HK1191377B - Stem cell-derived retinal pigment epithelial cells - Google Patents

Description

Stem cell-derived retinal pigment epithelial cells

The present application is a divisional application of the chinese patent application having the application number of 200880020748.0, the application date of 2008-27/4, entitled "stem cell-derived retinal pigment epithelial cells".

Technical Field

The present invention relates to methods and systems for producing differentiated retinal pigment epithelial cells (RPEs), and to therapeutic uses of the RPE cells obtained thereby.

List of related art

The following is a list of references deemed relevant for describing the state of the art in the field of the invention.

(1)StraussO.，Theretinalpigmentepitheliuminvisualfunction；Physiol.Rev.85：845-881，2005.

(2)LundRD.etal.，Celltransplantationasatreatmentforretinaldisease；ProgRetinEyeRes20：415-449，2001.

(3)HarutaM.，Embryonicstemcells：potentialsourceforocularrepair；SeminOphthalmol.20(1):17-23，2005.

(4)HarutaM.etal.，Invitroandinvivocharacterizationofpigmentepithelialcellsdifferentiatedfromprimateembryonicstemcells；InvestOphthalmolVisSci45:1020-1024，2004.

(5)AokiH.etal.，EmbryonicstemcellsthatdifferentiateintoRPEcellprecursorsinvitrodevelopintoRPEcellmonolayersinvivo；ExpEyeRes.82(2):265-274，2006.

(6)KlimanskayaI.etal.，Derivationandcomparativeassessmentofretinalpigmentepitheliumfromhumanembryonicstemcellsusingtranscriptomics；CloningStemCells6(3):217-245，2004.

(7)LundRD.etal.，Humanembryonicstemcell-derivedcellsrescuevisualfunctionindystrophicRCSrats；CloningStemCells8(3):189-199，2006.

(8) PCT application publication No. WO06/070370.

Background

Dysfunction, damage and loss of retinal pigment epithelial cells (RPEs) are prominent features of certain ocular diseases and disorders, such as age-related macular degeneration (AMD), hereditary macular degeneration, including Best's disease (early onset form of vitelliform macular dystrophy) and subtypes of Retinitis Pigmentosa (RP). A possible treatment for these diseases is the transplantation of RPE (and photoreceptors) into the retina of those affected by the disease. It is believed that RPE cell supplementation by RPE cell transplantation can delay, halt or reverse degeneration, improve retinal function and prevent blindness from these conditions.

The macula (macula), the central part of the retina, is responsible for fine visual detail and color perception, and is critical to many of our everyday visual tasks such as facial recognition and reading. In widespread retinal degeneration, such as Retinitis Pigmentosa (RP), and in different diseases that more specifically target areas of the macula, such as age-related macular degeneration (AMD) and Best's disease, the macula is often affected as part of the disease process. In many of these diseases, primary dysfunction and failure occurs in the retinal pigment epithelial cells (RPEs), which are located at the base of photoreceptors.

Highly specialized RPE cells play an important role in supporting photoreceptor function: they actively transport nutrients from the choroidal vessels, participate in the recycling of vitamin a, which is essential for chromophores in photoreceptors, and the outer segments of photoreceptors that are taken up and recycled back out as part of the normal renewal process of these cells¹。

In the subtypes Best disease and AMD of RP, failure of the RPE ultimately leads to vision loss and blindness. Replacement of these cells is a possible therapeutic intervention²However, it is difficult to obtain such cells from human donors or embryos. Human embryonic stem cells (hESCs) can serve as a potentially unlimited donor source of RPE cells if the manner in which they are directed to differentiate into functional RPE cells can be elucidated³. Methods for directed differentiation of hESCs into highly enriched cultures of neural precursor cells (NPs) have been described previously (ReubinfBE. et al., Neuriproleitor from neural stem cells; Natbiotechnol 19: 1134-1140, 2001; Itsyson P. et al., Derivaton neuroprosturroso from neural stem cells in presenting cell of noggin; MolCellNeurosci.30(1):24-36, 2005). In addition, hescs have been shown in rodents in vitro and in vivoPotential for generation of retinal cells following transplantation into the subretinal space (Banne. et. al., Retinal incorporation and Differencention of neural PrecursorDesriven from humann Embryonic StemCells; StemCells24(2):246-

The potential of mouse and non-human primate ESCs to differentiate into RPE cells, and to survive and slow retinal degeneration after transplantation has been demonstrated^4,5. Shows spontaneous differentiation of hESCs into RPE, however, the efficiency of the differentiation process is low, requires considerable differentiation time, and only a very low percentage is obtained after 4-8 weeks of differentiation: (<1%) of clusters containing RPE cells. Furthermore, although improved retinal function was observed in RCS rats following subretinal transplantation of these RPE cells, the function of the transplanted cells as authentic mature RPE cells was not exhibited, and this effect could potentially be correlated with non-RPE-specific trophic effects^6,7,9,10。

It has also recently been shown that hescs can be directed to differentiate reproducibly into RPE cells, with the outcome of directed, but not spontaneous, differentiation of hescs into RPE occurring in the presence of Nicotinamide (NA)⁸。

Summary of The Invention

According to a first aspect, the present invention provides the use of a member of the transforming growth factor-beta (TGF- β) superfamily for the preparation of a culture system for promoting the targeting and increased differentiation of human stem cells (hscs) into Retinal Pigment Epithelium (RPE) cells.

According to a second aspect, the present invention provides a method of promoting directed differentiation of hscs into RPE fates, said method comprising:

(a) providing a cell culture comprising hscs; and

(b) culturing cells in the cell culture in a culture system comprising a basal medium supplemented with one or more members of the TGF β superfamily, whereby the hSCs are urged towards directed differentiation to an RPE outcome.

According to a third aspect, there is provided a cell culture comprising RPE cells obtained by directed differentiation of hscs in the presence of one or more members of the TGF β superfamily. Preferably, the RPE cells are terminally differentiated (mature) RPE cells obtained by the methods disclosed herein. As will be shown herein, such RPE cells exhibit several characteristic traits that are different from those obtained when hscs spontaneously differentiate into RPE cells. Preferably, the RPE cells are capable of responding to TGF signaling during their differentiation.

According to a fourth aspect, there is provided a method of transplanting hSC-derived RPE cells obtained by directed differentiation of the hscs into the eye of a subject, the method comprising

(a) Providing a cell culture comprising hscs;

(b) culturing the cell culture in a culture system comprising a basal medium supplemented with one or more members of the TGF β superfamily, whereby the hscs are induced to differentiate into RPE cells;

(c) harvesting RPE cells from the cell culture; and

(d) transplanting the RPE cells into the eye of the subject.

According to a fifth aspect, there is provided a cell culture system comprising transplantable hSC-derived RPE cells obtained by directed differentiation of said hscs. The transplanted RPE cells exhibit one or more parameters indicating that the transplanted cells are functional in the eye of the subject. The functionality of the transplanted RPE cells is demonstrated by their ability to take up shed outer segments of photoreceptors while improving retinal function.

The hscs in the culture system of the methods disclosed herein are differentiating hscs, i.e., a population of hscs in a substantially undifferentiated state, or wherein at least a portion of the cells have been induced to undergo an initial stage of committed differentiation, and sometimes a majority of the cells have been induced to undergo an initial stage of committed differentiation. According to one embodiment, the initial stage of differentiation is achieved by prior exposure of the cells to NA, whereas the initial stage of differentiation will also occur when undifferentiated cells are co-exposed to NA and one or more members of the TGF β superfamily. Without being bound by theory, it is hypothesized that prior exposure to NA (prior to incubation with one or more members of the TGF β superfamily) causes the cells to tend to differentiate committed (as opposed to spontaneous) to RPE cells with a particular RPE morphology, as will be discussed further below.

According to a preferred embodiment, the hscs are human embryonic stem cells (hescs).

According to one embodiment, culturing the cells in a medium comprising one or more members of the TGF β superfamily is at least two days after the hscs have initial differentiation, committed differentiation, preferably committed differentiation by NA.

According to a fifth aspect, there is provided a method of treating or preventing a retinal disease or disorder (which includes dysfunction, damage and/or loss of retinal pigment epithelium) in a subject, the method comprising intraocular transplantation of hSC-derived RPE cells obtained by inducing hSC towards directed differentiation into the eye of the subject. Transplantable RPE cells are preferably obtained by the methods disclosed herein.

Brief description of the drawings

In order to understand the invention and to show how it may be carried out in practice, a preferred embodiment will now be described, by way of non-limiting example only, with reference to the accompanying drawings, in which:

FIGS. 1A-1E: real-time PCR, analyzing the expression of RPE markers in the presence of NA. Differentiation of hescs was induced by culturing them as free floating clusters (clusters). At 6 weeks of differentiation, the expression levels of the RPE markers MiTF-a (fig. 1A) and RPE65 (fig. 1B) were significantly increased in the presence of NA. Real-time PCR analysis at successive time points revealed that the expression levels of MiTF-a (fig. 1C) and RPE65 (fig. 1D) gradually increased over time in the presence of NA. Expression of other transcripts including RPE markers for wilting protein (Bestrophin), CRALBP and Mertk was demonstrated by RT-PCR analysis of plated pigmented clusters (fig. 1E). +/-indicates the presence or absence of reverse transcriptase, respectively.

FIGS. 2A-2F: the RPE differentiation-inducing effect of NA is not dependent on the specific medium composition. Dark field micrographs of hESC clusters differentiated for 12 weeks in KO medium (fig. 2A), or in Neurobasal medium supplemented with N2, replaced with DMEM/F12(NN medium) supplemented with B27 after 1 week (fig. 2C). In both media, NA increased differentiation towards pigmented cells (fig. 2B, D), while the size of the differentiated hESC clusters and their total number were smaller with NN media (white arrows mark the pigmented area within the cluster under differentiation). At the RNA level, the NA addition increased the expression level of MiTF-A and RPE65 in both media (FIGS. 2E and F, respectively).

FIGS. 3A-3L: the melanin-expressing cells within the free floating clusters of hescs are putative RPE cells. Dark field micrographs of free floating clusters of hescs in differentiation with defined areas highly enriched in pigmented cells (fig. 3A). Fluorescence (fig. 3B) and phase contrast (fig. 3C) images of pigmented cells immunoreactive with anti-Otx 2 and MiTF after dissociation and plating. Dark field micrographs of clusters in differentiation after plating showing restricted pigmented areas are shown (fig. 3D). Phase contrast images of cells with morphological features typical of RPE cells within the pigmented region (fig. 3E). Indirect immunofluorescent staining showed that these cells expressed markers for RPE cells, including MiTF (fig. 3F), ZO-1 (fig. 3G), wilting (fig. 3H), RPE65 (fig. 3I) and CRALBP (fig. 3J). After dissociation, low density plating and culture, pigmented cells lost pigmentation and acquired a fiber-like morphology (phase contrast image) (fig. 3K). After further prolonged culture and proliferation into high density cultures, the cells again acquired morphology and pigmentation characteristic of RPE cells (fig. 3L).

