Creating patient stem cell–based 3D retina models with transplant-ready light-sensing cells

Reprogramming and clonal expansion were performed using the Cell X™ platform (Cell X Technologies Inc), which is contained within a cGMP compliant Biospherix Xvivo system (BioSpherix, Ltd.) [10]. Dermal fibroblasts isolated from 3 mm dermal punch biopsies obtained from the individuals listed in Table S1 were cultured in Fibroblast Medium (FBM; DMEM, 10% Fetal Bovine Serum (FBS; Atlas Biologicals; Cat#F-0500-A) and 0.2% Primocin (InvivoGen; Cat#ant-pm-2)) at 37° C, 5% CO2, and 20% O2 as described previously [6]. Fibroblasts were transduced using the CytoTune 2.0 kit (Thermo Fisher Scientific; Cat#A16517), a non-integrating Sendai viral reprogramming kit [6] at 20% O2. After 5 days, cells were passaged onto rhlaminin 521 (5 µg/ml; Biolamina; Cat#LN521-02) coated dishes and the next day the media was changed to Essential 8 (E8) medium (Thermo Fisher Scientific; Cat#A1517001). To enhance reprogramming efficiency, the oxygen tension was reduced from 20% to 5% at 1 week following transduction. The number and diameter of iPSC colonies present in the entire plate was determined using automated iPSC colony detection and segmentation software as described previously [13]. At passage 10, iPSC lines were subject to karyotyping and scorecard analysis as described previously [10].

For TaqMan hPSC ScoreCard™ analysis, total RNA was isolated using the NucleoSpin RNA purification kit (Macherey-Nagel; Cat#740955). cDNA was generated from 1 µg of RNA using VILO cDNA synthesis kit (Thermo Fisher Scientific; Cat#11754050). cDNA was added to a TaqMan hPSC scorecard plate (Thermo Fisher Scientific; Cat#A15870) and amplified using a QuantStudio 6 Flex real-time PCR system. Results were analyzed using Thermo Fisher's cloud-based analysis suite.

Retinal differentiation was performed with modifications for GMP-compliance [8, 10, 11]. Briefly, iPSCs were cultured on recombinant human rhlaminin 521 coated cell culture plates in E8 medium. To generate embryoid bodies (EBs), iPSCs were lifted using ReLeSR™ (STEMCELL Technologies; Cat#05872) rather than dispase (as per our standard protocol [14]. Cell clusters were transitioned from E8 to Neural Induction Medium (NIM- DMEM/F12 (1:1) (Thermo Fisher Scientific; Cat#11320033), 1% N2 supplement (Thermo Fisher Scientific; Cat#17502048), 1% non-essential amino acids (Thermo Fisher Scientific; Cat#11140050), 1% Glutamax (Thermo Fisher Scientific; Cat#35050061), 2 µg/mL heparin (MilliporeSigma; Cat#H3149) and 0.2% Primocin (InvivoGen; Cat#ant-pm-2)) over a four-day period. On day 6, NIM was supplemented with 1.5 nM rhBMP4 (R&D Systems; Cat#314-BP-05/CF). On day 7, EBs were adhered to Matrigel™ (Corning; Cat#354234 – standard protocol) or the following xeno-free substrates: CELLstart™ (Thermo Fisher Scientific; Cat#A1014201), a mixture of recombinant human laminin 111 (LN111) (5 µg/ml; Biolamina; Cat#LN111-02), recombinant human nidogen-1 (5 µg/ml; R&D Systems; Cat#2570-ND-050) and recombinant human collagen IV (18 µg/ml; MilliporeSigma; Cat#CC076), or recombinant human LN111 only. After initial testing of all substrates (Fig. 1), further experiments were done with CELLstart. BMP4 was gradually transitioned out of the NIM over seven days. On day 16, the media was changed to Retinal Differentiation Medium (RDM - DMEM/F12 (3:1), 2% B27 supplement (Thermo Fisher Scientific; Cat#17504044), 1% non-essential amino acids, 1% Glutamax and 0.2% Primocin). On day 30 the entire outgrowth was mechanically lifted using the back end of a 1 ml pipette tip and transferred to ultra-low attachment flasks in xeno-free 3D-RDM (RDM plus 10% FBS (as per our standard protocol) or 10% KnockOut™ Serum Replacement (KSR; Thermo Fisher Scientific; Cat#10828028), 100 µM taurine (MilliporeSigma; Cat#T0625), 1:1000 chemically defined lipid concentrate (Thermo Fisher Scientific; Cat#11905031), and 1 µM all-trans retinoic acid (MilliporeSigma; Cat#R2625). The cells were fed three times per week with 3D-RDM until differentiation day 100 when media was switched to 3D-RDM minus (i.e., 3D-RDM without all-trans retinoic acid). Organoids were feed 3 days per week with 3D-RDM minus until harvest.

