The clinical and ERG data of one patient and one healthy unaffected daughter of the patient is included (Table 1). Complete ophthalmological examination by fundus photography and SD-OCT revealed features of RP (Hirji 2023) including bone spicule-like pigmentation changes, optic disc pallor and attenuation of retinal blood vessels (Figure 1A, Supplementary Figure 1), with outer retinal atrophy due to the loss of the photoreceptor layers, sparing the central foveal region (Figure 1B). While the central foveal region was relatively spared in this patient, the macular cones that remained were observed to be damaged by chronic edema, and photoreceptor and RPE atrophy had progressed into the macula. Genetic testing of the proband by next-generation sequencing (NGS) showed a complex duplication-rearrangement of chromosome 3q22, which encompassed the entire RHO coding sequence, 5’ and 3’ regulatory regions and flanking genes. The rearrangement consisted of a 48 kb triplicated region embedded within a 188 kb duplication, resulting in three apparently intact RHO genes on one chromosome and a fourth, unaltered RHO gene on the homologous chromosome. Within the duplications, RHO copies were flanked by H1FOO, IFT122, EFCAB12 and MBD4 while the inverted triplication of RHO was flanked by partially triplicated IFT122 and H1FOO (Figure 1C). Thus, the NGS data shows a duplication-inverted triplication-duplication event, in which the triplicated segment is inverted and located between correctly oriented genomic segments (Figure 1C). No other additional causative variants of inherited retinal degeneration candidate genes were identified. Whole genome sequencing supported the rearrangement and extra copies of RHO which were identified by NGS. The RHO copy number variants were not detected in the unaffected daughter of the patient. Clinically collected four-generation pedigree data of the family included two male family members affected with RP, indicating male-male transmission, consistent with an autosomal dominant inheritance. The proband’s affected father was not examined as he was deceased but had been diagnosed clinically with RP. The clinical findings were consistent with the hypothesis that CNV in the wild-type RHO causes adRP.

To assess the effects of RHO-CNV in human retinal organoids, we initially reprogrammed the peripheral blood mononuclear cells (PBMNCs) from one patient with four copies of RHO (RM) as well as the corresponding familial control (RC) into iPSCs (Supplementary Figure 2A-B). The iPSC lines had the colony morphology of tightly packed cells with a high nucleus-to-cytoplasm ratio, a well-defined border typical of stem cells, and expression of pluripotent markers (Supplementary Figure 2C-D). We then differentiated the patient and control iPSC lines into retinal organoids by following our previously published protocols(Bachu et al. 2022; Arthur et al. 2022), as per the differentiation timeline depicted in Figure 2A. Retinal organoids from the RHO-CNV patient and the control displayed a well-defined neuroepithelial lamination, indicating the arrangement of photoreceptors on the apical surface of the organoids and a dark central basal region consisting of inner retinal cells by phase contrast microscopy (Supplementary Figure 2E). Notably, there were no clear visible anatomical changes in apical-basal retinal cell type distribution during the early differentiation timeframe, when the cone and rod photoreceptors are usually born in 45-50-days-old and 90-120-days-old human retinal organoids, respectively (data not shown). Over the prolonged differentiation culture timeframe (>200 days), the control retinal organoids displayed long hair-like protrusions which were presumptive inner and outer segments at the apical side of the retinal organoids, a critical event indicating the start of photoreceptor maturation. Conversely, the patient retinal organoids showed short initial hair-like protrusions that did not elongate at the extended culture time of 260 and 300 days in culture by light microscopy (Supplementary Figure 2E, Figure 2B). To validate these observations, we examined the ultrastructure of photoreceptors in the RHO-CNV and control organoids using transmission electron microscopy (Figure 2C, Supplementary Figure 3E). Upon assessing the 300-days-old organoids, we observed that while the patient organoids developed connecting cilium and inner segments similar to control organoids, they failed to extend outer segments. This data was consistent across multiple rounds of differentiation from three independent clones of patient iPSCs. Taken together, these morphological changes suggest that iPSC-retinal organoids of RHO-CNV patient demonstrated the survival of photoreceptors with OS dysmorphogenesis over the time course of 300-days.