FIGS. 4A-4E: activin a induces RPE differentiation. Human ESCs were allowed to differentiate as free floating clusters for 6 weeks in the presence or absence of activin a, which was added after the first week of differentiation. Dark field micrographs of the clusters showed that activin a significantly increased the percentage of clusters that included pigmented cells (fig. 4A, B) (white arrows mark pigmented regions within clusters in differentiation and the boundaries of pigmented regions within some clusters are marked by dashed lines). In the presence of activin A, the boundaries of the pigmented regions are more clearly separated from the surrounding non-pigmented regions within the clusters. Furthermore, the pigmented cells were darker in the presence of activin a (fig. 4B). At the RNA level, real-time PCR analysis showed that the expression of RPE65 (fig. 4D) and wilting (fig. 4E) was significantly increased in the presence of activin a. The expression of MiTF-a was not altered by activin a treatment (fig. 4C).

FIGS. 5A-5G: BMPs and TGF β 3 play a role in RPE differentiation. Human ESCs were induced to differentiate as free floating clusters for 6 weeks. Spontaneous differentiation into pigmented cells was occasionally observed (FIG. 5A), but was significantly increased when the medium was supplemented with NA (FIG. 5B, left-most and left-most dark field images; white arrows mark pigmented areas within clusters in differentiation). In the absence (FIG. 5D) and presence (FIG. 5C) of NA, media supplementation with noggin blocked differentiation into pigmented cells. At the RNA level, real-time PCR analysis showed that noggin reduced the expression level of MiTF-a both in the presence and absence of NA (fig. 5E). When TGF β 3 was added to the medium during differentiation of hESC clusters in the presence of NA, it significantly increased the expression level of MiTF-a (fig. 5F) but not RPE65 (fig. 5G).

FIGS. 6A-6J: survival and integration of hESC-derived RPE cells transplanted in rat eyes. Pigmented cells can be readily identified in vivo in the eyes of albino rats following intraocular transplantation of hESC-derived RPE cells (fig. 6A, 6B). After enucleation of the eye (fig. 6B) and removal of the cornea and lens, the main grafts and other discrete pigmented spots were visible (fig. 6C). In tissue sections, grafts comprising darkened pigmented cells that also co-express GFP could be identified (FIGS. 6D-6G), demonstrating the fact that the cells are hESC derived. Transplanted cells can be found in the intravitreal area, between the retina and the lens (fig. 6H), in the retina (occasionally extending into the vitreous along the injected tract) (fig. 6I), and in the subretinal space (fig. 6I, 6O, 6P). Transplanted hESC-derived RPE cells (pigmented cells marked with arrows) integrated into the RPE layer of albino rats (fig. 6J). No pigmented cells were observed in the RPE layer of the control non-transplanted eye. Within the grafts, immunostaining with ZO-1 (FIGS. 6K-6N) showed tight junctions between the transplanted GFP + hESC-derived cells. This linkage is characteristic of RPE cells. After transplantation into the subretinal space of RCS rats with RPE dysfunction and retinal degeneration, relative preservation of the photoreceptor layer can be seen close to the graft compared to the area far from the graft (marked by the arrow) (FIG. 6O; the area within the rectangle is marked by an asterisk, enlarged in FIG. 6P). Note that a number of transplanted hESC-derived RPE cells had polygonal shape and cobblestone-like appearance (fig. 6P) (asterisks). In all cases shown here, RPE cells were derived from hescs in the absence of activin a.

FIG. 7: electronic retinal imaging recordings show that transplantation of hESC-derived RPE cells provides a rescue of retinal function in the eyes of dystrophic RCS rats. Full field ERG responses were higher in RCS rat eyes after transplantation of RPE cells from hescs compared to companion non-transplanted control eyes (n-11 rats). RPE cells used for these experiments were derived without addition of activin a to the culture medium. The b-wave amplitude of the dark adapted mixed cone-rod response to four stimuli of increasing intensity is shown.

FIGS. 8A-8I: the morphology of the effect of NA in inducing pigmented cell development from hescs and analysis of marker expression are shown. Dark field micrographs show the progressive appearance of pigmented cells during 4 weeks (fig. 8A, 8B), 6 weeks (fig. 8C, 8D), and 8 weeks (fig. 8E, 8F) of hESC-derived cluster culture in the presence (fig. 8A, 8C, and 8E) or absence (fig. 8B, 8D, or 8F) of NA (white arrows mark pigmented regions within clusters in differentiation). Histogram representation of the percentage of clusters containing pigmented regions at different time points during culture in medium supplemented with NA (thick line bars) and in control cultures (thin line bars) (FIG. 8G). Histogram representation of the percentage of pigmented cells (fig. 8H) and cells immunoreactive to early RPE markers against MiTF (fig. 8I) during 8 weeks of culture with NA supplementation. Scale post: (A)200 mm; p < 0.05; p < 0.001.

FIGS. 9A-9S: real-time PCR, immunostaining, and flow cytometry analysis of RPE developmental progression over time in clusters in hESC differentiation are shown. (FIGS. 9A-9L) real-time PCR, which analyzed the timing of expression of key genes in RPE development in clusters cultured in the presence (thick line bars) or absence (thin line bars) of NA. Stepwise expression of the following markers was analyzed at successive time points during the 8-week differentiation of hESC-derived clusters: the hESC-specific marker Oct4 (fig. 9A); early neural markers Otx2 (fig. 9B), Musashi (fig. 9C), and Pax6 (fig. 9D); retinal progenitor markers Six3 (fig. 9E), Rx1 (fig. 9F) and Chx10 (fig. 9G); RPE markers MiTF-a (fig. 9H), RPE65 (fig. 9I) and wilsonin (fig. 9J); photoreceptor progenitor marker Crx (FIG. 9K); melanocyte development marker Sox10 (bar with bar, fig. 9L) (M51 melanoma cell line was used as control). FACS analysis of stepwise expression of the hESC-specific marker TRA-1-60 (fig. 9M) and the neural progenitor marker PSA-NCAM (fig. 9O) was demonstrated in cluster differentiation for 8 weeks with (bold line) or in the absence (fine line bars) of NA. Within clusters differentiated for 2 and 4 weeks in the presence of NA, early neural markers were expressed: indirect immunofluorescence analysis of the percentage of cells of PSA-NCAM (bold bars column), nestin (horizontal bars column), Musashi (thin lines column), Pax6 (vertical bars column) (fig. 9N). Immunofluorescence images showing cells expressing these markers, PSA-NCAM (fig. 9P), nestin (fig. 9Q), musashi (fig. 9R), Pax6 (fig. 9S).

FIGS. 10A-10J: analysis of morphology, marker expression and function, showed that pigment expressing cells within free-floating clusters of hescs were putative RPE cells. Phalloidin staining revealed the distribution of F-actin in hESC-derived pigmented progeny that exhibited characteristics of RPE (fig. 10A); after dissociation, low density plating and culture, pigmented cells lost pigmentation and acquired a fiber-like morphology (phase contrast image, 1 week culture) (fig. 10B). After further prolonged culture and proliferation into high density cultures, the cells again acquired morphology and pigmentation characteristic of RPE cells (1.5 months of culture) (fig. 10C). Electron microscopy analysis of hESC-derived RPE cells, showed features that are characteristic of RPE: microvilli (fig. 10D), basement membrane (fig. 10E), melanin granules (fig. 10D), and tight junctions (fig. 10F). Phase contrast (FIG. 10G) and fluorescence images (FIGS. 10H-J) show phagocytosis (white arrows) of green fluorescent latex beads of hESC-derived pigmented cells; the cell membrane was stained with the red fluorescent dye PKH (grey). The three confocal fluorescence images represent sequential Z-axis slices (fig. 10H-J).

FIGS. 11A-11P: analysis of morphology and gene expression, showed that factors from the TGF β family promote differentiation towards RPE fates. Dark field micrographs of hESC-derived clusters that differentiated for 4 weeks showed the appearance of pigmented cells at this early stage in the presence of activin (fig. 11A), and an increase in the number of pigmented clusters differentiated in the presence of activin a and NA (fig. 11C), as opposed to NA alone (fig. 11B). Similar to activin a, supplementation with TGF β 1 also increased the appearance of pigmented clusters (fig. 11D). In contrast, the use of the inhibitor of the activin signaling pathway, SB431542, together with activin a and NA, reduced the efficacy of activin a for the appearance of chromatogenic clustering (fig. 11E). The development of pigmented clusters was also eliminated by culturing the cells in the presence of FGF β and NA (fig. 11F). Expression of activin receptor and activin a transcripts was demonstrated by RT-PCR analysis of 2-week-old clusters cultured in the presence or absence of NA, and undifferentiated hESCs as controls (fig. 11G). Histogram analysis of the percentage of clusters containing pigmented regions in the presence of NA, NA + ActA, NA + SB431542, NA + ActA + SB431542, NA + TGF β 1 after 4 weeks of culture (FIG. 11H). Histogram analysis of the percentage of pigmented cells after 4 weeks of culture with NA (bold bars) or activin a and NA (diagonal striped bars) (fig. 11I). Histogram analysis of the percentage of pigmented cells at different concentrations of activin a (fig. 11J), which is optimal for RPE induction of 140ng/ml, and the expression levels of the transcripts of the RPE markers, wilting (fig. 11K) and RPE65 (fig. 11L). Real-time PCR time-course analysis of the effect of activin a on expression levels of retinal and RPE genes, wilting (fig. 11M), MiTF-total (fig. 11N), Rxl (fig. 11O), and Chx10 (fig. 11P) in hESC differentiation with (diagonal striped bars) or without (bold bars) activin a addition, in the presence of NA. P <0.005. (white arrows mark pigmented regions within clusters in differentiation).

FIGS. 12A-12E: RPE cells from hescs treated with NA and activin a survived subretinal transplantation in the eyes of dystrophic RCS rats. The clustering of pigmented cells can be performed using a fundus imaging system (FIGS. 12A-12C); fundus (fig. 12A) and no red (fig. 12B) photographs were readily identified in vivo in the eyes of RCS rats, showing the sub-retinal location of the graft (noting that retinal blood vessels pass through the pigmented area). hESC-derived GFP expressing cells can be seen to emit fluorescence upon fluorescence excitation and use of an emission filter (fig. 12C). In the preparation of cups imaged ex vivo (exvivo) in a fluorescence microscope (fig. 12D-12E), large clusters of GFP positive cells under the retina (fig. 12D) and multiple discrete smaller clusters (fig. 12E) were visible.

FIGS. 13A-13F: histological appearance of subretinal hESC-derived activin a-treated RPE cell grafts in RCS rat eyes. Hematoxylin and eosin (fig. 13A and 1B) stained tissue sections showed transplanted hESC-derived pigmented cells in subretinal and occasionally intraretinal locations, appearing in clusters or as isolated cells (arrows). Immunostaining with GFP (FIGS. 13C-13F) confirmed that these cells were indeed hESC-derived. The grafts were often quite large and scattered (FIGS. 13C, 13E), and the co-expression of GFP in pigmented cells was evident (FIGS. 13D, 13F). Note that GFP-positive pigmented cells integrated within the RPE layer of the host (fig. 13D, arrow).

