Molecular Vision 2016; 22:1077-1094
Received 20 May 2016 | Accepted 07 September 2016 | Published 09 September 2016
1Neurobiology-Neurodegeneration and Repair Laboratory, National Eye Institute, National Institutes of Health, Bethesda, MD; 2Genetic Engineering Core, National Eye Institute, National Institutes of Health, Bethesda, MD
Correspondence to: Anand Swaroop, Neurobiology-Neurodegeneration and Repair Laboratory, National Eye Institute, National Institutes of Health, Bethesda, MD 20892; Phone: (301) 435 5754; FAX: (301) 480 9917; email: email@example.com
Purpose: The generation of three-dimensional (3D) organoids with optic cup–like structures from pluripotent stem cells has created opportunities for investigating mammalian retinal development in vitro. However, retinal organoids in culture do not completely reflect the developmental state and in vivo architecture of the rod-dominant mouse retina. The goals of this study were to develop an efficient protocol for generating retinal organoids from stem cells and examine the morphogenesis of rods in vitro.
Methods: To assess rod photoreceptor differentiation in retinal organoids, we took advantage of Nrl-green fluorescent protein (GFP) mice that show rod-specific expression of GFP directed by the promoter of leucine zipper transcription factor NRL. Using embryonic and induced pluripotent stem cells (ESCs and iPSCs, respectively) derived from the Nrl-GFP mouse, we were successful in establishing long-term retinal organoid cultures using modified culture conditions (called High Efficiency Hypoxia Induced Generation of Photoreceptors in Retinal Organoids, or HIPRO).
Results: We demonstrated efficient differentiation of pluripotent stem cells to retinal structures. More than 70% of embryoid bodies formed optic vesicles at day (D) 7, >50% produced optic cups by D10, and most of them survived until at least D35. The HIPRO organoids included distinct inner retina neurons in a somewhat stratified architecture and mature Müller glia spanning the entire retina. Almost 70% of the cells in the retinal organoids were rod photoreceptors that exhibited elongated cilia. Transcriptome profiles of GFP+ rod photoreceptors, purified from organoids at D25–35, demonstrated a high correlation with the gene profiles of purified rods from the mouse retina at P2 to P6, indicating their early state of differentiation.
Conclusions: The 3D retinal organoids, generated by HIPRO method, closely mimic in vivo retinogenesis and provide an efficient in vitro model to investigate photoreceptor development and modeling disease pathology.
Rod and cone photoreceptors are specialized retinal neurons that initiate the visual process. While cone photoreceptors mediate color and daylight vision, rod photoreceptors function in dim light and can capture even a single photon . Dysfunction of photoreceptors is the primary cause of vision impairment in most retinal and macular degenerative diseases . Genetic and genomic studies have uncovered almost 250 genes for inherited retinal diseases (RetNet), and at least 52 common and rare variants at 34 distinct genetic loci have been associated with age-related macular degeneration [3,4]. At this stage, few treatment options are available. Gene therapy appears to have some initial success in treating visual dysfunction caused by specific inherited genetic mutations [5-8]. However, extensive genetic and phenotypic heterogeneity observed in retinal and macular degeneration strongly argues for the development of additional therapeutic strategies. Network- or pathway-based design of drugs and optogenetics have demonstrated promise in preclinical studies [9,10], and the feasibility of photoreceptor cell replacement has been demonstrated for repairing degenerated retinas in mice [11,12]. Transplanted retinal progenitors or rod photoreceptors from embryonic pluripotent stem cells (ESCs) or induced pluripotent stem cells (iPSCs) have been shown to integrate into the mouse retina, express photoreceptor-specific proteins, and partially restore some vision of recipient mice [13-17]. These proof-of-principle studies strongly suggest that transplantation of ESCs or iPSC-derived photoreceptors can be used as a promising approach for the treatment of retinal degeneration diseases.
With the advent of a three-dimensional (3D) culture system, retinal organoids can be generated from pluripotent stem cells using serum-free floating cultures of embryoid body-like aggregates with quick reaggregation (SFEBq) methodology [18,19]. The 3D retinal organoids seem to mimic the dynamic processes of retinal morphogenesis in vivo, including evagination of neuroepithelia to form optic vesicles and subsequent invagination to produce optic cups. The neural retina in 3D cultures appears to be structurally similar to the mammalian retina in vivo, with some lamination, polarization, and distinct cell types [18-20]. Transplantation of post-mitotic progenitors or of a retinal sheet from organoid cultures into the mouse degenerative retina shows better although limited integration of the photoreceptors, which exhibit outer segment and synapse formation [21,22]. Thus, retinal organoids potentially offer a valuable platform to investigate retina development and disease and may provide a resource for regenerative medicine.