Retinal organoids were dissociated at differentiation day 160 as previously described (PMIDs 40504625, 38652563). Briefly organoids were allowed to settle and washed with EBSS buffer (Worthington Biochemical Corporation; Cat#LS006361). Organoids were dissociated with 35 units/mL of papain (Worthington Biochemical Corporation; Cat#LK003176) and 150 units/mL DNase I (Worthington Biochemical Corporation; LS006361) in EBSS buffer. Organoids were incubated in a 37˚C water bath for 45 min and were pipetted every 15 min to break up remaining tissue chunks. Following dissociation, cells were put through a 40 μm strainer, pelleted and resuspended in HBSS plus calcium, magnesium and N-acetyl-cysteine (NAC, 0.5 mM) (Spectrum Chemical Corporation; Cat#AC800).

The work has been reported in line with the ARRIVE guidelines 2.0. Animals were housed in a dedicated barrier facility at the University of Iowa in filtered polysulfone cages with water bottles and a standard twelve-hour light: dark cycle. Animals were given access to food and water ad libitum and co-housing offered as a supplemental environment enrichment, when appropriate. We recently described the generation of a Pde6b-null rat model of retinal degeneration via CRISPR-mediated genome editing [15]. This model is albino and has an early onset rapidly progressive disease resulting in near complete loss of photoreceptor cells by two-months of age, making it an ideal model to test survival of stem cell-derived photoreceptor cells. For the immunosuppression protocol development studies, 2-month-old Pde6b-null rats (N = 16; non-immunosuppressed controls = 3 M, 5 F; immunosuppressed = 3 M, 5 F) were used. For the subsequent cell replacement studies, two groups of 2-month-old Pde6b-null rats were used. Group 1 was sacrificed at 3-days (N = 11; 6 M, 5 F) and group 2 was sacrificed at 4-weeks (N = 11; 7 M, 4 F) following subretinal injection of 2,000,000 human iPSC-derived retinal cells. In each experiment, one eye received subretinal injections of cells while the contralateral eye was injected with sterile buffer as a vehicle control.

For immunosuppression prior to xenograft injections, rats were first treated with intraperitoneal injection of cyclosporine (10 mg/kg) daily for seven days starting two days prior to subretinal injection of human iPSC-derived retinal cells (i.e., day − 2). Following cyclosporine injection, immunosuppression was continued via cyclosporine-supplemented drinking water (100 µg/mL) ad libitum until animal euthanasia.

For subretinal injections of retinal cell suspensions, rats were placed under anesthesia with a weight-dependent cocktail of intraperitoneal ketamine (91 mg/kg) and xylazine (9.1 mg/kg). Once anesthetized, eyes were dilated with 1% tropicamide and administered artificial tears ointment to prevent drying of the cornea. Using an operating microscope, a limited conjunctival peritomy was created in the temporal quadrant using 0.12 mm forceps and Vannas scissors. Subretinal injections were performed using a 33-gauge blunt-tipped Hamilton syringe as previously described [15 –18]. All injections were performed by a fellowship-trained vitreoretinal surgeon (I.C.H.).