FIGS. 14A-14O: the transplanted hESC-derived pigmented cells express markers of mature RPE. Immunostaining revealed that a large number of transplanted cells within the graft expressed proteins characteristic of mature RPE cells, including RPE-specific markers RPE65 (fig. 14A-14E) and wilting protein (fig. 14F-14J) and the tight junction marker ZO-1 (fig. 14K-14O). FIGS. 14A, 14F and 14K show low magnification fluorescence images of grafts co-expressing GFP and associated markers. High magnification confocal images at each row show pigment (by Nomarski optics) at the single cell level as well as co-expression of GFP and different markers. These series confirmed that these cells were indeed hESC-derived, and that they expressed markers of mature RPE in vivo. In FIG. 14M, it is noted that the host RPE stained ZO-1 (dashed arrows), while it was GFP negative (corresponding regions dark) in FIGS. 14N, 14O, in contrast to ZO-1 positive hESC-derived cells (full arrows in FIG. 14M), which did co-express GFP (FIGS. 14N, 14O).

FIGS. 15A-15C: transplanted hESC-derived, activin a-treated RPE cells provided a functional rescue in the RCS rat retinal degeneration model. The full-field ERG responses recorded at 8 weeks of age were higher in the RCS rat eyes after transplantation of activator protein-treated RPE cells from hescs compared to the congenic non-transplanted control eyes, and compared to the subretinal injected eyes of medium alone. A series of lifts under dark adaptation conditions are shown in the transplanted eye (FIG. 15A) and its cognate control eye (FIG. 15B)FIG. 15C shows a significant difference in mean amplitude between transplanted eyes and control eyes of different groups (- ◆ -injected eyes (n ═ 13), - ■ -uninjected eyes (n ═ 13), - ● -uninjected eyes of medium (n ═ 5-Medium injected eyes (n ═ 5)). As shown, retinal function tended to be better preserved following transplantation of activin a-treated RPE cells (shown here) than the rescue effect achieved following transplantation of activin a-free derived RPE cells (fig. 7).

FIGS. 16A-16D: transplanted hESC-derived, activin a-treated RPE cells provided rescue of structures in the RCS rat retinal degeneration model. High resolution microscopy images of sections stained with hematoxylin and eosin were used to examine and quantify the effect of transplanted hESC-derived activator protein-treated RPE cells on degenerated host retina. Relative preservation of the outer nuclear (photoreceptor) layer (ONL) and the inner and outer photoreceptor segments (IS + OS) was observed near the subretinal RPE graft as compared to regions distant from the graft (two examples are shown in fig. 16A, 16B). The inset in fig. 16A demonstrates this difference (the rescued retina is shown with a relatively thick ONL in the right inset near the implant; severe thinning of the ONL is seen in the left inset away from the implant). Overall retinal thickness (fig. 16C) and ONL and IS + OS thickness (fig. 16D) increased significantly near the hESC-derived RPE grafts compared to regions distant from the graft (grey bars) (black bars, mean ± SEM, n ═ 7). This type of structural rescue was only observed at the subretinal and deep intraretinal grafts, but not when the graft was only intravitreal (not shown). For details of the quantitative technique, please see methods.

FIGS. 17A-17E: transplanted hESC-derived activin a-treated RPE cells take up rhodopsin in vivo. Confocal images of subretinal transplanted RPE cells showed co-localization of pigment, GFP, RPE65 and rhodopsin within the same single cell. Native RPE cells from RCS rats expressed RPE65 (fig. 17C, arrow), but did not express GFP (fig. 17D, arrow) and contained minimal rhodopsin (fig. 17B, 17E).

Detailed description of the invention

The present disclosure provides the use of one or more members of the transforming growth factor-beta (TGF β) superfamily for the preparation of a culture system for promoting differentiation of human stem cells (hscs), preferably human embryonic stem cells (hescs), into Retinal Pigment Epithelium (RPE) cells. It is noted that, in addition to the specific uses discussed in detail herein, also encompassed within the present disclosure are RPE cells obtained by directed differentiation of hscs in the presence of one or more TGF β superfamily; and methods of promoting directed differentiation of hscs into RPE fates, as well as methods of growing and maintaining such hSC-derived RPE cells, and methods of using such hSC-derived RPE cells. According to certain preferred embodiments, RPE cells obtained according to the teachings herein are mature (in other words, terminally differentiated) and functional RPE cells, as discussed and illustrated further below.

The present disclosure broadly relates to the use of one or more members of the TGF β superfamily of growth factors to promote/induce/enhance directed differentiation of hscs into RPE cells, preferably mature RPE cells.

In the following specification and claims, various terms will be used from time to time, the meanings of which should be construed in accordance with the teachings herein, as follows:

glossary

"transforming growth factor-beta (TGF-beta) superfamily growth factors," as used herein, refers to any member of the TGF-beta superfamily of growth factors, such as transforming growth factor-beta proteins, including TGF-beta 1, TGF-beta 2, and TGF-beta 3 subtypes, and homologous ligands, including activins (e.g., activin A, activin B, and activin AB), nodal, anti-mullerian hormone (AMH), certain Bone Morphogenetic Proteins (BMPs), such as BMP2, BMP3, BMP4, BMP5, BMP6, and BMP7, and Growth and Differentiation Factors (GDFs).

"human stem cells" or "hscs," as used herein, refer to cells of human origin that are capable of differentiating into other cell types with specific specialized functions under suitable conditions, and of self-renewal and maintenance in an undifferentiated pluripotent state under other suitable conditions, as described in detail below.

As used herein, "cell" refers to a single cell as well as a population of cells (i.e., more than one cell). The population may be a pure population comprising one cell type. Alternatively, the population may comprise more than one cell type. The hSC cells are preferably hematopoietic or mesenchymal stem cells obtained from bone marrow tissue of an individual of any age after birth, or from umbilical cord blood or tissue of a newborn individual, neural stem cells obtained from a fetus or cadaveric brain of any age after birth. The term cell may denote a single cell or a cluster of cells.

As used herein, "embryonic stem cell" and "pluripotent embryonic stem cell" refer to cells that can give rise to any differentiated cell type in an embryo or adult, including germ cells (sperm and ovum).

As used herein, "cell culture" or "cultured cells" refers to cells or tissues that are cultured, grown, or grown in an artificial in vitro environment.

As used herein, "undifferentiated pluripotent hSC" or "pluripotent hSC" refers to a precursor cell of human origin that has the ability to form any adult human cell. Such cells are true cell lines because they: (i) can proliferate in an undifferentiated state in vitro in a wide range; and (ii) derivatives capable of differentiating into all three embryonic germ layers (endoderm, mesoderm and ectoderm) even after prolonged culture. Other pluripotent hscs include, but are not limited to, pluripotent adult progenitor cells (MAPs), induced pluripotent stem cells (iPS cells), and amniotic fluid stem cells.

As used herein, "undifferentiated" refers to cells cultured when a substantial proportion (at least 20% and possibly more than 50% or 80%) of the cells and their derivatives in the population exhibit markers and morphological characteristics characteristic of undifferentiated cells, distinguishing them from differentiated cells of embryonic or adult origin. Cells are considered to proliferate in an undifferentiated state when they undergo at least 1 population doubling during a culture period of at least 3 weeks, while retaining at least about 50% after said culture period, or the same proportion of cells carry a marker or morphological feature characteristic of undifferentiated cells.

As used herein, "cell suspension" or "free-floating cells" refers to a culture of cells in which a majority of the cells float freely in a matrix, typically in a culture medium (system), as single cells, or as a cell population and/or as a cell aggregate. In other words, the cells survive and proliferate in the culture medium without adhering to the substrate.

As used herein, "culture system" refers to a culture system suitable for the proliferation of SC. The term denotes a combination of ingredients, including at a minimum a basal medium (a cell culture medium typically comprising a defined base solution, which includes salts, sugars, and amino acids) and one or more members of the transforming growth factor-beta (TGF β) superfamily of growth factors. The culture system according to the invention may further comprise other components, such as, without limitation, serum or serum replacement, culture (nutrient) matrix and other exogenously added factors, which together provide suitable conditions for supporting the growth of the SC, as well as other components typically used in cell culture systems. The above ingredients may be collectively classified as soluble ingredients. However, in the context of the present invention, the ingredients may also be associated with a carrier, i.e. an insoluble ingredient. The association may be by chemical or physical attachment/binding. For example, the components may be immobilized on a matrix (e.g., extracellular matrix), presented by cells added to the system or incorporated on a biodegradable material. Further, the components may be released from a carrier, which may be a cell or a vesicle encapsulating or embedding the components. Thus, in the text that follows, the components that supplement the basal medium to form the culture system include soluble and insoluble components.

As used herein, "differentiation" refers to the process of switching cell states from one cell type to another, and more particularly in the context of the present disclosure, to the process of human stem cells obtaining a cell type of Retinal Pigment Epithelial (RPE) cells that has at least one characteristic feature indicating that the RPE cells are mature (terminally differentiated) cells. As used herein, the term "cell type" refers to a unique morphological or functional form of a cell.

As used herein, "differentiating hscs" refers to undifferentiated hscs that are capable of differentiating into a predetermined fate in an incremental, directed manner under appropriate conditions; the term also refers to a population of hscs, wherein at least a portion thereof has been induced to undergo at least initial differentiation, i.e., committed differentiation, or a combination thereof.

"cause", "increase", "promote" or "cause.

"differentiation inducer" or "differentiation promoter", "differentiation initiating agent" or "differentiation promoting factor" are used interchangeably herein to mean any agent capable of initiating, augmenting, promoting differentiation of pluripotent SC into somatic cells or directing differentiation of pluripotent SC into somatic cells, preferably into RPE cells.

"retinal pigment epithelial cells," "RPE" may be used interchangeably, where the context permits, to refer to cells of a cell type that is functionally similar to native RPE cells that form the pigmented cell layer of the retina (e.g., they exhibit functional activity similar to native RPE cells when transplanted into the eye). Thus, the terms "retinal pigment epithelial cells", "RPE cells" or "RPE" may be used to refer to native RPE cells of the pigmented layer of the retina and RPE cells that are directly differentiated from hscs according to the present disclosure.

The term "hSC-derived RPE cells" is used herein to denote RPE cells obtained by directed differentiation from hscs. According to a preferred embodiment, the hSC-derived RPE cells are mature (terminally differentiated) and functional RPE cells, as demonstrated by the parameters defined below. The term "directed differentiation" is used interchangeably with the term "RPE-induced differentiation" and is understood to refer to the process of manipulating hscs under culture conditions that induce/promote differentiation into RPE cell types only.

"functional RPE cells" as used herein means cells obtained by directed differentiation of hscs in the presence of one or more members of the TGF β superfamily, said RPE cells exhibiting at least one of the following characteristics:

during differentiation, the cultured cells respond to TGF signaling;

-the RPE cells are mature, terminally differentiated cells as exhibited by expression of markers indicative of terminal differentiation, such as wilting or RPE65, simultaneously or alternatively, by their lack of potential to proliferate in vivo.

After transplantation (i.e. in situ), RPE cells exhibit a trophic potency to support photoreceptors adjacent to RPE cells;

-further, in situ, the RPE cells are capable of functioning as phagocytosis of shed photoreceptor outer fragments as part of the normal renewal process of these photoreceptors.

Thus, RPE cells according to the present invention are particularly suited for regeneration of host RPE, thereby providing improved vision after transplantation thereof into the eye of a subject.