Control and patient retinal organoids were analyzed at two developmental stages: at rod photoreceptor birth (D120), and maturation (D300). For each sample, qRT-PCR was carried out by utilizing primers designed to nine key genes that regulate the development and maturation of rod photoreceptors. mRNA levels were analyzed for the expression of genes specific to early pan photoreceptors (OTX2, CRX, RCVRN), early rod photoreceptors (NRL, NR2E3), rod-specific phototransduction (PDE6B, SAG, RHO) and ciliary (IFT122) genes (Supplementary Table 4). To equilibrate the data to equivalent number of photoreceptors in organoids, we normalized the data to CRX expression, a transcription factor highly enriched in photoreceptors in the retina (Yamamoto et al. 2020). The expression of all the target genes were detected at each time point (D120 and D300) in the retinal organoids (Figure 3A). Pan-photoreceptor and early rod marker genes showed similar expression levels with no noticeable variations in RC and RM organoids. In contrast, RHO and SAG were expressed at a higher level in the patient compared to control organoids. There was a significant eight fold increase in the RHO levels at D120 and D300 in the patient organoids compared to controls. We also observed a small two-fold statistically significant increase in rod/visual arrestin (SAG) at D300 timepoint in the patient organoids compared to control organoids. Relatedly, a one-fold, but non-statistically significant, increases in patient organoids was observed in IFT122, a gene partially triplicated in NGS along with RHO (Figure 1C).

To advance a better molecular understanding on detailed genomic expression and gene regulatory networks, we performed bulk-RNA sequencing comparing 300-days-old retinal organoids from patient and control iPSCs (n=3 independent biological replicates). Patient retinal organoids demonstrated upregulated transcriptomic levels of RHO (∼eight-fold) and SAG (∼four-fold) compared to control organoids, comparable to the differences observed in the qRT-PCR data (Figure 3B). Additionally, we also observed increased expression in disc structure support gene including PRPH, visual cycle genes RDH8 and HCN1, and a synaptic gene, PTPRT in patient relative to control organoids (Figure 3B). Following the Gene Ontology enrichment analysis (EnrichGO) of significant differentially up-regulated genes, we confirmed increases in phototransduction cascades including the visual/light perception and membrane potential/Ca+ signaling pathways in patient organoids compared to control organoids. Additionally, increases in genes associated with the synaptic signaling pathway were also detected, suggestive of photoreceptor dysfunction (Figure 3C). Pathway enrichment analysis of significantly differentially expressed genes for cellular components category pointed to perturbation in pathways associated with defects in glycosylation especially N-linked glycosylation as well as ER and Golgi transport (Figure 3D, Table 2), highlighting potential pathophysiology that may drive RHO-CNV associated RP. Although glycosylation is not required for the rhodopsin biosynthesis, N-linked glycosylation (at N2 and N15), a post-translational modification, is a necessary step for interacting with chaperones during ER transport. This is also an essential step towards incorporation of the heptahelical G-protein coupled receptor rhodopsin in the rod outer segments. Thus, RNAseq data suggests that the defects in rhodopsin glycosylation decreased the ability of rhodopsin to exit ER, and lead to an adRP phenotype (Tam and Moritz 2009; C.LH. Sung and Tai 1999).

Immunofluorescence staining on the cross sections of control and patient retinal organoids was examined for spatial location of the photoreceptors at three time points (D120, D200, D300). Pan-photoreceptor precursor and progenitor proteins (OTX2, CRX), and rod specific proteins (NR2E3) were expressed in the apical layer of the organoids in a similar pattern both in the control and patient organoids with equivalent photoreceptors and outer nuclear layer thickness (Supplementary Figure 3A-C). These results corroborated our gene expression and transcriptomics data (Figure 3A-B) indicating no defects in rod biogenesis within the patient retinal organoids. As the organoids began to mature, the distribution of RHO protein was observed in the outer segments of control organoids starting at D200, earliest timepoint for detection in human retinal organoids using our differentiation protocol. Compared to the control, the patient organoids had mis-localized RHO protein accumulating in the photoreceptor cell soma, at all analyzed time-points (D200, and >D250) (Figure 4A-B). The patient organoids rarely showed RHO in IS/OS which were notably consistent with our results discerned by light microcopy (Supplementary Figure 2E, Figure 2B). Co-staining with NRL and CRX confirmed mislocalized RHO expression in the rod photoreceptor soma within patient organoids. We also evaluated the expression of rod-specific phototransduction protein, SAG, which was increased in the outer nuclear layer of patient retinal organoids (Figure 4C). We did not observe any differences in the cone markers, ARR3, BCO or pan phototransduction marker, RCVRN when comparing >D250 patient to control organoids (Supplementary Figure 3D). There was also no difference in apoptosis between the patient and control organoids (Supplementary Figure 3F).