Each eye was immediately imaged post-injection using a rodent-specific fundus camera and optical coherence tomography instrument (Micron IV + OCT; Phoenix MICRON Image-Guided OCT2, Phoenix Laboratories) to confirm the presence and location of a subretinal bleb and to check for the absence of complications (e.g., perforation of the retina into the vitreous cavity or subretinal hemorrhage) as described previously [15 –18]. The contralateral eye of each animal was injected with sterile buffer as a vehicle control. Following subretinal injection, animals were monitored daily for 72 h and weekly thereafter and evaluated for signs of inflammation, ocular injury, general distress including social withdrawal, emaciation, stilted gait, hyperactivity or vocalization. Any adverse events were immediately noted and, if necessary, animals immediately humanely euthanized via CO₂ inhalation or Euthasol. At appropriate study timeline endpoints animals were euthanized in the same manner.

Retinal organoids were fixed with 4% paraformaldehyde for 30–60 min at room temperature and equilibrated to 15% sucrose in PBS, followed by 30% sucrose. Organoids were cryopreserved in 50:50 solution of 30% sucrose/PBS: tissue freezing medium (Electron Microscopy Sciences; Cat#72592) and cryosectioned (15 μm).

For eyes that received subretinal injection of a single cell suspension, the injection site was identified by fundus examination, and a limbal suture was placed at the corresponding clock hour of the injection for orientation when embedding tissues for sectioning, as described previously [15 –18]. Rat eyes were enucleated and fixed in 4% paraformaldehyde for at least 1 h prior to dissection of the anterior segment. The posterior eye cup was fixed in 4% paraformaldehyde overnight at 4 °C and rinsed in increasing concentrations of sucrose. The eye cup was oriented using the previously-placed limbal suture to identify the injection site, embedded in 2:1 solution of Tissue-Tek OCT compound (VWR International; Cat#4583) and 20% sucrose, flash-frozen in liquid nitrogen, stored at -80 °C, and sectioned at a thickness of 7 μm using a Microm HM505E (Microm) or Leica cryostat as previously described [15 –18].

Retinal organoid sections were blocked in PBS containing 5% normal donkey serum (MilliporeSigma; Cat#S30), 3% bovine serum albumin (BSA; Research Products International Corp.; Cat#A30075), and 0.1% triton-x. Retinal sections were blocked in PBS containing 3% BSA at room temperature for an hour. Primary antibodies (Table S2) were diluted in blocking buffer and allowed to incubate for 2 hours at room temperature or overnight at 4°C. Following washes in PBS, sections were incubated with appropriate Alexa Fluor fluorescent-conjugated secondary antibodies (Table S2). 4',6-Diamidino-2-phenylindole dihydrochloride (DAPI; Thermo Fisher Scientific; Cat#62248) was used to counterstain cell nuclei. Sections were mounted using Aqua-Mount Mounting Medium (Thermo Fisher Scientific; Cat#14-390-5) and then visualized using a Leica TCS SPE upright confocal microscope system (Leica Microsystems).

H&E staining was carried out in standard fashion. Briefly, slides were: rinsed in fresh distilled water for 60 s, incubated in Hematoxylin (Epridia, Portsmouth, NH; Cat#: 7211) solution for 90 s, rinsed via repeated dipping (10 times) in fresh distilled water in two consecutive containers, dipped twice in Clarifier 1 Solution (Epridia; Cat#: 7401), rinsed via a single dip in fresh distilled water, incubated in Bluing Reagent (Epridia; Cat#: 7301) for 60 s, rinsed once in distilled water and incubated in 80% ethanol for 60 s. Slides were then incubated in Eosin-Y (Epridia; Cat#: 71304) for 30 s and then dipped repeatedly (10 times) in 95% ethanol in two consecutive containers, before two rinses in the same manner in 100% ethanol. Lastly, slides were dipped repeatedly (10 times) in Clear-Rite-3 (Epridia; Cat#: 6901), placed on a flat surface to dry overnight at room temperature and coverslipped the following day.

Although inclusion of animal-derived products (such as Matrigel and FBS) in therapeutic stem cell differentiation pipelines is not prohibited by regulatory authorities, use of xeno-free components is strongly encouraged. In addition to the increased risk of containing pathogenic contaminants, animal-derived products often suffer from significant lot-to-lot variability. Inconsistency of supplements used in the manufacturing process can have a major impact on protocol reproducibility and potency of the therapeutic product. As depicted in Fig. 1A, the most robust retinal differentiation protocol that we have used to date is based on that first published by Meyer et al. [7], which uses a series of adherent and non-adherent cell culture steps to produce free-floating retinal organoids. This protocol begins with passage of adherent iPSCs from LN521 coated tissue culture plates using ReLeSR into ultra-low adhesion dishes in 75% Essential 8 (E8) media and 25% Neural Induction Media (NIM) (Fig. 1A).