"similar," when used in the context of differentiating RPE cells, means that the differentiated RPE cells share one or more unique morphological or functional characteristics with native RPE cells. For example, sufficient similarity can be obtained, for example, by determining that the differentiated cells express one or more markers of naturally occurring RPE cells, such as MiTF, ZO-1, wilting protein, RPE65, Otx2, Mertk, and CRALBP; or the cells exhibit one or more physical morphological characteristics of the RPE cells, such as typical F-actin distribution within the cells, pigmentation of pigmented microparticles, polygonal (e.g., hexagonal) shape, cobblestone-like appearance, and ultrastructural characteristics of the RPE exhibited by electron microscopy. In addition, any of the above listed functions may be included, for example, supporting the trophic effects of photoreceptors adjacent to RPE cells; the functionality of phagocytosis of shed photoreceptor outer fragments carrying rhodopsin or lacking the potential for proliferation in vivo.

As used herein, "large scale" with respect to cell culture and expansion refers to the production of RPE cells under conditions that allow at least doubling of cells in cell culture after 4 weeks, with the cell population after 4 weeks consisting essentially of RPE cells.

As used herein, "cell marker" refers to any phenotypic characteristic of a cell that can be used to characterize the cell or to distinguish the cell from other cell types. The marker may be a protein (including secreted, cell surface or internal proteins; synthesized or taken up by the cell); a nucleic acid (e.g., an mRNA, or an enzymatically active nucleic acid molecule) or a polysaccharide. Included are determinants of any of these cellular components, which can be detected by antibodies, lectins, probes, or nucleic acid amplification reactions specific for markers of the cell type of interest. The markers may also be identified by biochemical or enzymatic analysis or biological reaction depending on the function of the gene product. Associated with each marker is a gene encoding a transcript, and the events that lead to the expression of the marker. A marker is said to be preferentially expressed in an undifferentiated or differentiated cell population if its expression is at a level that is at least 50% (for total gene product measured in antibody or PCR assays) or more frequently 30% (for positive cells in the population) higher than an acceptable control such as actin or glyceraldehyde 3-phosphate dehydrogenase (GAPDH). Markers that express more or more frequently by a factor of 2, 10, 100 or 10,000 are increasingly more preferred.

The current disclosure utilizes RPE-derived hscs due to the induced and directed differentiation of hscs in the presence of a unique culture system applied to suspended hscs.

Non-limiting examples of hscs are neural stem cells obtained from a fetus or from any age after birth or from a cadaver, hematopoietic stem cells obtained from bone marrow tissue of a human subject at any age after birth or from umbilical cord blood of a newborn subject, mesenchymal stem cells, amniotic fluid stem cells, induced pluripotent stem cells, or stem cells obtained from gonads of a human subject at any age after birth. Preferred human stem cells according to the invention are human embryonic stem cells (hescs).

The hscs can be obtained using well-known cell culture methods.

Commercially available hscs may also be used according to the present invention. HSCs can be purchased from NIH human embryonic stem cell registry. Non-limiting examples of commercially available embryonic stem cell lines are BG01, BG02, BG03, BG04, CY12, CY30, CY92, CY10, TE03 and TE 32.

The potential applications of hescs and cells derived from them are broad, including drug development and testing, production of cells, tissues and organs for transplantation, production of biomolecules, testing compounds for toxicity and/or teratogenicity, high throughput screening of molecules for toxicity, regeneration, protection or other efficacy, and facilitating development and other biological processes. For example, diseases currently contemplated to be treatable by therapeutic transplantation of hescs or hESC-derived cells include parkinson's disease, cardiac infarction, juvenile onset diabetes, and leukemia [ geohartj. science 282: 1061-1062, 1998; RossantandNagy, nature biotech.17: 23-24, 1999].

However, there are significant obstacles to the practical development of hescs. Two such obstacles include: maintaining hescs in an undifferentiated, pluripotent state without spontaneous differentiation; and directing differentiation of hescs into specific types of somatic cells. Several culture systems have been described for the maintenance and proliferation of stem cells, in particular hescs, in an undifferentiated state⁸。

Because of the potential of differentiated cells derived from stem cells in myriad therapeutic applications, it is of great interest to direct differentiation of stem cells in culture or to force differentiation of stem cells in culture towards a particular somatic fate.

In certain ocular diseases and disorders, such as those of the retina and macula (macula), failure of RPE cells ultimately leads to vision loss and even blindness. Transplantation of RPE cells to replace and support failing host RPE has been suggested as a possible therapeutic intervention, but obtaining such cells from human donors or embryos is difficult. Hscs can therefore serve as a potentially unlimited donor source of RPE cells if the way in which hSC cells are committed to differentiate into functional RPE cells can be elucidated.

It has now surprisingly been found that contacting hscs with a large number of members of the TGF β superfamily of growth factors strongly promotes differentiation of hscs towards the outcome of RPE. In other words, these growth factors have an inducing effect on hscs. Thus, the use of members of the transforming growth factor-beta (TGF- β) superfamily for the preparation of culture systems for inducing differentiation of human stem cells (hscs) into Retinal Pigment Epithelium (RPE) cells is envisaged.

While many members of the TGF-beta superfamily of growth factors are known (some non-limiting examples are listed above), according to preferred embodiments, the members of the TGF-beta superfamily are preferably TGF-beta 1, TGF-beta 3 growth factor or activin A or combinations thereof.

It was previously found that Nicotinamide (NA) in cell culture has an inhibitory effect on the differentiation of stem cells into extra-embryonic cells, and furtherNA promotes somatic differentiation towards the nerve, and further towards RPE cell-like fate⁸NA, also known as "nicotinamide", is an amide-derived form of vitamin B3 (niacin), which is believed to preserve and improve β cell function₆H₆N₂And O. NA is essential for growth and conversion of food to energy, and it has been used in arthritis treatment and diabetes treatment and prevention.

In the context of the present disclosure, the term NA also denotes a derivative of NA.

The term "derivative of Nicotinamide (NA)" as used herein represents a compound that is a chemically modified derivative of natural NA. Chemical modifications may include, for example, substitutions on the pyridine ring of the underlying NA structure (via a carbon or nitrogen member of the ring), through a nitrogen or oxygen atom of the amide moiety, and deletions or substitutions of groups, for example, to form thiobenzamide analogs of the NA, all as understood by those skilled in organic chemistry. Derivatives in the context of the present invention also include nucleoside derivatives of NA (e.g. nicotinamide adenine). Various derivatives of NA are described, some of which are also involved in the inhibitory activity of the PDE4 enzyme (WO 03/068233; WO 02/060875; GB2327675A) or as VEGF receptor tyrosine kinase inhibitors (WO 01/55114). For example, processes for the preparation of 4-aryl-nicotinamide derivatives (WO 05/014549).

In connection with the above, it has now surprisingly been found that when hscs differentiate in the presence of NA, their properties are altered, so that they acquire the ability to respond to the induced effects of one or more members of the TGF β superfamily, which direct their differentiation towards RPE fates, preferably mature and functional RPE cells. Thus, when the hscs are subsequently exposed to one or more members of the TGF β superfamily of growth factors, the RPE differentiation-inducing effects of NA may be significantly increased in combination with the pre-exposure of the cells to NA in culture.

Thus, according to one embodiment, the method comprises treating the cells with one or more members of the TGF β superfamily of growth factors, in combination with prior exposure to NA. Combinations for the preparation of culture systems comprising TGF β and NA; and for the preparation of a culture system comprising only one or more members of the TGF β superfamily for inducing/promoting differentiation and/or further differentiation of hscs that have been exposed to NA. Without being limited by theory, it is believed that NA acts as a differentiation inducer/promoter and, similarly, one or more members of the TGF β superfamily act as RPE differentiation promoting factors. Furthermore, without being limited by theory, it is believed that prior exposure of hscs to NA provides cells with properties that allow them to respond to the RPE differentiation promoting effects of one or more members of the TGF β superfamily.

Thus, according to an embodiment of the invention, the hscs are first cultured in a culture system comprising a basal medium supplemented with NA for at least several hours, preferably for at least one day, more preferably for at least 2 days, before culturing the cells in the cell culture in a basal medium (the same or different) supplemented with one or more members of the TGF β superfamily.

According to another embodiment, the undifferentiated hscs are cultured in a culture system comprising a basal medium supplemented with NA and one or more members of the TGF β superfamily.

It is noted that various basal media are known in the art for cell cultures and preferably for SC cultures. A non-limiting list of basal media that can be used according to the present disclosure includes Neurobasal^TM(CAT#21103-049，Gibco1998/1999)、KO-DMEM(CAT#10829-018，Gibco1998/1999)、DMEM(CAT#41965-039，Gibco2004)、DMEM/F12(CAT#21331-020，Gibco2004)、Cellgro^TMStem cell growth medium (CAT #2001, CellGenix2005) or X-Vivo^TM(CAT#04-380Q，LONZA2007)。

The present disclosure also provides a method of inducing directed differentiation of hscs into RPE fates, the method comprising:

(a) providing a cell culture comprising hscs;

(b) culturing cells in the cell culture in a culture system comprising a basal medium supplemented with one or more members of the TGF β superfamily, thereby promoting directed differentiation of hscs into RPE fates.

Differentiation may occur within free floating clusters or attached cultures of hscs. Somatic differentiation in adherent culture is described [ U.S. patent No.7,112,437 ]. Such an attached culture may thus serve as a basis for inducing RPE differentiation by a culture system supplemented with at least one or more members of the TGF β superfamily of growth factors.

The cells in the cell culture may be a population of undifferentiated hscs, or a population of cells in which at least a portion of the hscs have begun to differentiate. The initial differentiation is directed differentiation. Thus, in the context of the present disclosure, the cells provided in the methods are sometimes referred to as differentiating cells.

As already indicated above, the basal medium can be supplemented by introducing soluble and insoluble components therein. For supplementation with one or more members of the TGF β superfamily of growth factors, the members may be present in soluble form, or immobilized or associated with a matrix or cells added to the culture system, or the components may be bound or complexed to other substances. The member may also be secreted into the culture system from the cells present in the latter.

The hscs may be provided in an undifferentiated state, and after exposure to differentiation promoting factors (differentiation initiating agents) such as NA. Undifferentiated hscs can be obtained from various culture systems in which the hscs can be maintained in an undifferentiated pluripotent state. For example, the cells may be cultured in feeder-free (feeder-free) attachment or suspension systems (WO06/070370) or on feeder cells. Commonly used feeder cells include Primary Mouse Embryonic Fibroblasts (PMEF), Mouse Embryonic Fibroblasts (MEF), Mouse Fetal Fibroblasts (MFF), Human Embryonic Fibroblasts (HEF), differentiated human fibroblasts obtained from human embryonic stem cells, human fetal muscle cells (HFM), human fetal skin cells (HFS), human adult human skin cells, Human Foreskin Fibroblasts (HFF), human cells obtained from the umbilical cord or placenta, human adult human fallopian tube epithelial cells (HAFT), and human bone marrow stromal cells (hMSC). Clusters of hscs can be obtained from adherent cell cultures by dissociating cells from the feeder layer or extracellular matrix to form a suspension of cells. The suspension of cells may comprise a free-floating population or a suspension of substantially single cells from which the population of cells may grow to form a cell population.