To investigate protein expression levels in RHO-CNV organoids, we quantitated the level of RHO and SAG by western blot by comparing to control organoids. Patient retinal organoid homogenates displayed a significant 16-fold and 9-fold higher fractions of ∼40 kDa monomer and ∼80 kDa dimer rhodopsin content respectively in patient organoids relative to controls despite loading equal amounts of protein lysates by western blot (Figure 4D-D’’). A significant 1.5-fold increase in ∼48 kDa SAG, a rhodopsin interacting protein was also observed (Figure 4E-E’). Additionally, we analyzed the ER stress-UPR pathway in the patient organoids and did not detect any differences compared to control (data not shown). These findings suggest that RHO-CNV organoids actively translated the excessive rhodopsin protein which is not getting transported to the outer segments.

Since, the excess RHO protein production due to the extra RHO copies is likely not being processed appropriately, we aimed to test potential therapies that may reduce RHO protein levels in rods. We targeted orphan nuclear receptor, NR2E3, using a small-molecule, PR3, to assess its effect on RHO regulation and expression in the human patient organoid model (Figure 5A). PR3has been previously described to regulate the rod gene expression in Rho^P23H mice following a screen for molecules targeting NR2E3, suggesting its potential for the use in the treatment of RP (Nakamura et al. 2017). We treated long-term cultured 300-days-old, RHO-CNV patient retinal organoids with varying concentrations of PR3 (0.1, 0.25 and 0.5 µM) for one week and assessed the effects on RHO mRNA expression and protein localization. Immunofluorescence staining of PR3-treated organoids displayed a partial rescue of RHO localization (Figure 5B). 0.5 µM treatment lead to the strongest reduction in RHO with very few rods still expressing the protein. 0.25 µM treatment lead of best outer segment region localization of RHO. None of the organoids showed any evidence of toxicity (by TUNEL, Supplementary Figure 4C) post-treatment. Following qRT-PCR analysis of PR3 treated organoids compared to vehicle treated ones, we observed a four to thirty fold decrease in RHO expression in a dose-dependent manner, along with smaller decreases in other rod-specific genes including NR2E3, GNAT1 and PDE6B (Figure 5C). We did not see any significant effects of PR3 on blue-, green- and red-cone opsin genes or protein expression for these protein or cone arrestin (Supplementary Figure 4A, B). Upon comparison of PR3-treated patient organoids with control (RC) organoids. RHO expression levels in the patient organoids treated with 0.1 and 0.25 µM PR3 was not significantly different from control organoids. However, 0.5 µM PR3 resulted in significantly decreased RHO expression, much lower than the RHO levels observed in control organoids (Figure 5D).

We further carried out bulk RNA-sequencing analysis to comprehensively characterize three different groups of organoids, 0.25 µM PR3-treated and vehicle-treated patient organoids and control (RC) organoids from three independent differentiation experiments. Consistent with the qRT-PCR gene expression analysis, the results showed a significant downregulation in RHO and other rod phototransduction genes including SAG and GNAT1 between the PR3 and vehicle treated patient organoids (Figure 6A). Additionally, we confirmed that PR3 did not have any adverse effects on cone opsin transcripts in patient organoids. Principal component analysis (PCA) and normalized read counts analysis of sequenced data for rod (RHO, SAG, GNAT1, NR2E3) and other genes demonstrated that the PR3-treated organoids were more alike to control organoids than the vehicle treated patient organoids (Figure 6B-C, Supplementary Figure 5B). Upon KEGG analysis of differentially expressed genes associated with the rod phototransduction pathway between the three groups of organoids, we observed that several rod pathway components which were upregulated in patient organoids compared to control organoids were potentially salvaged following PR3 treatment (Figure 6D). EnrichGO analysis of significantly downregulated genes in visual perception/phototransduction pathways confirmed the specific activity of PR3 in retina (Figure 6E). While analysis for significantly upregulated genes showed changes in cilium organization, axoneme assembly, ER to Golgi transport and glycosylation, all of which are critically required for outer segment formation and protein trafficking (Figure 6F, Supplementary Figure 5A-B) suggestive of rod photoreceptor maturation recovery. Thus, the data presented strongly suggests that PR3 could potentially rescue rod photoreceptor homeostasis in RHO-CNV patients.