Cultures are fed with 50% E8 and 50% NIM on differentiation day 1, 25% E8 and 75% NIM on differentiation day 2 and 100% NIM on differentiation day 3. On differentiation day 6, NIM is supplemented with BMP4 to bias retinal cell fate specification [8, 19] (Fig. 1A). In the standard non-xenofree protocol on differentiation day 7, resulting embryoid bodies are plated onto Matrigel coated cell culture surfaces to enable attachment, outgrowth of neural progenitor cells, and formation of optic vesicle-like structures (Fig. 1A). Matrigel, which is derived from the mouse chondrosarcoma Engelbreth-Holm-Swarm cell line, is particularly prone to lot-to-lot variability (e.g., lot 11524001 = 11.1 mg/ml total protein and lot 27624003 = 7.9 mg/ml total protein). To identify a suitable Matrigel replacement for embryoid body attachment, on differentiation day 7, embryoid bodies were plated onto dishes coated with 1 of 4 different substrates (Fig. 1B, F, J, & N = Matrigel; Fig. 1C, G, K & O = recombinant human laminin 111, collagen IV, and nidogen-1; Fig. 1D, H, L & P = CELLstart; and Fig. 1E, I, M & Q = recombinant human laminin 111). BMP4 was gradually transitioned out of the NIM over seven days (100% on Day 6, 50% on day 9, 25% on day 12, 12.5% on day 14). On differentiation day 16, NIM was switched to BMP4-free Retinal Differentiation Media (RDM). At differentiation day 30, cultures were mechanically lifted using the back end of a 1 ml pipette tip and cultured in 3DRDM in suspension until differentiation day 100, at which time the media is switched to 3DRDM- (no retinoic acid). No difference in early neural epithelial layer development (Fig. 1B-E) or retinal lamination (Fig. 1F-I) were detected by phase microscopy between any of the substrates used for embryoid body attachment. Likewise, the number of organoids present in each condition was unaffected by plating substrate. The impact of embryoid body plating substrate on retinal organoid maturation was evaluated at differentiation day 180 via phase microscopy (Fig. 1J-M) and immunohistochemical analysis using primary antibodies targeting OTX2 and recoverin (Fig. 1N-Q). Again, we did not detect a significant difference between retinal organoids generated from embryoid bodies plated on any of the xeno-free substrates as compared to those cultured on Matrigel. All organoids developed clearly demarcated inner and outer nuclear layers (Fig. 1N-Q). Furthermore, the outer nuclear layer of all retinal organoids contained OTX2 and recoverin positive photoreceptor cells that extended outer segment-like structures (Fig. 1N'-Q'). Since all substrates showed similar results, CELLstart was used for further experiments because it is a fully-defined xeno-free product, has lot-to-consistency and produced under cGMP.

As indicated above, optic vesicle-like structures are lifted at differentiation day 30, during which time the cell culture media is switched from RDM to 3DRDM (Fig. 2A). Unlike RDM, 3DRDM contains fetal bovine serum (FBS), which is added to improve long term cell survival and retinal organoid maturation (Fig. 2A).

One of the most well characterized xeno-free FBS alternatives that is currently available as a clinical grade reagent is knockout serum replacement (KSR). To determine if KSR could be used in place of FBS, we differentiated iPSCs and used CELLstart in place of Matrigel for embryoid body attachment. At differentiation day 30, half of the organoids were placed in 3DRDM supplemented with FBS and half were placed in 3DRDM supplemented with KSR. Retinal organoids were assessed at differentiation day 160. Those grown in FBS (Fig. 2B-E) and KSR (Fig. 2F-I) had similar morphology with clear lamination and extension of outer segment like projections, the length of which was not significantly different (Supplemental Fig. 1). Immunohistochemical analysis revealed expression of the photoreceptor cell markers OTX2 and Recoverin (Fig. 2D, H) as well as the cone phototransduction protein ARR3 (Fig. 2E, I).