According to a preferred embodiment, the cell culture comprises a cell suspension, preferably a free floating population in suspension culture, i.e. an aggregate of cells from human embryonic stem cells (hESC). The source of free floating stem cells was previously described in WO06/070370, which is hereby incorporated by reference in its entirety.

The culturing step according to the present disclosure may comprise culturing the cells in the cell culture with one or more different culture systems, at least one of which comprises one or more members of the TGF β superfamily.

According to one embodiment of the present disclosure, the cells in culture are cultured in a culture system comprising a basal medium supplemented with NA in addition to one or more members of the TGF β superfamily of growth factors.

According to another embodiment, the cells are first cultured in a culture system comprising a basal medium and NA, said cells being undifferentiated hSC, preferably after differentiation of the hSC is induced (i.e. after a predetermined time, or after confirmation of cell differentiation by techniques available in the art), the cells in the cell culture are cultured in a culture system comprising one or more members of the TGF β superfamily of growth factors. The second culture system may also comprise NA, i.e. may be the same as the initial culture system, to which a member of the TGF superfamily is added. As a result, directed differentiation into RPE cells was induced.

According to this embodiment, the hSCs in the initial cell culture are cultured in the NA-containing culture system for at least the time required to initiate differentiation of the hSCs. According to a particular embodiment, the cell culture system is cultured in a culture system comprising NA for several days, preferably for at least two days, preferably for at least one week, more preferably for at least two weeks.

Without being limited by theory, the inventors provide that NA induces a process of directed differentiation, which is also accelerated in its development compared to spontaneous differentiation (i.e., in the absence of NA exposure or exposure to NA and TGF β members). It has been shown here that in directed differentiation, undifferentiated stem cells are eliminated from the culture system more rapidly. Therefore, NA is used in the culture system of differentiating hscs as a means to promote and accelerate the process of committed differentiation for the complete elimination of undifferentiated stem cells, thereby preventing potential complications such as teratoma tumor formation after transplantation due to the presence of undifferentiated cells.

It has been shown herein that exposure of hscs to NA, followed by exposure to one or more members of the TGF β superfamily, induces differentiation into cells with different phenotypes compared to spontaneously differentiating cells (i.e., absence of these factors).

Further, without being limited by theory, the inventors postulate that NA induces differentiation into cells expressing receptors for one or more members of the TGF β superfamily that are not expressed by spontaneously differentiating stem cells, thereby allowing directed differentiation into mature and functional RPE cells. Such receptor expression allows for the inductive effect of TGF β superfamily members on directed differentiation towards RPE fate in differentiated cells in culture, i.e. towards mature and functional RPE cells.

As noted above, there are a number of members of the TGF β superfamily of growth factors. For example, the growth factor according to the present invention may be one or more of TGF β 1, TGF β 2, TGF β 3, activin a, activin B, activin AB, nodal, anti-mullerian hormone (AMH), BMP3, BMP4, BMP5, BMP6, BMP7, or a Growth and Differentiation Factor (GDF). Preferably, however, the growth factor of the TGF β superfamily is TGF β 3 or TGF β 1 or activin a or a combination thereof.

The basal medium according to the invention is any known cell culture medium known in the art for supporting the growth of cells in vitro, generally a medium comprising a defined base solution comprising salts, sugars, amino acids and any other nutrients required to maintain the cells in culture in a viable state. Non-limiting examples of commercially available basal media that can be used according to the invention include Neurobasal^TM、KO-DMEM、DMEM、DMEM/F12、Cellgro^TMStem cell growth media or X-Vivo^TM. The basal medium may be supplemented with various reagents known in the art for processing cell cultures. The following are non-limiting examples of various additives that may be included in the culture system for use in accordance with the present disclosure:

media containing serum or serum substitutes, such as, without limitation, knockout serum substitutes (KOSR), Nutridoma-CS, TCH^TMN2, N2 derivatives or B27 or a combination;

extracellular matrix (ECM) components such as, without limitation, fibronectin, laminin and gelatin. The ECM may be used to carry one or more members of the TGF β superfamily of growth factors;

antibacterial agents such as, but not limited to, penicillin and streptomycin;

-nonessential amino acids (NEAA),

neurotrophic factors known to play a role in promoting survival of SC in culture, such as, without limitation, BDNF, NT3, NT 4.

Once the cells are driven into the RPE fate, RPE cells can be retrieved/harvested from culture by known methods for various applications.

The present disclosure also provides RPE cells obtained by directed differentiation of hscs in the presence of one or more members of the TGF β superfamily. According to one embodiment, the RPE cells are obtained by the method of the invention.

Further to the above, RPE cells produced by directed differentiation according to the present disclosure have specific properties compared to RPE cells that develop during spontaneous differentiation.

Differentiated cells have the potential to respond to TGF signaling in their development and differentiation.

The RPE cells produced are mature cells (terminally differentiated);

mature RPE cells show darker pigmentation compared to RPE cells formed during spontaneous differentiation.

Mature RPE cells express significantly higher levels of transcripts of markers of mature RPE cells, such as wilting and RPE65, compared to their expression in RPE cells produced by spontaneous differentiation. In this regard, for example, with reference to fig. 9J, 11M and 11K, the expression of wilting protein in spontaneous differentiation (no NA) compared to differentiation in the presence of NA (fig. 9J), and the increased potency of activated protein a for directed differentiation (fig. 11K, 11M and 4E) are shown. With further reference to fig. 1B and 9I, expression of RPE65 in spontaneous differentiation (no NA) is shown compared to differentiation in the presence of NA, further compared to the increased potency of activator protein a in fig. 4D.

In Electron Microscopy (EM) analysis, RPE cells exhibit morphological features of mature authentic RPE cells, which are not exhibited in RPE-like cells from spontaneously differentiating hscs, such as apical villi, tight junctions and basement membrane.

RPE cells produced by the presently disclosed methods can be used for large scale and/or long term culture of these cells. To this end, the method of the invention is performed in a bioreactor suitable for large scale production of cells, in which undifferentiated hscs are to be cultured according to the invention. The general requirements for culturing cells in a bioreactor are well known to those skilled in the art.

Alternatively, RPE cells produced by the presently disclosed methods can be expanded following their derivation. For amplification, they were dissociated, plated at low density on extracellular matrix, preferably poly-D-lysine and laminin, and cultured in serum-free KOM with NA. Under these culture conditions, pigmented cells release pigmentation and acquire a fiber-like morphology. After further prolonged culture and proliferation into high density cultures, the cells regain polygonal shape morphology and pigmentation characteristic of RPE cells.

RPE cells may be expanded in suspension or as a monolayer. The expansion of RPE cells in monolayer culture can be modified to large scale expansion in a bioreactor by methods well known to those skilled in the art.

It is well understood by those skilled in the art that derivatization of RPE cells from hscs is of great benefit. They can be used as in vitro models for the development of new drugs to promote their survival, regeneration and function. hSC-derived RPE cells can be used for high throughput screening of compounds that have toxic or regenerative effects on RPE cells. They can be used to reveal mechanisms, novel genes, soluble or membrane-bound factors important for the development, differentiation, maintenance, survival and function of photoreceptor cells.

RPE cells can also serve as an unlimited source of RPE cells for transplantation, supplementation and support of dysfunctional or degenerated RPE cells in retinal degeneration. In addition, the genetically modified RPE cells can serve as a vector to carry and express genes in the eye and retina after transplantation.

Thus, according to a further aspect of the present disclosure, there is provided a method of transplanting RPE cells into an eye of a subject, the method comprising:

(a) providing a cell culture comprising hscs;

(b) culturing cells in a culture system comprising a basal medium supplemented with one or more members of the TGF β superfamily, whereby the hscs are caused to differentiate into RPE cells;

(c) harvesting RPE cells from the cell culture; and

(c) transplanting the differentiated RPE cells into the eye of the subject.

Harvesting of the cells may be performed by various methods known in the art. Non-limiting examples include mechanical dissection and cleavage with papain. Other methods known in the art may also be applied.

hSC-derived RPE cells can be transplanted to various target locations in the eye of a subject. According to one embodiment, the transplantation of RPE cells will be into the subretinal space of the eye, which is the normal anatomical location of the RPE (between the photoreceptor outer segment and the choroid). In addition, depending on the migratory capacity of the cells and/or the active paracrine effect, transplantation into other ocular compartments including the vitreous space, the interior of the outer retina, the periretinal portion and the interior of the choroid may be considered.

Further, transplantation may be performed by various techniques known in the art. Methods for performing RPE transplantation are described, for example, in U.S. patents No.5,962,027, 6,045,791 and 5,941,250, and in eyegraefeses archclinexp oppthalmolmarch 1997; 235(3) 149-58; biochem biophysis resummunfeb.24, 2000; 268(3): 842-6; OpthalmicSurgFebruary 1991; 22(2): 102-8. Methods for performing corneal transplants are described, for example, in U.S. patent No.5,755,785 and in Eye 1995; 9(Pt6Su) 6 to 12; curropinoptalmollugst 1992; 3(4): 473-81; OphthalmicSurgLasersApril 1998; 29(4): 305-8; ophthalmologiclaril 2000; 107(4): 719-24; andJpJOphthalmolNovember-December 1999; 43(6): 502-8. If the paracrine effect is to be used primarily, cells may also be delivered and maintained in an eye sealed with a semi-permeable container, which will reduce exposure of the cells to the host immune system (Neurotech USACNTFdelivery system; PNASMArch7, 2006vol.103(10) 3896-.

According to one embodiment, the transplantation is performed by a parsanana vitrectomy procedure followed by delivery of cells into the subretinal space through a small retinal opening, or by direct injection. Alternatively, cells may be delivered to the subretinal space by a transscleral, transchoroidal approach. In addition, direct transscleral injection into the vitreous space, or delivery to the outer peripheral portion of the anterior retina near the ciliary body, may be performed.

RPE cells can be transplanted in various forms. For example, RPE cells may be introduced into a target site in the form of a cell suspension, or attached to a matrix, extracellular matrix, or substrate, such as a biodegradable polymer, or a combination thereof. RPE cells may also be transplanted (co-transplanted) with other retinal cells, for example, with photoreceptors.

Thus, the present invention also relates to a composition comprising hSC-derived RPE cells obtained by the method of the invention. The composition is preferably suitable for implantation into the eye.

By introducing the RPE cells obtained by the methods of the invention into the eye of a subject, various ocular conditions can be treated or prevented. The eye condition may include a retinal disease or disorder generally associated with retinal dysfunction, retinal damage, and/or loss of retinal pigment epithelium. A non-limiting list of conditions that may be treated according to the present invention includes retinitis pigmentosa; congenital leber's amaurosis, hereditary or acquired macular degeneration, age-related macular degeneration (AMD), Best's disease, retinal detachment, gyratory atrophy, choroideremia, pattern dystrophy (patterndystrophy) and other dystrophies of the RPE, Stargardt's disease, RPE and retinal damage due to damage caused by any of light, laser, inflammation, infection, radiation, neovasculature or traumatic injury.

Without being limited by theory, transplanted RPE cells may exert their therapeutic effects through a variety of mechanisms. One mechanism is a trophic supporting effect that promotes the survival of degenerating photoreceptors or other cells within the retina. RPE cells from hscs by the methods of the present disclosure, and in the presence of TGF β superfamily members, are able to potentially protect the photoreceptors adjacent to them by trophic effects.