Both the proband (Male patient, 60-70 year-old) and his unaffected daughter (30-35-year-old) provided written informed consent, which was approved by the UCSF Institutional Review Board (IRB # 18-26409) and adhered to the tenets set forth in the Declaration of Helsinki. The participants gave consent to participate in the current study and have the results of this research work published. The proband (unidentified Lab ID # RM) and his asymptomatic daughter (unidentified Lab ID # RC) were examined at the University of California San Francisco (UCSF, CA, USA) including the pedigree data collection, clinical examination, and genetic diagnosis. Retinal examination included visual acuity testing, fundus photography, spectral domain optical coherence tomography (SD-OCT), full-field electroretinogram testing (ERG). For genetic diagnosis, peripheral blood (PB) samples collected from the proband and the unaffected daughter of the proband were screened by targeted Next Generation Sequencing (NGS) at Blueprint Genetic, a College of American Pathologists–and Clinical Laboratory Improvement Amendments–certified laboratory. The details of the methodology are available in our previous publication(Tuupanen et al. 2022). In brief, the sequencing was conducted using a retinal dystrophy NGS panel consisting of 266 genes, which was derived from an in-house tailored whole exome sequencing assay. The sequence reads were mapped to the human reference genome (GRCh37/hg19). CNV analysis was conducted using CNVkit (Talevich et al. 2016). Following a manual review of the identified RHO-CNV, discordant read-pairs suggesting a duplication-inverted triplication-duplication event was detected(Carvalho et al. 2011).

Peripheral blood samples collected from the human subjects (unidentified Lab ID # RC and RM) were reprogrammed into iPSCs as per the schematic outlined in Supplementary Figure 2A. Blood samples were processed by density-gradient centrifugation using Ficoll-paque™ PLUS (17-1440-02, GE Healthcare Biosciences, Sweden) to isolate the peripheral blood mononuclear cells (PBMNCs) (Supplementary Figure 2B). A small fraction of ∼1-2 x 10⁶ PBMNCs was cultured in 1-well of a 12-well suspension plate with peripheral blood mononuclear cells expansion medium (Supplementary Table 1) by changing half media every day. Date of seeding the PBMNCs in culture was designated as Day (D) −4. Four days later, ∼0.2-0.4 x 10⁶ PBMNCs were reprogrammed using CytoTune™-iPS 2.0 Sendai Reprogramming Kit (A16517, Thermo Fisher Scientific, USA), at 2.5:2.5:1.5 (KOS:c-myc: Klf4) multiplicity of infection. 24h post-transfection on D1, a full media change was done. Two days post transfection on D3, cells were plated onto 1-2 wells of a Matrigel (354234, Corning, USA, diluted as 1 µg/mL in DMEM/F12)-coated 6-well plate with reprogramming medium (Supplementary Table 1). Half media change was done every other day from D3 until D6. For all the suspension cultures, medium change was done by centrifugation at 200x g for 10 minutes and resuspending the cell pellet with the respective medium and culturing back into the plates. On D8, cells were transitioned to iPSC medium (Supplementary Table 1) with further media changes done on every other day. About 14-21 days post transfection, colonies with typical characteristics of iPSC morphology were manually picked. Pure, compact, and tightly packed iPSC clones were lifted using dissociating solution (Supplementary Table 1) for further expansion, characterization (Supplementary Figure 2C-G), and retinal organoid differentiation.

Retinal organoids were differentiated (three to six independent differentiation from each clones) using three different clones from each iPSC lines via the three step embryoid body (EB) approach following our previously published protocols and as per the schematic depicted in Figure 3A(Bachu et al. 2022; Arthur et al. 2022). Briefly, iPSC colonies were lifted and cultured in 6-well suspension plate. Small iPSC colonies self-aggregated as EBs within 24h were gradually transitioned to complete Neural Induction Medium (NIM) (Supplementary Table 1) from D0 to D3. On D6, EBs were treated with 1.5 nM BMP4 (120-05ET, PeproTech, USA). By D7, the EBs were transferred onto a Matrigel-coated plate to facilitate the adherence of EBs to the plate with NIM and 1.5 nM BMP4. Brief exposure of BMP4 in the differentiation process included the treatment of EBs with 1.5 nM (from D6-D8) to 0.75 nM (D9-D11) and 0.375 nM (D12-14). On D15, BMP4 was completely removed and EBs were fed with just NIM. From D16 through D30, the adherent EBs were fed Retinal Differentiation Medium (RDM) (Supplementary Table 1) at the intervals of every 2 days. Regions of the plate displaying clear, shiny borders indicative of retina-like morphology were manually lifted using a sterile P100 pipette tip. Lifted neural retina was transferred to a 6-well suspension plates and allowed to self-acquire the 3D organoid configuration within the next 2 days. From D31, the developing retinal organoids were fed 3D-RDM medium (Supplementary Table 1) along with 1 µM All-trans retinoic acid (R2625, Millipore Sigma) every 3days until D120. From D120 onwards, organoids were fed with just 3D-RDM medium without All-trans retinoic acid every 3 days until the completion of the study. Organoids were periodically assessed for morphological characteristics using phase contrast microscopy (Olympus IX70) at regular intervals. Various time-points of rod photoreceptor differentiation and maturation in the retinal organoids were utilized for phenotypic characterization and drug assessments (Supplementary Figure 3B).