The optic cup formation and subsequent differentiation of retinal neurons are inefficient using the current protocols as less than 70% of the organoids produce optic vesicles (evagination of neuroepithelia) at day (D)7 and only 18–35% generate optic cups (subsequent invagination of optic vesicles) by D10 [18,19,23,24]. Using most protocols, the neural retina in organoid cultures survives for approximately 25 days and exhibits limited development of inner retina neurons and a relatively smaller number of rod photoreceptors [21,23-25]. iPSCs derived from rods (compared to fibroblasts) exhibit higher efficiency of differentiation ; however, such a scenario might not be possible for human therapy.
The purpose of this study is to achieve high-efficiency differentiation of stem cells into retinal organoids, which recapitulate in vivo retinogenesis and include 70–75% of rod photoreceptors. To examine rod differentiation directly, we generated ESC and iPSC lines from Nrl-GFP mice, in which green fluorescent protein (GFP) expression is controlled by the promoter of the neural retina leucine zipper transcription factor Nrl . As Nrl determines the fate of the rod cells [27,28] and is the first rod lineage-specific marker , rod birth and subsequent differentiation can be followed with GFP expression. Using modified culture conditions (High Efficiency Hypoxia Induced Generation of Photoreceptors in Retinal Organoids, HIPRO), the retinal organoids from Nrl-GFP ESCs and iPSCs exhibited longer survival, high differentiation efficiency, and better stratification with the development of inner retina neurons. The rod photoreceptors in the retinal organoids at D35 accounted for almost 70% of all cells, and the rod photoreceptors’ transcriptome appeared similar to that reported for P6 rods from the mouse retina . The retinal organoid cultures, we report, thus provide a useful model for investigating early photoreceptor development, modeling degenerative disease, and exploring novel treatment paradigms.
ESC and iPSC clones were derived based on published protocols [31,32]. In brief, ESC clones were isolated from the inner cell mass (ICM) of 3.5-day blastocysts of Nrl-GFP transgenic mice on feeders of mouse embryonic fibroblast (MEF) in the presence of leukemia inhibitory factor (LIF; Millipore, Darmstadt, Germany) and 3i  and maintained under the same conditions at 37 °C in a humidified environment of 5% CO2. iPSC clones were generated by infection of the E14.5 MEFs from Nrl-GFP mice of Dox-inducible lentiviral vectors carrying Oct3/4, Sox2, Klf4, and c-Myc genes individually and then cultured on MEF feeder cells for 3 weeks. Embryonic stem cell–like colonies were picked and transferred onto fresh feeder cells, followed by evaluation of quality by morphology, proliferation rate, and Nanog expression. Nrl-GFP ESCs and iPSCs were maintained on feeder cells in knockout Dulbecco's Modified Eagle Medium (DMEM; Life Technologies, Carlsbad, CA, Catalog number: 11995073) supplemented with 1X Eagle′s minimum essential medium (MEM) non-essential amino acids (Sigma; St. Louis, MO, catalog number: M7145), 1X GlutaMAX (Life Technologies), 1X penicillin-streptomycin (PS; Life Technologies), 1×β-mercaptoethanol (Life Technologies), 2,000 U/ml Leukemia Inhibitory Factor (LIF; Millipore, Darmstadt, Germany), and 15% embryonic stem cell–qualified fetal bovine serum (FBS; Life Technologies). Cells were passaged using TrypLE Express (Life Technologies).