Although cellular reprogramming has been well characterized, the process of converting human dermal fibroblasts to iPSCs following forced expression of OCT4, SOX2, KLF4 and c-MYC is often inefficient [20 –22]. Many strategies designed to enhance reprogramming efficiency have been evaluated, the vast majority of which rely on addition of small molecule inhibitors [23, 24] or modification of the reprogramming factor cocktail [20]. In 2009, Yoshida and colleagues first demonstrated that culturing human dermal fibroblasts under 5% oxygen for 1, 2, or 3 weeks, beginning 1-week post-transgene expression, enhanced reprogramming efficiency by as much as 4-fold [25]. As modifying oxygen tension within the cell culture incubator does not require additional cGMP compliant reagents, we have incorporated hypoxia into our cGMP iPSC generation pipeline [10]. For donors such as the one presented in Fig. 3, reprogramming under reduced oxygen tension is essential. To demonstrate, dermal fibroblasts were passaged 5-days following viral transduction and split at an equal density into two separate LN521 coated cell culture dishes (Fig. 3A & B). At 7 days post-transduction one plate of cells was placed in an incubator maintained at 5% oxygen tension and the other plate was placed in an incubator maintained at 20% oxygen tension. The number of iPSC colonies present two weeks later was determined to be approximately 30-fold greater for cultures maintained at 5% oxygen tension versus those maintained at 20% oxygen tension (Fig. 3C, E, F & G – colony analysis was performed using images collected via the CellX device and an automatic iPSC detection algorithm as we have previously described [13]). Interestingly, iPSC colonies generated under 5% oxygen tension were larger in diameter and as such ready to pick sooner than those generated under 20% oxygen tension (Fig. 3E-G).

To evaluate the impact of oxygen tension on retinal differentiation, iPSCs lines generated at 5% oxygen were split and either maintained at 5% oxygen or cultured under 20% oxygen for 2 passages prior to initiating differentiation. Cells were differentiated using the newly developed xeno-free differentiation protocol described above. As compared to iPSCs maintained at 5% oxygen, cells maintained at 20% oxygen were contained in densely packed colonies, had a high nucleus to cytoplasm ratio, lacked phase bright edges and had increased self-renewal scores as assessed by the TaqMan hPSC ScoreCard analysis (Fig. 4B, G & L). In addition, as determined by RNA abundance, at 7 days post-differentiation there were significantly more embryoid bodies present in iPSC cultures maintained under 20% oxygen tension than those cultured under 5% oxygen tension (Fig. 4C, H & N).

Interestingly, there was no significant difference between lines maintained in 5% oxygen versus 20% oxygen when expression of germ layer specific genes was evaluated, suggesting that oxygen tension alters embryoid formation potential rather than cell fate decision (Fig. 4M). By 30 days of differentiation, optic vesicles were identified in both 5% and 20% oxygen conditions. Interestingly, as with embryoid body formation, there was significantly fewer optic vesicles present in cultures maintained at 5% oxygen than those maintained at 20% oxygen (Fig. 4D & J). Since iPSCs differentiated under 20% oxygen generated significantly more embryoid bodies and in turn optic vesicles that go on to develop into retinal organoids (Fig. 4E, F, J & K, immature retinal organoids containing OTX2 and recoverin positive photoreceptor precursor cells), all subsequent studies were done at this oxygen level.

