responsible for inactivating complement C3 cleavage product C3b, which is a component of C3 convertase in the AP that feeds the amplification loop.⁸ Notably, GT005 is in phase I/II clinical trials designed to increase the bioavailability of factor I in patient's eyes and reduce the overall area of GA.⁷² Increasing fI, like CR2-fH, would reduce overall complement activation initiated by the alternative pathway and the AP amplification loop overall, reducing complement activation in a proportional manner, rather than inhibiting at a single point. In contrast, Hemera Biosciences (Waltham, MA, USA) has developed an intravitreal injection of an AAV vector (AAVCAGsCD59) that expresses a soluble MAC (sCD59) inhibitor.⁷³ MAC is the terminal pathway that forms cytolytic pores in the membranes of cells. This complement inhibitory approach is being used in phase II clinical trials (NCT03585556) on patients with advanced AMD. Soluble CD59 has been shown to significantly attenuate CNV in a mouse model of wet AMD.⁴⁶ However, targeting only the lytic segment of complement does not affect upstream AP activation and the AP amplification loop, which will continue to lead to increased MAC deposition and anaphylatoxin production, perpetuating the immune response. Thus, it will be of great interest to determine the long-term consequences of affecting the whole complement cascade, by modulating the overall activity (fI gene replacement and CR2-fH), when compared to inhibition at particular breakpoints (C3, C5, and MAC). Regardless, advancements in efficacious ocular gene therapy delivery and accurate complement targeting show promising potential for the future of complement-dependent AMD treatment. 

Currently, multiple invasive intraocular injections of anti-VEGF therapeutics are the standard of care for patients with wet AMD. However, efforts are under way to develop alternative approaches to monthly injections since they subject patients to complications like potential bleeding, cataracts, infection, and inflammation.⁷⁴ In our previous studies, we have shown that complement inhibition reduces CNV lesion sizes. Specifically, we have shown that CR2-fH localizes to CNV lesions and reduces complement and VEGF expression in CNV mice.¹⁴ Likewise, in RPE monolayers, CR2-fH reduced complement-mediated VEGF secretion and prevented concomitant VEGF-induced loss of barrier function.⁴⁴ Here, we confirmed that CNV lesion sizes and subretinal fluid accumulation were both reduced by AAV5-pC3-CR2-fH. This suggests that complement therapeutics alone or used in conjunction with anti-VEGF therapeutics may reduce features of wet AMD. 

Numerous attempts are being made to develop gene therapy for wet AMD. Specifically, Adverum (Redwood City, CA, USA) has developed a gene therapy vector to express Eylea, which is currently in phase I clinical trials with no posted results (NCT04418427). Similar to our approach, and acknowledging that constant VEGF inhibition has been shown to lead to macular atrophy,⁷⁵ Reid et al.⁷⁶ are using mouse models of wet AMD to deliver a recombinant adeno-associated viral expression cassette containing a tetracycline-regulatable riboswitch for modulating Eylea expression in vivo. This approach significantly reduced CNV lesion sizes.⁷⁶ Overall, these studies suggest that the future of wet AMD treatment might also be a onetime injection of a long-term gene therapy that potentially targets both complement inhibition and anti-VEGF therapeutics. 

In summary, our experiments report the proof of concept that the C3 promoter can be regulated in a complement-dependent manner in vitro. In vivo AAV5-pC3-CR2-fH significantly reduces the AP of complement in mouse eyes as well as CNV lesion size and subretinal fluid accumulation. The efficacy of a complement-dependent regulator that restores ideal homeostatic levels of complement activation provides an avenue for treatment of other complement-associated diseases. 

Acknowledgments

The authors thank the MUSC Molecular Analytics core facility for providing access to quantitative RT-PCR support and Dr. Gloriane Schnabolk (MUSC) for critical review. 

Supported in part by the Department of Veterans Affairs awards RX000444, BX003050, and IK6BX004858 (to BR); the National Institutes of Health award EY019320 (to BR); the Smart State Endowment from the State of South Carolina (to BR); the Interdisciplinary Research Training in Otolaryngology and Communication Sciences T32 Support DC014435 (to J. R. Dubno); the NIH National Institute of Neurological Disorders and Stroke Diversity Specialized Predoctoral to Postdoctoral Advancement in Neuroscience Grant F99NS124188 (to NBP); and the Southern Regional Education Board Scholars Award (to NBP). 

Disclosure: N.B. Parsons, None; B. Annamalai, None; B. Rohrer, utilization of CR2-fH in complement-dependent diseases (P)