ESCs and iPSCs (passages 15–25) were differentiated using the SFEBq protocol  and after modifications in the culture conditions as follows: (i) The hypoxic condition was applied from D0 to D10; (ii) 36 uncut retinal organoids were cultured in a 100 mm Poly(2-hydroxyethyl methacrylate) (poly-HEMA)-coated (Sigma) dish after D14; (iii) β-mercaptoethanol was freshly added to the media every time; (iv) retinoic acid (RA; Sigma) was added to the culture from D14 to D26; (v) retinal organoids were supplied with DMEM/Nutrient Mixture F-12 (F12) with GlutaMAX (Life Technologies, catalog number: 10565042) supplemented with 2% FBS (Life Technologies), 1X penicillin-streptomycin (PS, Life Technologies), 1X MEM non-essential amino acid (NEAA, Sigma), 1X N2 supplement (Life Technologies), and 1X B27 supplement (Life Technologies); and (vi) 1X β-mercaptoethanol (Life Technologies) was added to all media every time. Then, 72 retinal organoids were transferred at D7 to a 100 mm poly-HEMA-coated Petri dish with 10 ml retinal maturation medium (DMEM/F12 with GlutaMAX with 1X PS). The maturation medium of the retinal cultures was changed at D10, and 1 mM taurine and 500 nM RA were added. At D14, half of the media was changed and the culture density reduced to 36 organoids per 100 mm dish. The cultures were maintained until D26 with half of the media changed every 3 days. At D26, the full media were replaced with DMEM/F12-containing GlutaMAX that was supplemented with 2% FBS, 1X PS, 1X NEAA, 1X N2, and 1X B27. Half of the media was then changed every 2 days. The cultures were incubated at 5% O2 from D0 to D10 and with 20% O2 from D10 onward.
ESCs and iPSCs were plated into four-well Nunc® Lab-Tek® II Chamber Slides (Sigma) at a density of 56×103 cells per well. After 48 h, the cells were washed once with DPBS (Dulbecco's phosphate-buffered saline (DPBS, 2.6 mM KCl, 1.47 mM KH2PO4, 137.93 mM NaCl, 8.06 mM Na2HPO4-7H2O, pH 7.0-7.3; Life Technologies, catalog number: 14190250) and fixed for 30 min using 4% paraformaldehyde (PFA; Electron Microscopy Science, Hatfield, PA). The cells were washed three times with DPBS and incubated for 1 h at room temperature in blocking solution containing DPBS with 10% FBS (Atlanta Biologicals, Flowery Branch, GA) and 0.2% Triton X-100 (Sigma). The cells were then incubated with anti-Nanog antibody (Mouse, 1:200 in blocking solution, ReproCELL, Kanagawa, Japan) overnight at 4 °C, followed by three washes of 5 min each with DPBS. Then 4',6-diamidino-2-phenylindole (DAPI; Life Technologies) and Alexa Fluor 568 anti-mouse antibody (1:250 in blocking solution, Life Technologies) were added, and the cells were incubated at room temperature for 1 h. After washing three times with DPBS (5 min each with gentle shaking), the cells were rinsed once with Ultrapure water (Life Technologies) to remove the sodium ions before the mounting solution was added. Fluorescence images were captured using the EVOS® XL system (Life Technologies).
For immunohistochemistry (IHC) of the differentiating neural retina, the organoids were fixed in 4% PFA (Electron Microscopy Science) for 1 h and cryoprotected in 15% sucrose for at least 2 h, followed by 30% sucrose overnight, before embedding in optimum cutting temperature (OCT) compound (Sakura Finetek, Torrance, CA). The OCT blocks were sectioned at 10 μm thickness and incubated at room temperature for at least 1 h, before immunostaining or storage at −80 °C. The immunostaining procedures have been recently described . Primary antibodies were used at the following dilutions: laminin (rabbit, 1:50; Sigma), Zonula occludens-1 (ZO-1, rabbit, 1:100; Life Technologies), brain-specific homeobox/POU domain protein 3A (Brn3a, rabbit, 1:200; Santa Cruz, Dallas, TX), calbindin (Calb, rabbit, 1:1,000; Calbiochem, San Diego, CA), S-opsin (Opnsw, goat, 1:200; Santa Cruz), Prospero homeobox 1 (Prox1, rabbit, 1:200; Abcam, Cambridge, MA), Ceh-10 homeodomain-containing homolog (Chx10, sheep, 1:200; Abcam), protein kinase C alpha (PKCα, rabbit, 1:1,000; Sigma), GFP (chick, 1:500; Abcam), rhodopsin (Rho, mouse, 1:1,000; a gift from Dr. Robert Molday, University of British Columbia, Canada), glutamine synthetase (GS, rabbit, 1:200; Abcam), ADP-ribosylation factor-like protein 13B (Arl13b, rabbit, 1:1,000; Proteintech, Rosemont, IL), gamma-tubulin (γ-tubulin, mouse, 1:500, Sigma), and synaptophysin (mouse, 1:200, Abcam). Sections were incubated with the primary antibody at 4 °C overnight. Species-specific secondary antibodies, conjugated with Alexa Fluor 488 or 568, were diluted in antibody dilution buffer (1:250, Life Technologies) and incubated at room temperature for 1.5 h. DAPI (Life Technologies) was used for nuclear staining. For comparison of intensity, images in the same section were captured using identical gain and voltage. Fluorescence images were captured using an LSM 880 confocal microscope (Zeiss, Jena, Germany).