Transplanted RPE cells may also exert their effects by replenishing the regenerative mechanisms of dysfunctional and/or degenerated host RPE cells. RPE cells derived from hscs can be supplemented with dysfunctional host RPEs by the methods of the present disclosure and in the presence of a TGF β superfamily member. The transplanted cells are mature and have the functional ability to phagocytose the shed outer segment of the photoreceptor that includes rhodopsin.

As described above, RPE cells derived from hscs by the methods of the present disclosure and in the presence of a member of the TGF β superfamily are mature and thus they do not proliferate in vivo following transplantation. Thus, RPE cells derived from hscs by the methods of the present disclosure are safer for transplantation therapy with reduced risk of developing teratoma tumors or tumors of proliferative precursor cells.

As used herein, the term "treating" refers to both therapeutic and prophylactic effects of the hSC-derived RPE cells of the present invention on a condition of the eye of a subject, which effects generally include improving symptoms associated with the condition, reducing the severity of the condition, or curing the condition, and more specifically, which effects may include reversing damage to the retina and RPE of the treated subject, improving the function of the retina of the subject, reconstructing the retina and RPE of the subject, either directly or through paracrine effects by replacing and/or supporting the failing host retina and RPE cells, and the prophylactic effects exhibited by attenuating, inhibiting, or halting the damage caused to the retina of the subject as a result of the condition.

As used in this specification and the claims, the forms "a", "an" and "the" include both singular and plural referents unless the context clearly dictates otherwise. For example, the term "a growth factor" includes one or more growth factors, and the term "growth factor" includes one growth factor and more than one growth factor.

As used herein, the term "or" refers to one or a combination of two or more of the listed options. Further, the term "selected from" is used to include one or a combination of two or more of the listed options, followed by a list of options separated by the terms "and".

Further, as used herein, the term "comprising" is intended to mean that the method or composition includes the recited elements, but does not exclude others. Similarly, "consisting essentially of … …" is used to define methods and systems that include the recited elements but exclude other elements that may be of essential importance to the functionality of the culture system of the invention. For example, a culture system consisting essentially of a basal medium, medium supplements, and feeder cells will not include, or will only include, insignificant amounts (which will have insignificant effects on cell proliferation and differentiation in the culture system) of other substances that have an effect on the cells in the culture. Also, a composition consisting essentially of the elements defined herein will not exclude trace contaminants from the separation and purification process. "consisting of … …" will mean excluding more than trace amounts of other elements. Embodiments defined by each of these transition terms are within the scope of the present invention.

Further, all numbers, such as concentrations or dosages or ranges thereof, are approximations that vary (+) or (-) from the stated value by up to 20%, and sometimes up to 10%. It is understood that all numerical symbols are preceded by the term "about," even if not necessarily explicitly stated. It is also to be understood that the reagents described herein are exemplary only, although not necessarily explicitly stated, and that their equivalents are known in the art.

Certain exemplary embodiments

Materials and methods

HES cell culture

Human ESCs (HES1 cell line) and hESCs [ GroppM, Itsykson P, SingerO, Ben-HurT, Reinhartze, Galune, and ReubinfBE.Stablegeneticmodificationno humanmanemexcellent cells based on viral vectors, molecular therapy7:281-7(2003)]Cultured on human foreskin fibroblast feeder layer in KO medium (KOM) consisting of 86% KO-DMEM (Gibco, Invitrogen, Gaithersburg, Md), 14% KOSR (Gibco), 1mM glutamine, 1% non-essential amino acids, 50 units/ml penicillin (Gibco), 50. mu.g/ml streptomycin, (Gibco) and 4ng/ml LBFGF (R)&DSystems, inc., Minneapolis, MN). hES cells were passaged weekly with collagenase type IV (1 mg/ml; Gibco) and plated on fresh feeder layers. One week prior to induction of differentiation by treatment with Ca/Mg free supplemented with 0.05% EDTA (biologicals industries, BeitHaemek, Israel)⁺⁺PBS was dissociated into a near single cell suspension to passage cells and replated on feeders.

EB formation in suspension cultures

Six-eight days after plating the hES cells isolated as single cells as described above; they were removed from the feed by treatment with collagenase IV. The pellet was cultured in suspension for up to 12 weeks in a bacteriological plate precoated with 0.1% low melting temperature agarose in the presence or absence of 10mM Nicotinamide (NA) (Sigma, St. Louis, Mo., USA) in KO medium (KOM) consisting of 86% KO-DMEM, 14% KOSR, 1mM glutamine, 1% non-essential amino acids, 50 units/ml penicillin and 50. mu.g/ml streptomycin. In some experiments, the medium used was Neurobasal supplemented with N2 supplement (1:100) (Gibco)^TMMedium (Gibco) (NN medium) which was replaced after 1 week with DMEM/F12(Gibco) supplemented with B27(1:50) (Gibco).

hESC differentiation into RPE cells in the presence of TGF-beta growth factor or inhibitor

Human ESCs were allowed to differentiate in the presence of Nicotinamide (NA)10mM as free-floating clusters in KOM as described above for up to six weeks. After the first or 2 weeks of differentiation, the cultures were supplemented with activin A20-180ng/ml (PeproTechInc, RockyHill, NJ), TGF β 3(1 ng/ml; R & DSystems Inc, Minneapolis, MN), TGF β 1(1ng/ml-20 ng/ml; R & DSystems Inc) or SB431542 (5. mu.M-50. mu.M, Sigma). Control media was supplemented with NA alone.

Human ESCs suspended in KOM were also supplemented with Bone Morphogenetic Protein (BMP) antagonist noggin (700ng/mlR & dsystemss inc, Minneapolis, MN) in the presence or absence of NA after one week, or FGF β (20ng/ml pepropetechinc) in the presence of NA at weeks 3 and 4, and allowed to differentiate as free-floating clusters in suspension up to 6 weeks of age.

Description of amplification of RPE cells

To expand the RPE cells, the pigmented clusters were gently mechanically dissociated into small pieces, low density plated on poly-D-lysine (30-70kDa, 10. mu.g/ml) and laminin (4. mu.g/ml), and cultured in KOM with NA. Under these culture conditions, pigmented cells lose pigmentation and acquire a fiber-like morphology. After further culture for 1.5 months and proliferation into high density cultures, the cells regained the polygonal shape morphology and pigmentation characteristic of RPE cells.

All cultures were immunostained and subjected to real-time RT-PCR as described below.

Indirect immunofluorescence staining of differentiated cells within a cluster

To characterize the immunophenotype of cells within aggregates, clusters cultured for 2,4, 6, or 8 weeks were gently dissociated using 0.04% trypsin/0.04% EDTA, or with a papain dissociation system (worthington biochemical, Lakewood, NJ), resulting in clumps and single cell platingIn KO medium supplemented with NA on poly-D-lysine (30-70kDa, 10-20. mu.g/mL) alone or with laminin (4. mu.g/mL) or fibronectin (10-20. mu.g/mL; all from Sigma, St. Louis, http:// www.sigmaaldrich.com). After 2 hours the cells were fixed with 4% paraformaldehyde and examined for nestin (1:200), polysialic acid NCAM (PSA-NCAM) (1:100), Musashi (1: 200; all from Chemicon, Temecula, from CA), Pax6(DSHB, 1:100 or Chemicon, 1:250), Otx2(Chemicon, 1:200), MiTF (LabVision corporation, Fremont, CA; mouse IgG; mouse₁1: 50).

For immunostaining of enriched preparations of pigmented cells, pigmented (Brown) clusters of cells within 8-10 week differentiated floating clumps were mechanically dissected and separated by glass micropipettes or dissecting blades (No 15; Swann-Morton sheffield, Eng).

The isolated clusters enriched in pigmented cells were mechanically dissociated into smaller pieces by digestion with/without trypsin (0.025%, 3mM EDTA in PBS) or papain dissociation (papain dissociation system; Worthington Biochemical corporation, Lakewood, New Jersey) assisted grinding. Small clusters of cells were plated on poly-D-lysine coated (30-70kDa, 10. mu.g/ml; Sigma) and laminin coated (4. mu.g/ml; Sigma) glass coverslips and cultured for an additional 3-5 weeks in the media used for suspension culture of hESC clusters. Differentiated cells within the growth were fixed with 4% paraformaldehyde at room temperature for 30 minutes. For immunostaining with anti-intracellular marker antibodies, cell membranes were permeabilized with 0.2% triton x100(Sigma) in PBS supplemented with normal goat serum for blocking (5%, biologicals industries) for 30 min. Cells were incubated with the following primary antibodies: anti-MiTF (LabVision corporation, Fremont, Calif.; mouse IgG)₁1: 50); anti-RPE 65(Novus biologicals, Littleton, CO; mouse IgG)₁1: 300); anti-wilting protein (Novus biologicals; mouse IgG)₁1: 150); anti-ZO-1 (zymed laboratories Inc., san Francisco, Calif.; Rabbit polyclonal, 1: 10); anti-Ki 67 (DakoDenemarkA/S;, 1:50) and anti-CRALBP (by john c. saari,university of washington, Seattle, a benefit; rabbit polyclonal, 1: 100). Cells were also incubated with phalloidin (1:200 Sigma).

Primary antibody localization was achieved by using Fluorescein Isothiocyanate (FITC) -conjugated goat anti-mouse immunoglobulin (DakoDenmark A/S; 1:20-1:50), conjugated to Cy^TH3 (1:500) (jackson immunoresearch laboratories inc, WestGrove, PA), rabbit anti-goat IgG conjugated to Cy2 (1: 200; jackson immunoresearch laboratories inc), and pig anti-rabbit Ig conjugated to Fluorescein Isothiocyanate (FITC) (Dako; 1: 50).