To test the utility of our amended xeno-free retinal differentiation protocol for derivation of transplantable photoreceptor cells, subretinal injections were performed in the Pde6b-null rat model [15]. The Pde6b-null rat has a rapidly progressive retinal degeneration with near complete loss of photoreceptor cells by 2-months of age [15], which makes it ideal for evaluation of post-transplant donor cell integration (i.e., the lack of host photoreceptor cells at the time of transplantation mitigates concerns pertaining to donor cell antigen transfer [26 –28]). As previously reported in other rodent models of retinal degeneration [29 –34], disease induced activation of retinal microglia can be detected as early as post-natal day 14 in Pde6b-null rats (Supplemental Fig. 2). In addition to expressing IBA1 (which stains for both resident and infiltrating immune cells) and being rounded in shape compared to their normal ramified appearance in the non-diseased state, these cells also express CD68, a lysosomal marker typically present in activated, phagocytosing microglia (Supplemental Fig. 2). By 28-days of age, activated microglia are still present within the degenerating photoreceptor cell layer and subretinal space of the Pde6b-null rat (Supplemental Fig. 2). To determine if the systemic immunosuppression protocol described in the methods section mitigates immune cell infiltration and xenograft rejection, 500,000 iPSC derived retinal cells were injected into the subretinal space of immunosuppressed (N = 8) versus naïve (N = 8) animals. Compared to non-immunosuppressed controls (Supplemental Fig. 3A, C, E, F), animals treated with subretinal injection of cyclosporine had an appreciable decrease in the number of Iba1-postive cells in the neural retina (Supplemental Fig. 3B, D, G, H). Importantly, no influx of CD45 positive infiltrating monocytes or other leukocytes was observed from cyclosporine-treated animals (Supplemental Fig. 3E-H).

Following confirmation of immune suppression, xeno-free iPSC-derived retinal organoids were dissociated, and 2 million cells were injected into the subretinal space of one eye of 22 rats. Immunohistochemical analysis was performed at 3- (N = 11) and 30-days (N = 11) post-injection. To visualize human iPSC-derived cells, the human-specific antibodies human nuclear antigen (HNA), Stem101 (AKA Ku80), Stem121, and human mitochondrial antigen (HMA) were evaluated (Supplemental Fig. 4). As HNA was found to be the most specific with the least amount of background labeling, this antibody was used in subsequent experiments.

At 3-days post-transplantation, HNA-positive cells were present in all animals (N = 11) as a continuous, multi-layered sheet within the subretinal space immediately under the degenerative host inner nuclear layer (Fig. 5A-E), demonstrating successful transplantation. HNA-positive cells expressed the photoreceptor cell marker recoverin (Fig. 5B - expressed in both rods and cones) and the photoreceptor transcription factor OTX2 (Fig. 5C – this marker is also expressed in mature bipolar cells). HNA positive cells did not express the retinal ganglion/amacrine cell marker Calretinin (Fig. 5D) or the bipolar cell marker PKCα (Fig. 5E). There were pigmented RPE cells in the injected area positive for CRALBP (Fig. 5F) that were easily identified by H&E staining (Fig. 5G). As expected, no HNA positive cells were observed in the contralateral buffer-injected eye (Supplemental Fig. 5A-B).

At 30 days post-transplant, host photoreceptor cells were largely absent (Fig. 6A), but recoverin positive photoreceptor cells were present almost exclusively within the transplant region (Fig. 6B-D'). While fewer HNA positive donor cells remained at 30 days post-transplantation, HNA-positive cells were identified in all animals (N = 11). Most HNA-positive donor cells expressed the photoreceptor cell marker recoverin (Fig. 6B). Similarly, s-opsin positive cones were almost exclusively HNA positive (Fig. 6C). Again, no HNA positive cells were detected in the contralateral buffer-injected control eye (Supplemental Fig. 5C-D). Donor HNA positive iPSC-derived photoreceptor cells within the transplant eye remained at the outer limits of the neural retina juxtaposed to PKCα positive bipolar cells (Fig. 6D-D'). Staining for the synaptic marker synaptophysin revealed expression between recoverin positive photoreceptor cells and host bipolar neurons only (Fig. 6D-D'), suggesting that transplanted cells are attempting to form new synaptic connections with the residual host retina. As expected, the number of human iPSC-derived donor cells present in the subretinal space decreased significantly between 3- and 30-days post-transplantation (Fig. 6E). Interestingly, at 3-days following subretinal injection approximately 35–40% of HNA positive donor cells expressed the pan photoreceptor cell marker recoverin (Fig. 6F). By 30-days post-transplant this number increased to approximately 75–80%, indicating that donor photoreceptor cells were selectively preserved (Fig. 6F). Disproportionate survival of transplanted photoreceptor cells likely results from the positive trophic effect afforded by establishment of synaptic connectivity.