ESCs and iPSCs were dissociated using TrypLE Express (Life Technologies). The cells were centrifuged at 180 ×g for 5 min and transferred to a T25 flask (Corning, Corning, NY) in 4 ml maintenance media to remove the feeder cells, which adhered to the flask in 20 min. Suspended cells were then fixed and stained with the FIX & PERM® Cell Fixation & Permeabilization Kit (Life Technologies) as suggested by the manufacturer. The cells were stained with anti-Nanog antibody overnight at 4 °C, followed by secondary antibody for 2 h at room temperature. The cells were then resuspended in DPBS containing 1 mM EDTA, filtered by 40 μm cell strainer (BD Bioscience, San Jose, CA), and kept on ice until fluorescence-activated cell sorting (FACS) analysis was performed using a MACSQuant Analyzer (Miltenyi Biotec, San Diego, CA).
For sorting the GFP+ cells, the retinal organoids were dissociated in 0.25% trypsin-EDTA (Life Technologies). The cells were centrifuged at 400 ×g for 5 min, resuspended in DPBS containing 1 mM EDTA, and filtered with a 40 μm cell strainer (BD Bioscience). The samples were kept on ice until sorting with FACSAriaII (BD Bioscience). Cell viability was evaluated by integrity (gated by DAPI) and size (gated by forward scatter, FSC). Gating of GFP+ cells was based on wild-type retinal organoids without GFP at the same differentiation day.
Total RNA was purified from tissue homogenates or flow-sorted GFP+ photoreceptors using TriPure isolation reagent (Roche, Basel, Switzerland). RNA quantity was assessed with a NanoDrop 1000 spectrophotometer (Thermo Scientific, Rockville, MD). Reverse transcription (RT) was performed using a SuperScript III Reverse Transcription-PCR kit (Life Technologies) and complementary DNA (cDNA) was generated using oligo(dT) primer. The resulting cDNA was amplified using gene-specific primers (Table 1). PCR products were separated by 1% agarose gel (Lonza, Walkersville, MD) and detected under ultraviolet (UV) illumination (PerkinElmer, Waltham, MA).
RNA sequence (RNA-seq) profiles of high-quality total RNA (RNA integrity number ≥7.5) from the GFP+ photoreceptors were obtained using the Illumina platform. These paired-end RNA-seq data were compared to reported single-end mouse rod transcriptome  as follows. After the Illumina adapters were trimmed, defined in the TruSeq3-PE-2.fa file, by Trimmomatic (version 0.36)  from paired-end data, each sample was quantified against Ensembl release 84  cDNA FASTA file by Kallisto (version 0.42.5)  via a custom pipeline in Perl (Perl version 5.16.3). The quantifications were then normalized and summarized at the gene level with tximport  with the option “countFromAbundance=lengthScaledTPM.” The statistical environment R (version 3.3.1) installed on a 64-bit GNU Red Hat Linux system (Centos 7.2) was used to further analyze the data and to generate heatmaps. The count values from tximport were converted to count per million (CPM) via edgeR . Logarithmically transformed (base 2) CPM values were used for the PCA and clustering. All GFP+ data were averaged for each time point to avoid replicates. The heatmap function has been used previously . Photoreceptor genes data were clustered with the affinity propagation algorithm  using the negative distance matrix option as the similarity measure. PCA analysis was performed with custom plotting, using prcomp function in R without scaling and centering the data. To highlight the time matching, the first principal component was plotted against the 11th component on the x- and y-axes, respectively.
All data were expressed as mean ± standard error of the mean (SEM), unless specified. An unpaired two-tailed Student t test was used to compare the mean between two groups. Results with a p value of less than 0.05 were considered statistically significant. At least three independent biologic replicates were performed (n = number of experiments).
