Generation and Staging of Human Retinal Organoids Based on Self-Formed Ectodermal Autonomous Multi-Zone System

Abstract

Methods for stem cell-derived, three-dimensional retinal organoids induction have been established and shown great potential for retinal development modeling and drug screening. Herein, we reported an exogenous-factors-free and robust method to generate retinal organoids based on "self-formed ectodermal autonomous multi-zone" (SEAM) system, a two-dimensional induction scheme that can synchronously generate multiple ocular cell lineages. Characterized by distinct morphological changes, the differentiation of the obtained retinal organoids could be staged into the early and late differentiation phases. During the early differentiation stage, retinal ganglion cells, cone photoreceptor cells (PRs), amacrine cells, and horizontal cells developed; whereas rod PRs, bipolar cells, and Müller glial cells were generated in the late differentiation phase, resembling early-phase and late-phase retinogenesis in vivo. Additionally, we modified the maintenance strategy for the retinal organoids and successfully promoted their long-term survival. Using 3D immunofluorescence image reconstruction and transmission electron microscopy, the substantial mature PRs with outer segment, inner segment and ribbon synapse were demonstrated. Besides, the retinal pigment epithelium (RPE) was induced with distinct boundary and the formation of ciliary margin was observed by co-suspending retina organoids with the zone containing RPE. The obtained RPE could be expanded and displayed similar marker expression, ultrastructural feature and functional phagocytosis to native RPE. Thus, this research described a simple and robust system which enabled generation of retina organoids with substantial mature PRs, RPE and the ciliary margin without the need of exogenous factors, providing a new platform for research of retinogenesis and retinal translational application.

Keywords: RPE; SEAM system; ciliary margin; human retinal organoid; photoreceptor cell; retinogenesis.

FIGURE 1

Robust retinal organoids induced by the modified SEAM method. (A,B) Expression pattern of key transcription factors during the induction of retinal organoids. Immunofluorescent staining of eye field marker SIX3 (red) and PAX6 (green) on Day 10 (A); the retinal progenitor cell marker CHX10 (red) on Day 10 (A’); the neural progenitor cell markers OTX2 (red) and SOX2 (green) on Day 20 (B). Scale bar = 100 μm. (C) The ratio of OTX2-, SOX2-, and CHX10-labeled clusters to total clusters (marked by DAPI) on Day 28 in the SEAM- and RDM-treated groups, respectively. The bars represent the mean ± SEM. **P < 0.01 (n = 3). (D) Bright-field imaging showing the derivation of cup-like neural structure and the surrounding pigment cell on Day 31. The identification of the retinal progenitor cell marker CHX10 (red, in the 3D cup-like neural structure) and retinal pigment epithelial cell marker MITF (green, at the bottom of the 3D cup-like neural structure) (D’). Scale bar = 100 μm. (E) Average number of retinal organoids obtained per well in 6-well plates on Day 28, Day 32, Day 37, Day 40, and Day 44 (n = 3). (F) Schematic diagram of the induction of human retinal using the modified SEAM method.

FIGURE 2

Morphogenesis of retinal organoids. (A) Bright-field imaging showing the cup-like structure of the retinal organoid immediately after isolation. Scale bar = 100 μm. (B) Variation in the growth rate of retinal organoids at different time points. Scale bar = 500 μm. (C,D) The growth rate was quantified by sphere size [(C), n = 10 retinal organoids from three independent experiments] and neural retina thickness [(D), n = 15 retinal organoids from three independent experiments]. (E) Representative images at Week 4, Week 6, and Week 23 showing the changes in the thickness of the neural retina (dashed line). Scale bar = 100 μm. (F) Ki67 immunostaining of the neural retina at Week 3, Week 7, Week 19, and Week 36, indicating the downregulation of proliferation as differentiation proceeded. The neural retina thickness was measured as illustrated by the arrows. Scale bar = 50 μm.

FIGURE 3

The cellular composition of retinal organoids during the early-stage differentiation. (A) The cellular dynamics of retinal ganglion cells were marked at Week 5 (BRN3, red; TUJ1, green), and Week 8 (BRN3, red; TUJ1, green). (B) Statistics analysis of the ratio of BRN3 + cells versus total cells (marked by DAPI) at Week 5 and Week 8, respectively. Mean ± SD, ***P < 0.0005. (C) Localization of CRX⁺ cone photoreceptor cells at Week 5. (D) The onset of AP2α + amacrine cells and AP2α⁺/PROX1⁺ horizontal cells were detected (AP-2α, green; PROX1-red) at Week 5 and confirmed by CALBINDIN (green, D’) staining at Week 7. Scale bar = 50 μm.

FIGURE 4

The cellular composition of retinal organoids during the late-stage differentiation. (A) Robust derived cone and rod photoreceptor cells at Week 13 as indicated by CRX (green) and RECOVERIN (red), respectively. Scale bar = 100 μm. (B) The co-localization of CHX10 (red) and α-PKC (green) in the presumptive inner nuclear layer (INL), suggesting the generation of bipolar cells at Week 18. (C) The nuclei-free layer between the outer nuclear layer (ONL) and the INL was illustrated by DAPI immunostaining, indicating the outer plexiform layer (OPL, arrowhead). (D) Müller glial cells were detected at Week 23 and were rapidly generated at Week 36 (D’), as shown by immunostaining of the Müller glial cell markers SOX9 (green) and GS (red). Scale bar = 50 μm.