Analysis of hESC clusters by RT-PCR and real-time PCR

Total RNA was extracted from hescs grown at consecutive time points up to 8 weeks during culture of hESC-derived clusters in the presence or absence of 10mM nicotinamide, grown under serum-free conditions (1 week after passage) and with or without supplementation with TGF β superfamily growth factors or antagonists. RNA was isolated using TRIzol reagent (Invitrogen, http:// www.invitrogen.com) or TRI-reagent (Sigma). cDNA synthesis was performed using Moloney murine leukemia virus reverse transcriptase (M-MLVRT) and random primers according to the manufacturer's instructions (Promega corporation, Madison, Wis., http:// www.promega.com). Polymerase Chain Reaction (PCR) was performed using TaqDNA polymerase (GIBCO-BRL) using standard protocols. The amplification conditions were as follows: denaturation at 94 ℃ for 15 seconds, annealing at 55 ℃ for 30 seconds, and extension at 72 ℃ for 45 seconds. The number of cycles varied between 18 and 40 depending on the particular mRNA abundance. The length of the primers human sequence and amplification product used to identify the human gene transcription products (forward and reverse 5 '-3') are as follows (SEQ ID NOs: 1-12):

MiTF-A(GAGCCATGCAGTCCGAAT，GACATGGCAAGCTCAGGACT；486bp)；

RPE65(GCTGCTGGAAAGGATTTGAG，CAGGCTCCAGCCAGATAGTC；231bp)；

wilsonin (GAATTTGCAGGTGTCCCTGT, ATCCTCCTCGTCCTCCTGAT; 214 bp);

CRALBP(AGCTGCTGGAGAATGAGGAA，CAAGAAGGGCTTGACCACAT；218bp)

MERTK(AAGTGATGTGTGGGCATTTG，TCTAAGGGATCGGTTCTCCA，189bp)；

ACTRIA(AATGTTGCCGTGAAGATCTTC，CTGAGAACCATCTGTTGGGTA；699bp)；

ACTRIB(CACGTGTGAGACAGATGGG，GGCGGTTGTGATAGACACG；346bp)；

ACTRIIA(AACCATGGCTAGAGGATTGGC，CTTTCACCTACACATCCAGCTG；551bp)；

ACTRIIB(CACCATCGAGCTCGTGAAG，GAGCCCTTGTCATGGAAGG；611bp)；

activin A (CTTGAAGAAGAGACCCGAT; CTTCTGCACGCTCCACCAC; 262 bp);

beta-actin (TTCACCACCACGGCCGAGC, TCTCCTTCTGCATCCTGTCG; 351 bp);

GAPDH(AGCCACATCGCTCAGACACC；GTACTCAGCGCCAGCATCG；301bp).

for real-time PCR, TaqMan from a commercially available source was used^RTaqMan primers and probes for the Assays-on-Demand Gene expression product (applied biosystems, Foster City, Calif.) monitor the level of transcripts:

oct4, IDHs 01895061; musashi, IDHs 01045894; pax6, IDHs 00240871; six3, IDHs 00193667; rx1, IDHs 00429459; chx10, IDHs 01584048; MiTF-A, IDHs 01115553; MiTF-total, IDHs 01115557; bestrophin, IDHs 00188249; RPE65, IDHs 00165642; sox10, IDHs 00366918; crx, IDHs 00230899. Sequence detection systems and methods using ABIPrism7000HT and ABIPrism7900HTHousekeeping gene β -glucuronidase (gusB, analytical IDHs99999908) was selected as an internal reference standardized in real-time RT-PCR quantitation, the relative expression level of each gene being shown when the expression level of day 0 (or untreated cells) was set to 1Relative values. Amplification reactions were performed in duplicate or triplicate according to the manufacturer's protocol (applied biosystems).

Transmission electron microscopy and phagocytosis of latex beads

Human ESC-derived clusters were cultured in KOM in suspension. The pigmented areas were then mechanically separated and processed for transmission electron microscopy. Cells were fixed with 2% glutaraldehyde and 4% formaldehyde in 0.1M cacodylate buffer pH 7.4. After three washes in 0.1M cacodylate buffer, the tissues were post-fixed with 1% osmium tetroxide and 1.5% potassium ferricyanide, dehydrated with increasing concentrations of ethanol, and embedded in Agar100 resin. Ultrathin sections cut by lkbutrotome 3 were stained with uranyl acetate and lead citrate. Micrographs were taken with a Tecnai12 electron microscope (Phillips, Eindhoventhenerlandeshttp:// www.philips.com) equipped with a MegaviewIICCD camera and Analysis version 3.0 software (Softi imaging System, http:// www.soft-imaging. net).

To examine phagocytic capacity, the pigment was clustered and concentrated at 1.0 × 10⁹1 μm latex beads (Polysciences Inc., Warrington, Pa.) per bead/ml were incubated at 37 ℃ for 18 hours. The pigmented clusters were then washed with PBS +, dissociated into single cells or small pieces using a papain dissociation system and plated on poly-D-lysine. After fixation, the cell membrane was stained with the red fluorescent dye pkh (sigma). Phagocytosis was analyzed using confocal microscopy (olympus fluoviewfv 1000).

Flow cytometry

Flow cytometry analysis was performed to determine the number of PSA-NCAM and TRA-1-60 positive cells at different time points within hESC-derived clusters differentiated with or without nicotinamide supplementation. The clusters were dissociated with 0.04% trypsin/0.04% EDTA. Single cells were then stained with anti-PAS-NCAM or anti-Tra-1-60 antibody (both from Chemicon; 1:100), detected with goat anti-mouse immunoglobulin conjugated to FITC (Dako; 1:100), and counterstained with the cell viability dye propidium iodide (0.005 mg/ml; Sigma). Control cells were incubated with secondary antibody only. Cell-associated immunoreactivity was analyzed using CellQuest software with FACScalibur (Becton Dickinson immunocytometric systems).

Intravitreal and subretinal transplantation of hESC-derived differentiated RPE cells

For intraocular transplantation, hescs engineered to express eGFP [ as described earlier in gropepetal, statistical modificationofhumanorganizing cells based analysis methods molecular therapeutics 2003; 7:281-7 ] was used to produce RPE cells in culture as described above. Briefly, clusters enriched for pigmented cells were mechanically isolated by dissection after 6-8 weeks of differentiation in the presence of NA alone or in the presence of NA supplemented with activin a. To allow injection through small bore glass capillaries, the pellet was further dissociated into smaller clusters of cells by papain (Papaiin DissociationSystem; Worthington Biochemical corporation, Lakewood, NewJersey) digestion for 30 minutes at 37 ℃ followed by grinding.

Fifteen adult albino rats (weighing 230-. In RCS rats, mutations in the Mertk gene cause RPE dysfunction, which leads to retinal degeneration in the first few months of life. All animal experiments were performed according to the ARVO statement of the use of animals in ophthalmological and visual studies, approved by the committee of the animal research institute of Hadassah medical school, university of Hebrew.

For transplantation (and for electroretinal image recording), animals were anesthetized with ketamine HCl (Ketalar, ParkeDavis, UK; 100mg/kg) and injected intraperitoneally with the relaxing agent xylazine (2.0 mg/kg). Local anesthetic drops (oxybuprocaine hcl 0.4%; fischer pharmaceuticals, Israel) were administered. The pupil was dilated with tropicamide 0.5% (Mydriamide, Fisher pharmaceuticals, Israel) and phenylephrine HCl 2.5% (Fisher pharmaceuticals, Israel). Approximately 100,000 cells in 4 μ L of medium were injected into the vitreous or subretinal space by transscleral, or suprachoroidal, methods via glass capillaries connected to pneumatic Pico syringes (PLI-100; medical System Corp., Greenvale, NY, http:// www.medicalsystems.com) under visualization by a dissecting microscope (Stemisv11, Zeiss, Germany), and the uninjected, same type of eye served as one type of control. As an additional control, eyes were injected with saline (sodium chloride injection BP, 0.9%, b.braun Melsungen ag, Melsungen, Germany).

No choroidal hemorrhage was observed during and after injection. The animals were kept warm throughout and after operation using heating lamps. After transplantation, all animals received the immunosuppressive agent cyclosporin A (Sandimmune, Novartis PharmaAG, Basel, Switzerland) at a concentration of 210mg/l in their drinking water.

In vivo and ex vivo imaging of transplanted cells

To monitor the survival and location of transplanted cells in vivo, anesthetized animals were imaged using a color fundus camera (Zeiss, Germany) and fluorescence of GFP expressing cells was detected using a fluorescein filter on a scanning laser ophthalmoscope (HeidelbergHRA, Germany). In some eyes, the location of GFP positive grafts was also determined ex vivo in an eyecup preparation using a fluorescence microscope (Canon, Japan).

Assessment of host retinal function following intraocular transplantation of hESC-derived RPE cells

Retinal function was assessed in transplanted and control RCS rat eyes by Electroretinography (ERG) four to six weeks after transplantation. Full field ERG was recorded after dark adaptation overnight. Animals were paralyzed in dim red light with ketamine and xylazine and the pupils were dilated with tropicamide and phenylephrine. Monopolar rat ERG lens electrodes (lentiselectrode) (medical works, Amsterdam, Netherlands) were placed on each eye after additional local anesthesia, with the reference and ground electrodes placed on the tongue and tail, respectively. A commercially available computerized ERG system (LKC technology, UTAS3000) was used to record visual acuity for full-field stimulation using xenon strobe flash (Grass, PS-22) placed on a Ganzfeld bowlAnd (4) performing omentum reaction. A faint blue flash under scotopic conditions is used to elicit a primary rod-driven response. Mixed cone-rod responses were recorded under a more intense stimulus intensity of blue and white flash under dark adaptation conditions. Using a rod suppression of 34cd/m under light adaptive (photopic) conditions²White flashing of the white background produced cone responses at 1Hz and 16 Hz. The signal was filtered between 0.3-500Hz using signal averaging.

Histological and immunohistochemical evaluation of transplanted eyes

Animals were sacrificed 4-8 weeks post-transplantation and eyes were enucleated for histological and immunohistochemical examination. After fixation in Davidson's solution, the eyes were embedded in paraffin and sectioned in 4 μm serial sections. Every five sections were stained with hematoxylin and eosin for histomorphological assessment and quantification. For the indirect immunofluorescence studies, to characterize the state of differentiation of the transplanted cells, the samples were deparaffinized in xylene and dehydrated in gradient alcohol, rinsed with phosphate buffered saline (PBS, ph7.4), and incubated with 10mM citrate buffer (ph6.0) at 110 ℃ for 4 minutes. After washing with PBS, samples were blocked with PBS solution containing 1% Bovine Serum Albumin (BSA), 0.1% triton x100(Sigma-Aldrich) and 3% normal goat or normal donkey serum for 1 hour at room temperature. Subsequently, the sections were incubated in a humidified room for 1 hour with the appropriate combination of the following primary antibodies: anti-green fluorescent protein (anti-GFP), conjugated with Fluorescein (FITC) or rhodamine (TRITC) (Santa Cruz Biotechnology, Inc, Santa Cruz, Calif.; monoclonal for mice, 1: 100); anti-RPE 65(Novus biologicals, Littleton, CO; mouse IgG)₁1: 100); anti-wilting protein (Novus biologicals; mouse IgG)₁1: 100); anti-ZO-1 (zymed laboratories Inc., san Francisco, Calif.; rabbit polyclonal, 1: 100); and anti-rhodopsin (Santa Cruz Biotechnology, Inc, Santa Cruz, CA; rabbit polyclonal, 1: 100). After washing in PBS, by using Cy^TM2 conjugated goat anti-Rabbit IgG (1:200), Cy^TM2 conjugated goat anti-mouse IgG (1:200), Cy^TM3 conjugated goat anti-Rabbit IgG (1:200), Cy^TM2 affixConjugated donkey anti-mouse IgG (1:200), Cy^TM5 conjugated donkey anti-rabbit IgG (1: 200; all from Jackson ImmunoResearch laboratories, Inc, WestGrove, Pa.) for primary antibody localization. Nuclei were counterstained with 4, 6-diamino-2-phenylindole (DAPI) containing mounting media (vector laboratories, Burlingame, Calif.) or with 1. mu.g/ml propidium iodide (BioLegend, SanDiego, Calif.). To determine the specificity of the antigen-antibody reaction, a corresponding negative control with irrelevant isotype-matched antibodies was performed. An Olympus bx41 microscope equipped with a DP70 digital camera (Olympus, Japan) was used for fluorescence and light microscopy imaging. Confocal images were acquired on an Olympus fluoview300(FV300) confocal microscope (Olympus, Japan) constructed around an IX70 inverted microscope. 488-nmAr, 543 HeNe-green and 633HeNe red lasers were used in combination with Nomarski optics.