Nrl-GFP ESCs and iPSCs were evaluated by the morphology, proliferation rate, and expression of Nanog, which is a critical transcription factor for pluripotency . There was no significant difference between the ESCs and iPSCs in terms of morphology (Figure 1A) and proliferation rate (Figure 1B), both of which were consistent with other publications [41,42]. Nanog expression was detected in the nuclei of all ESC and iPSC colonies (Figure 1C). Flow analysis showed that all ESCs and iPSCs were pluripotent and exhibited Nanog expression (Figure 1D).
The SFEBq protocol was modified to efficiently obtain a more mature neural retina by changing the oxygen level and medium components during differentiation (Figure 2A). Oxygen tension has been demonstrated to impact the viability and cell fate commitment of neurons during early neuronal development in mice and humans [43,44]. Thus, the hypoxic condition was applied from D0 to D10 in the modified protocol. Under normoxia (20% O2) and hypoxia (5% O2) conditions, mouse ESCs and iPSCs could be differentiated into embryoid bodies with a neuroepithelium at D4, an optic vesicle at D7, and an optic cup at D10, without significant morphological differences (Figure 2B). To assess the efficiency, differentiating organoids with optic vesicle and optic cup formation were quantified at D7 and D10, respectively. Organoids in hypoxic condition demonstrated significantly higher differentiation into optic vesicles and optic cups (Figure 2C). For ESCs and iPSCs, more than 70% of the organoids formed optic vesicles by D7 (n = 10, range from 50% to 95%), and more than 50% formed optic cups on D10 (n = 10, range from 30% to 85%), compared with 40% at D7 (n = 10, range from 30% to 50%) and less than 20% (n = 10, range from 8% to 30%) at D10 with the original SFEBq method. To avoid bias in counting, 60 organoids at D10 were randomly selected and harvested for RNA. RT–PCR analysis was performed to measure the expression of early eye field transcription factors. As predicted, the embryoid bodies under hypoxic conditions revealed higher expression of Pax6 and Rax, suggesting commitment to the retinal cell fate by a greater number of cells (Figure 2D).
To investigate whether the neural retinas generated by the modified protocol mimic the in vivo mouse retina in terms of polarity, lamination, and the presence of all cell types, immunostaining was performed using various retinal markers. Apical-basal polarity of the neural retina is critical for the proper functioning of photoreceptors . The neural retinas generated using the modified protocol revealed polarity, with the outer side basal and the inner side apical (Figure 3A).
Ganglion cells and horizontal cells are the two earliest-born cell types in 3D retinas, as evident by the expression of Brn3a and calbindin (Calb), respectively (Figure 3B). As early as D14, ganglion cells could be detected at the basal side of the neural retina, with a substantial increase in the number at D18. However, the number of ganglion cells decreased at D21, which may be due to the lack of axon formation or other secretory factors from the brain in culture [46,47]. Horizontal cells started to be born at D14, and few cells were detected at the middle part of the neural retina. At D18, the horizontal cells migrated to the basal side, and their number greatly expanded at D21, with limited connections among the cells.
The birth of ganglion cells and horizontal cells is followed by the generation of cone photoreceptors and amacrine cells (Figure 4). At D18, cones and amacrine cells were detected with minimal expression of S-opsin (Opnsw) and Prox1, respectively, at the middle part of the neuroblastic layer (NBL). At D26, cone cells were found at the apical side of the NBL, while amacrine cells were mainly located at the basal side. With the development of amacrine cells, the inner plexiform layer (IPL) started to form at this stage. Further maturation of the cone photoreceptors and amacrine cells was indicated by the significant increase in expression of Opnsw and Prox1, respectively. At D32, the cone photoreceptors had typical morphology and were located in the outer nuclear layer (ONL), which is at the apical side of the neural retina. Notably, the outer plexiform layer (OPL) and the IPL were detected at this stage. Most amacrine cells were located at the inner nuclear layer (INL), with some misplaced at the ganglion cell layer (GCL).
The Nrl-GFP ESC and iPSC lines were also differentiated using the original SFEBq protocol for comparison (Appendix 1). At D26, the inner retina neurons were not fully developed, and only the NBL and the GCL could be observed. The morphology of the neural retina in these experiments was similar to that reported by others [23,24,48].