FIGURE 5

Maturation of photoreceptor cells in retinal organoids. (A) Representative images showing the onset of hair-like microvilli on the surface of the retinal organoid at Week 22, Week 26, and (A’) Week 32. Scale bar = 100 μm. (B) Progressive increase in the number of retinal organoids with hair-like microvilli (n = 3 independent experiments per group; each group contained 18, 12, and 13 retinal organoids, respectively). (C,D) Representative images showing the maturation of cone and rod PRs, as indicated by the mature PRs markers Opsin Green/Red (C, Opsin G/R, green) and RHODOPSIN (D, red) at Week 18, Week 23, and Week 29. Scale bar = 50 μm. (E) Immunofluorescent staining of the PR marker ARRESTIN 3 (red) and CRX (green) showed that a 4/5-nuclei–thick layer was formed. (E’) Higher magnification of (E) showing the distinct morphology of the cone outer segment (asterisks) and the rod outer segment (triangle). Scale bar = 50 μm. (F) The co-staining of ARRESTIN 3 (red) and vesicular transporter marker VGLUT1 (green) indicated the formation of the PR synapse. Scale bar = 50 μm. (G) Electron microscopy showed the formation of outer segments-like (OS) and inner segments-like structure (IS), outer limiting membrane (OLM, indicated by black arrow) and outer nuclear layer (ONL). Scale bar = 5 μm. The insert image showed the infolding disk-like structure in OS. Scale bar = 0.5 μm. (H) The 4/5-nuclei–thick ONL was shown. The black arrow indicated that the ribbon synapses-like structure were present at basal side of photoreceptors. Scale bar = 10 μm. The insert image showed the mitochondria-rich inner segment-like structure. The white arrow indicated the OLM. Scale bar = 2 μm. (I) The clearer image of ribbon synapse-like structure (RS) was shown, surrounded by various vesicles. Scale bar = 500 nm (The insert image scale bar = 50 nm).

FIGURE 6

The uneven distribution of rod and cone photoreceptor cells. (A) The ortho X-Z display of a random section of the organoid at Week 42 showed that the hinge distal part was mainly composed of cone PRs (Opsin B/G/R, green, purple frame) and rod PRs (Rhodopsin, red, green frame) mainly located at the hinge proximal part. (B) Ortho display of maximum intensity projection of the whole organoid at Week 42. The fluorescence intensity from the hinge proximal to distal part (marker by the red box) was calculated in (B’). Scale bar = 200 μm. (C) The ratio of the numbers of cone PRs versus rod PRs were paired calculated in the proximal and distal part, respectively. **P < 0.005.

FIGURE 7

Autonomous generation of the retinal pigment epithelium and the ciliary margin. (A) Representative images showed the autonomous formation of a four concentric-zone cluster. Immunofluorescent staining showed that zone 1 and zone 2 were CHX10- (green) and MITF- (red) positive, respectively. Scale bar = 100 μm. (B) Compared with isolation without zone 2 [zone 2 (–)], isolation of retinal organoids with zone 2 [zone 2 (+)] generated more RPE spheres (n = 3 independent experiments per group; each group contained 20 or more retinal organoids). (C) Representative images showing the gradual formation of the peripheral neural retina between the neural retina and the RPE (dashed circle). Scale bar = 500 μm. (D) Co-staining of the ciliary margin marker PAX6 (green) and CX43 (red) confirmed the generation of ciliary margin zone at Week 32 (triangle). Scale bar = 200 μm.

FIGURE 8

Expansion and validation of the retina pigment epithelium (RPE). (A,B) Representative images showing the expansion and pigmentation loss of the RPE on Day 1, Day 4, and Day 7 (A); and the re-pigmentation on Day 24 and Day 36 (B). (B’) Higher magnification images showing the formation of elevated domes (asterisks). (C) Expression of the naïve RPE marker OTX2 (red, left), PAX6 (green, middle) and the RPE marker ZO-1 (green, right) was shown on Day 7, while the mature RPE marker RPE65 (red, right) was not found. (D) On Day 44, RPE65 (red, left) coupled with the RPE marker MITF (red, middle) and ZO-1 (green, right) indicated the maturation of the RPE. Scale bar = 100 μm. (E) Electron microscopy showed that on Day 44, RPE sheet has developed melanin granules (mg), microvilli (mv) and the abundant tight junctions at the apical side (indicated by black arrow). Scale bar = 2 μm. (F) Different cell junctions of RPE were clearly shown (apical tight junctions, indicated by black arrow; desmosomes, indicated by white arrow). Scale bar = 500 nm. (G) Phagocytosis assay showed that the 2-week-old RPE sheets, labeled by ZO-1 (green), were able to phagocytose FluoroSpheres (red). Scale bar = 20 μm.