Retinal organoids and confluent colonies of iPSCs were fixed in 4% paraformaldehyde (PFA) (157-8, Electron Microscopy Sciences, USA) in 1X PBS for 20 minutes. Organoids were cryopreserved in 15% through 30% sucrose (made in 1X PBS), and frozen in 2:1 mixture (20% sucrose: OCT, Sakura, USA). 10 µm sections of retinal organoids were collected on Super frost® Plus microscopic slides (12-550-15, Fisher Scientific, USA) using Leica CM3050 S cryostat. Immunohistochemistry was done as described previously. Organoid sections were pap-pen and PFA-fixed iPSC clones were permeabilized at room temperature for 15 mins with 0.1% Triton® X-100 (0694-1L, VWR Life Science, USA) in 10% Normal Donkey Serum (NDS, S30-100ML, EMD Millipore Corp, USA; made in 1X PBS). Following to that, sections were incubated for 1 hour in 10% NDS. Primary antibodies (Supplementary Table 3A) diluted in 10% NDS were added to the slides and incubated overnight (12-18 h) at 4°C. The slides were washed thrice in 1X PBS at 5 mins intervals and further incubated for 1 hour at room temperature with fluorescently conjugated secondary antibodies (Supplementary Table 3B) diluted at 1:250 in 10% NDS. Cell nuclei were counter stained with DAPI (1ug/mL, Roche, USA) for 10 minutes. The slides were then washed thrice in 1X PBS at 5 mins interval and cover slipped using Fluoromount G (Electron Microscopy Sciences, USA). TUNEL analysis was carried out per manufacturer’s protocol (Sigma-Aldrich). Images were acquired using ZEN software on LSM700 confocal microscope (Zeiss, Inc.) and processed by Image J (NIH, USA).

Total RNA was extracted from retinal organoids using RNeasy® Mini Kit (74104, Qiagen, USA) as per the manufacturer’s instructions. The quality and purity of the extracted RNA was assessed by NanoDrop One (Thermo Fisher Scientific, USA). cDNA was synthesized using iScript cDNA Synthesis Kit (1708891, Bio-Rad, USA). Real-time qRT-PCR was performed on CFX96 system (Bio-Rad, USA) using iTaq™ Universal SYBR® Green Supermix (64047467, Bio-Rad, USA). Primer sequences used for qRT-PCR is listed in Supplementary Table 4. The amplification reactions set were: 95°C for 30 s, 40 cycles at 95°C for 5 s, 60°C for 25 s, 95°C for 5 s and final extension at 95°C for 5 s. The Ct values of the target genes were first normalized to the endogenous control β-actin. The corrected Ct values were then utilized to compare and validate the control and patient retinal organoids. Log₂ fold change (log2FC) in gene expression of all targets were then calculated.

The quality control, RNA library preparations and sequencing reactions of the extracted RNA from retinal organoids were performed at Novogene Corporation Inc (CA, USA). FASTQ files received from Novogene following QC were quantified using Salmon package (V 1.9.0) by pseudo-aligning against homo sapiens hg19 genomic assembly in Galaxy(Afgan et al. 2016). Low-counts (<10) were filtered out and batch correction was carried out using Combat package in DEBrowser (v1.24.1) R package (Kucukural et al. 2019). Differential expression analysis was carried out using DESeq2 in DEbrowser. Genes with an adjusted p-value < 0.05 were assigned as differentially expressed. Gene ontology enrichment analysis and KEGG pathway analysis of differentially expressed genes was carried out in DEBrowser and Beavr (Perampalam and Dick 2020)packages. Differential expression analysis and pathway enrichment data have been uploaded as supplementary files (Table 2). The data is uploaded to NCBI GEO database (GSE245545).