During in vivo retinal development, formation of the OPL and the IPL is followed by the development of bipolar cells to connect the two plexiform layers . Chx10 is a marker for proliferative cells or bipolar cells, depending on the differentiation stage. At an early stage of differentiation (D18), proliferative cells were found throughout the NBL, as indicated by Chx10 staining. As the cells started to commit bipolar cell fate, a small portion of Chx10+ cells were located at the basal side of the NBL at D26, while most remained in the NBL, which were proliferative retinal progenitor cells. With further maturation of the neural retina, all cells exited the cell cycle at D32, and Chx10+ cells were detected only at the ONL, indicating the proper location of bipolar cells. PKCα is the marker for ON-bipolar cells, and its expression indicates the maturation of bipolar cells. A substantial expression of PKCα was detected only in the late stage of differentiation, connecting the OPL and the IPL (Figure 5).
Nrl is the earliest marker of post-mitotic rods; Nrl expression is detected at rod birth, followed by Rho expression as the differentiation proceeds [26,29]. RT–PCR analysis of the neural retina showed that Nrl expression began at D15, with a continuous increase during differentiation (Figure 6A). A minimal amount of rhodopsin (Rho) expression was detected at D18 and increased with time as the rod photoreceptors matured (Figure 6A). As GFP is under the control of the Nrl promoter in ESCs and iPSCs, differentiating cells committed to rod cell fate would express GFP in organoids. Flow analysis of the dissociated organoids revealed a progressive increase in GFP+ cells and in the intensity of GFP (Figure 6B). Notably, the GFP+ rod cells constituted more than 70% of the total population at D35, closely mimicking the mature mouse retina in vivo  (Figure 6C). At D18, there were few GFP+ cells with weak signal intensity, but more GFP+ cells with higher intensity were detected by D21 (Figure 6B,D). After D25, GFP seemed polarized to the apical side of the neural retina, with D35 having the highest GFP+ cell number and signal intensity (Figure 6D). Thus, the retinal organoids generated by the HIPRO protocol demonstrated longer survival and more GFP+ rods compared to the original SFEBq protocol (Appendix 1).
Most rod photoreceptors and Müller glia are born postnatally in the mouse retina . At D18, rod photoreceptors were evident by the minimal expression of Reep6, a marker for rod photoreceptors , at the apical side of the neural retina. Expression of Reep6 increased at D25, indicating the maturation of the rods. In addition, GS, a marker for Müller glia, was mainly located at the basal side, while Reep6 was polarized to the apical side. From D32 and onward, the Müller glia matured further and spanned the entire neural retina, and the outer limiting membrane (OLM) was evident at the apical side of NR, with Reep6 protruding beyond the OLM. With the HIPRO protocol, the organoid cultures were maintained until D35, with an intact structure of the neural retina (Figure 7).
Photoreceptor outer segments originate as modified primary cilium [52,53], and limited ciliogenesis has been one of the shortcomings of 3D retinal organoid cultures [18,19,21,54]. Using the HIPRO protocol, we facilitated cilia growth in retinal organoids and detected elongation of the ciliary axoneme at the apical side of the neural retina (Figure 8A). At D25, basal bodies, indicated by immunostaining of γ-tubulin, were polarized to the apical side of the neural retina, suggesting the start of ciliogenesis. At this stage, immunostaining of ADP-ribosylation factor-like protein 13B (Arl13b), a marker of ciliary axoneme, was barely detectable. As the photoreceptors continued to mature, more cells with elongated cilium were observed at D32, as indicated by Arl13b. Colabeling with Rho and Arl13b demonstrated their partial colocalization, with Rho immunostaining extending beyond Arl13b, suggesting the initiation of outer segment biogenesis (Figure 8B).
In addition, we observed the formation of plexiform layers, as indicated by synaptophysin immunostaining. At D26, only the IPL was detected with a trace amount of synaptic vesicles. As the development proceeded, the OPL was detected with significant labeling with synaptophysin. Increased synaptic staining was also detected in the IPL at D32, suggesting further maturation of the INL cell types (Figure 8C).
After the transcriptome data from the GFP+ rods derived from the D25, D28, and D35 organoids were normalized with the published RNA-seq of the developing mouse rods , we applied PCA to the whole data set (Figure 9A). The principal component 1 (PC1) captured more than 30% of all variations in the data and perfectly captured the variance between time points. The PCA plot showed that the D25 and D28 data coprojected on PC1 with the P2 and P4 time points of the mouse rods, whereas D35 falls between P6 and P10 but closer to the P6 developmental stage. A heatmap showing the expression profiles of the photoreceptor genes validated the PCA results and further demonstrated that the D35 rods in the organoid culture were more mature than in the P6 retina (Figure 9B). Notably, the developing rod photoreceptors in the retinal organoids also expressed cone genes, consistent with the proposed concept of the potential origin of mammalian rods from S-cones .
Here, we described a modified differentiation protocol, called HIPRO, for mouse retinal organoid culture, with efficient production of a more mature neural retina mimicking in vivo retinogenesis. A striking observation in the study is the efficient generation of optic vesicles and optic cups by applying hypoxic conditions to organoid cultures from D0 to D10. The efficiency of optic cup formation is reportedly low under normoxic condition [18,23-25], except in one study that demonstrated high efficiency of optic cup formation by maintaining Matrigel until D10 instead of D7 . Although differentiation efficiency varies among clones and batches , the concurrent comparison of organoids derived from ESCs and iPSCs in the present study showed a significant improvement in optic cup formation when hypoxic conditions are provided from D0 to D10. Hypoxia has been shown to promote the proliferation and differentiation of neural stem cells . Formation of the optic cup largely depends on the size of the optic vesicles [19,23], and the vesicles formed under hypoxia tend to be bigger with a higher probability of optic cup formation. The present results are consistent with those of a study that demonstrated higher efficiency and number of rod photoreceptors cells in two-dimensional (2D) culture under hypoxia conditions .
ESCs and iPSCs derived from Nrl-GFP mice were used to investigate rod birth and differentiation in vitro. As Nrl is the earliest rod-specific marker , rod photoreceptors in organoid culture are marked by GFP and can be quantified as differentiation proceeds. Thus, we can reliably document the progressive increase in the number of GFP+ cells and GFP intensity in retinal organoid cultures, which include more than 70% GFP+ rods at D35. The GFP+ cells may have been underestimated as intact retinal organoids with some non-retinal tissues were used for quantification. Nevertheless, the data are comparable to 75–80% rods in the mouse retina in vivo  and consistent with the reported percentage of photoreceptors in retinal organoids from mouse ESCs . Transcriptome analysis of D35 GFP+ cells from retinal organoids revealed their maturation beyond the developing rods in the P6 mouse retina. Efficient differentiation of rods from Nrl-GFP ESCs and iPSCs should facilitate their analysis and use in transplantation studies, without additional steps for labeling .
With the use of the HIPRO protocol, the organoid cultures were maintained until at least D35. At this stage, the neural retina contained all major cell types, including rod and cone photoreceptors. Limited differentiation of the inner retina neurons from iPSCs derived from fibroblasts has been reported to reflect epigenetic memory ; however, the neural retina derived with the HIPRO protocol was stratified, with a substantial number of bipolar and amacrine cells and Müller glia. Consistent with other studies [21,24], retinal ganglion cells are scarce, especially at the late stage of differentiation. As the survival of ganglion cells requires sufficient oxygen and nutrients [57,58], static culture may not be useful for maintaining their long-term survival, and dynamic culture (e.g., using a bioreactor) may be required for retinal organoids.
Another significant observation in the retinal organoids in the present study was the growth of cilia and the initiation of outer segment morphogenesis. As reported previously, proper development of the outer segments requires signaling from the RPE . Previous studies have reported the presence of the RPE in organoid culture but only at an early stage of differentiation [18,19,21]. The RPE appears to be lost as retinal cells proliferate and detach. Coculture or explant culture of the RPE with retinal organoids would be necessary for outer segment biogenesis and phototransduction.
Retinal organoid cultures have wide applications for studies of organ development, disease modeling, and/or treatment design [12,60-64]. With the development of genome editing tools, such as CRISPR/Cas9, a specific cell type can be labeled by fluorescence markers and purified to decipher gene regulatory networks [20,65] or as a source of transplantation [21,23,66]. The present study provides a useful system for efficiently generating retinal organoids with a mature neural retina and provide a framework for developing a more advanced differentiation platform by augmenting nutrients and oxygen supply using bioengineering tools (such as a bioreactor).
We are grateful to Pinghu Liu, Samantha Papal, Jung-woong Kim, Hiroko Shimada-Ishii, Jacklyn Mahgerefteh, Tyler DiStefano and Jacob Nellissery for insightful discussions and constructive comments. We thank Linn Gieser and Matthew Brooks for assistance with RNA-seq, and Julie Laux, Jessica Albrecht, and Rafael Villasmil for help with flow cytometry. This work was supported by intramural research program of the National Eye Institute (ZIAEY000450, ZIAEY000474).