Retinal organoids (300-days-old) were washed in 1X PBS three times at 10 mins intervals and fixed with Karnovsky’s fixative (4% paraformaldehyde/2% glutaraldehyde in 0.1 M PBS, pH 7.4) overnight at 4°C. For TEM, organoids were washed in 1X PBS and stained in 1% osmium tetroxide (19180, Electron Microscopy Sciences, USA; made in distilled water) for 1 hour at room temperature followed by staining with 2% uranyl acetate (22400, Electron Microscopy Sciences, USA; made in distilled water). After three washes in distilled water, organoids were then dehydrated in a graded series of ice-cold ethanol (50%, 70%, 95% and 100%) for 20 mins each. Organoids were incubated in propylene oxide (00235-1, Polysciences Inc, USA) twice for 5 mins, followed by a 5-mins incubation in 1:1 mix of propylene oxide: Epon 812 (13940, Electron Microscopy Sciences, USA). Organoids were allowed to infiltrate in Epon 812 overnight at room temperature. Next day, the organoids were embedded in PELCO silicone rubber molds (105, Ted Pella Inc, USA) using Epon 812, and polymerized for 48 hours at 60°C. Ultrathin (70 nm) sections were collected and imaged using a Philips Tecnai 10 electron microscope at the VA Medical Center, San Francisco, USA.

Retinal organoids were collected and washed in cold 1X PBS twice at 5 mins intervals. Protein was extracted by homogenizing the organoids using hand-held pestle (1415-5390, USA Scientific) in ice-cold RIPA buffer (See supplementary Table 2A) with 1% complete™ Protease inhibitor cocktail (11697498001, Roche Diagnostics, GmbH). Lysates were incubated at 4°C for 10 minutes and then centrifuged at 14,000 xg for 15 minutes at 4°C. Supernatant was collected and the protein concentration was measured using Pierce™ BCA Protein Assay Kit (23227, Thermo Scientific, USA) as per the manufacturer’s instructions. Equal amounts (20 µg) of protein lysates were mixed with 4x Laemmli sample buffer (161-0747, Bio-Rad, USA) and 10% dithiothreitol (DTT, R0861, Thermo Scientific, USA). Samples were resolved on a precast gel (12% Mini-Protean TGX Gels, 456-1044, Bio-Rad, USA) for 20 mins at 70 V until the loading dye entered the resolving layer, then increased to 150 V for 40-60 mins or until the run was completed (See supplementary Table 2A). After electrophoresis, the proteins were transferred onto an Immobilon^®-FL PVDF membrane (IPFL20200, Merck Millipore, USA) ice-cold transfer buffer (See supplementary Table 2A) at 100V for 90 minutes. The blots were blocked with blocking solution for 30 minutes. The blots were incubated overnight at 4°C with primary antibodies diluted in blocking solution (See supplementary Table 3A for primary antibody details). Thereafter, blots were washed thrice with TBS-T buffer (10 mins interval each) and incubated for 1 hour at room temperature with host-specific secondary antibodies diluted in blocking solution (See supplementary Table 3B for secondary antibody details). The blots were washed another three times with TBS-T buffer (10 mins interval each) and visualized using LiCor Odyssey XF scanner. For quantification of protein expression, the background was subtracted for each band and normalized to internal control band using Image J software. Statistical calculations were performed using multiple technical and five independent biological replicates.

Patient-specific RHO-CNV retinal organoids (∼300-days-old) were treated with small-drug like molecule Photoregulin3 (PR3, SML2299-5MG, Sigma, 5 mM stock made in DMSO) supplemented in 3D-RDM for 1 week by changing medium every other day. Three different concentrations of PR3 utilized in this study were 0.1, 0.25, and 0.5 µM. DMSO was used as a vehicle control. One-week, post-PR3 treatment, retinal organoids were collected and assessed by qRT-PCR, bulk RNA sequencing and imaging experiments.

All the data were obtained from three to six independent retinal differentiation experiments. The quantified data values were provided as mean ± SEM. The intergroup differences for all the analysis were determined with the GraphPad Prism v9, using a two-tailed student’s t-test or ANOVA. Significant differences were indicated by p values listed in the figures. For imaging studies, at least 3 sections were averaged to account for regional variability in iPSC reprograming and retinal organoid differentiation.