Quantification of photoreceptor layer rescue in proximity to RPE graft

To quantify the efficacy of hESC-derived RPE transplantation on degenerated host retinas, high resolution microscope images of hematoxylin and eosin stained sections were obtained, and Photoshop software (Adobe, USA) was used to construct full-length montage of the retinas. The total retinal thickness, the thickness of the outer nuclear (photoreceptor) layer, and the thickness of the inner and outer fragment layers were measured using the J-image program (NIH) at subretinal displacement plants in proximity to hESC-derived RPE cells. These were compared to measurements obtained in the corresponding contralateral side of the retina distal to the graft. Since the degenerative process in RCS rats is position-dependent, the thickness is measured in regions equidistant from the ciliary body. In each zone, at least three equidistant measurements are averaged.

Results

Characterization of differentiated RPE

Differentiation of hescs was induced by culturing them as free-floating clusters in KO medium supplemented with NA. Under these culture conditions, a defined region of highly concentrated pigmented cells developed within the cluster under differentiation, as shown in FIG. 3A. These pigmented regions appeared after 4 weeks of differentiation, with 72.9 ± 2.5% of the clusters having pigmented regions after 8 weeks. No pigmented regions were observed after 4 weeks of differentiation without NA supplementation, and under these conditions only 13.1 ± 4.8% developed into pigmented regions after 8 weeks (fig. 8A and 8B). Thus, NA treatment increases/promotes differentiation into pigmented cells within hESC clusters compared to hESC clusters that are spontaneously differentiating.

Within the 8-week differentiated cluster in the presence of NIC, 5.7 ± 1.0% of the cells were pigmented, 5.4 ± 1.1% expressed the early RPE marker MiTF, and expression of MiTF was mostly associated with pigmentation (fig. 8C). The partially dissociated and plated clusters of differentiated hescs developed into colonies consisting of monolayers of pigmented cells, along with other types of differentiated cells, as shown in the dark-field micrographs (fig. 3D) and the phase-contrast images (fig. 3E). The cells within these colonies assumed a polygonal shape, forming a "cobblestone" like sheet of cells with tight junctions between them (FIG. 3E), with the highly characteristic features of native RPE cells. Intracellular F-actin distribution was close to their membranes, as demonstrated by staining with phalloidin, comparable to authentic RPE cells (fig. 10A). Pigmented RPE cells co-expressed RPE markers Otx2 (fig. 3B) and MiTF-a (fig. 3B and fig. 3F) as well as ZO-1 (fig. 3G), wilting protein (fig. 3H), RPE65 (fig. 3I) and CRALBP (fig. 3J).

After dissociation, low density plating and culture, the pigmented cells lost pigmentation and acquired a fiber-like morphology as shown by phase contrast imaging (fig. 3K and 10B). After further prolonged culture and proliferation into high density cultures, the cells regained the polygonal shape morphology and pigmentation of the RPE cells (fig. 3L and 10C).

Electron Microscopy (EM) analysis revealed that hESC-derived pigmented cells had the morphological features of native RPE cells, including microvilli on their top surface (fig. 10D) and basement membrane on their basal surface (fig. 10E). The cells contained melanin microparticles (fig. 10D) and were attached by tight junctions (fig. 10F).

One of the most important functions of RPE cells is phagocytosis of external fragments shed by photoreceptors. To examine whether hESC-derived pigmented cells were phagocytic, they were incubated with 1 μm fluorescent latex beads. The three confocal fluorescence images represent sequential Z-axis slices (fig. 10H-10J). Confocal microscopy analysis showed that putative RPE cells were able to phagocytose fluorescent beads (fig. 10G-10J).

Differentiation-inducing Effect of Nicotinamide

Differentiation of hescs towards RPE fates was examined by culturing them as free-floating clusters in KOM with or without NA (clusters in spontaneous differentiation were not provided as controls). After 6 weeks of differentiation, the expression levels of the markers MiTF-a and RPE65 of RPE cells were significantly increased in the presence of NA, as determined by real-time RT-PCR (fig. 1A and 1B, respectively). The expression of MiTF-A was almost two-fold increased in the presence of NA, while the expression of RPE65 was almost 30-fold increased. Since most pigmented cells co-expressed MITF-a (fig. 8C), it appears that there was a significant and prominent increase in the level of RPE65 expression per pigmented cell in the presence of NA. Thus, in addition to its inductive effect towards the outcome of RPE, NA promotes the maturation of RPE cells and has an effect on the phenotype of the cells. Q-PCR analysis at successive time points between 2 and 6 weeks showed an increase in MiTF-a and RPE65 expression levels in the presence of NA (fig. 1C and 1D, respectively). Expression of the marker was upregulated after four weeks of differentiation, and levels of expression continued to increase during the next four weeks (last two weeks not shown). Similar increased expression of the RPE markers, wilson, CRALBP and Mertk, was shown by RT-PCR of cells in plated, pigmented clusters (fig. 1E).

To find whether the in vitro development of RPE cells recapitulates the key developmental steps of RPE in vivo, time course experiments were performed, including analysis of the expression of key markers within hESC clusters during RPE development. Cluster differentiation in the presence or absence of NA was analyzed by real-time PCR of expression of markers of undifferentiated hescs, early neural differentiation, retinal and RPE development (fig. 9A-9L). First, it was shown that the expression of the marker Oct4 of undifferentiated hESC (fig. 9A) decreased more rapidly during differentiation in the presence of NA. Consistently, FACS analysis revealed that the expression of the surface membrane marker TRA-1-60 of undifferentiated cells was also decreased more rapidly in NA-treated samples (fig. 9M). Thus, differentiation in the presence of NA can be used to eliminate undifferentiated cells from culture and can help avoid teratoma tumor formation after transplantation.

In addition, NA treatment enhances the process of early neural differentiation. In the presence of NA, expression of transcripts of the early neural markers Otx2 (fig. 9B), Pax6 (fig. 9D), and Musashi (fig. 9C) increased significantly after 2, 2-6, and 4-6 weeks of differentiation, respectively. Similar results were shown at the protein level by FACS analysis of expression of the neural precursor marker PSA-NCAM (fig. 9O). After 4 weeks of differentiation with NA, 81.4 ± 6.3% of the cells expressed PSA-NCAM compared to 14.4 ± 5.9% in the control cluster, indirect immunofluorescent staining confirmed that, at 4 weeks, most of the cells within the NA-treated cluster acquired a neurophenotype and expressed PSA-NCAM (74.2 ± 4.1%), nestin (55.9 ± 10.1%) and Musashi (71.4%; fig. 9N).

Expression of transcripts of the key regulatory genes for retinal specification and morphogenesis, Rx1 and Six3 (fig. 9F and 9E, respectively), were shown at 2 weeks post-differentiation. NA treatment increased the expression of these genes.

In the presence of NA, expression of the transcript of the early RPE marker MiTFA was induced after 4 weeks of differentiation (fig. 9H). Expression of the markers of more mature RPE, wilforin and RPE65, were primarily upregulated after 4 and 8 weeks, respectively (fig. 9J and 9I, respectively). Expression of these transcripts was also higher in NA treated cultures compared to spontaneously differentiating hESC clusters as controls (fig. 9I and 9J). The expression of RPE65 increased more than 100-fold, further confirming that NA has an effect on the phenotype of the cells in addition to inducing effects. To exclude that the obtained pigmented cells were not neural crest derived melanocytes, expression of Sox10 was demonstrated, which is a developmental marker for these cells, which was low compared to control cells of the M51 melanoma cell line and not dependent on NA supplementation. The culture thus does not consist of melanocytes. Thus, it was concluded that NA promotes the induction of differentiation towards the outcome of RPE.

NA treatment also increased expression of Chx10, which modulated proliferation of neural retinal progenitor and photoreceptor progenitor marker Crx (fig. 9K).

In summary, the inventors concluded that the process of RPE differentiation within hESC clusters goes through a similar step to true RPE development in vivo and is augmented by NA. In addition, NA has an effect on the phenotype of the RPE cells obtained. These cells are distinct from RPE cells obtained after spontaneous differentiation and express markers of mature RPE cells at significantly higher levels. NA showed an inducing effect independent of the culture medium

NA supplementation increased pigmentation in differentiated cells cultured for 12 weeks in KO medium and NN/DMEM medium, as shown by dark field micrographs of clusters of differentiating hescs (fig. 2A-2D). In NN medium further replaced with DMEM/F12-B27(NN/DMEM) one week later, the percentage of pigmented hESC clusters in the total number of clusters was higher compared to KO in the presence of NA, while their size and total population number were smaller relative to clusters cultured in KO medium. RT-PCR analysis showed that NA supplementation increased the expression of MiTF-A by about 3-fold in KOM and about 2.5-fold in NN/DMEM medium (FIG. 2E). Expression of RPE65 was approximately doubled in KOM and increased almost 6-fold in NN/DMEM medium when supplemented with NA compared to no NA (FIG. 2F). Thus, the differentiation-inducing effect of NA was shown in differentiated RPE cells, independent of the medium in which hescs were cultured and differentiation occurred.

Effect of members of the TGF-beta superfamily on SC differentiation

First analyzed was the expression of activin receptors and activin a in the 2-week-old cluster. Analysis was performed at this time point, since expression of early eye region (eyefield) markers occurred at this time, and thus the differentiating cells were likely in a developmental stage parallel to the early blebs. It was shown that expression of the receptors ACTRIB and ACTRIIB was high in the presence of NA compared to the lack or lesser expression when cells differentiated without NA (figure 11G). Thus the presence of NA has an effect on the phenotype of cell differentiation.

Activin a was found to enhance differentiation towards RPE fate. Dark field micrographs of hESC-derived clusters differentiated for 4 weeks showed the appearance of pigmented cells at this early stage in the presence of activator protein (fig. 11A). It was further shown that activin a significantly enhances hESC differentiation towards RPE cells in the presence of NA (fig. 4A-4B and 11B-11C). The percentage of clusters including pigmented cells (50.7 ± 6.5vs17.7 ± 3.2), and the percentage of pigmented cells from the total number of cells (9.9 ± 1.4vs2.4 ± 1.2), was significantly higher when differentiation was induced in the presence of activin a compared to control cultures supplemented with NA without activin a (fig. 11H and 11I). This result was confirmed with RT-PCR, which showed that activin a treatment significantly increased (more than five-fold and more than four-fold, respectively) the expression of markers RPE65 (fig. 4D) and wilting (fig. 4E) specific for mature RPE cells. Furthermore, the morphological characteristics of the clusters of pigmented cells that develop in the presence of activin a are different. Their pigmentation was darker, they showed a very clear boundary with surrounding non-pigmented cells (fig. 4B). The expression of the marker MiTF-a, which occurs earlier during RPE development, is not affected by the complement of activator protein a. Thus, activin a, a member of the TGF β superfamily of growth factors, enhances differentiation of hescs into RPE cells. Supplementation of the activin inhibitor SB431542 significantly reduced the appearance of pigmented clustering under these conditions (fig. 11E and 11H). The effect of activin a on RPE differentiation was studied at various concentrations. The effect was found to be dose dependent on the percentage of pigmented cells and the expression of the RPE markers wilsonin (fig. 11K) and RPE65 (fig. 11L), with the best enhancement at 140 ng/ml. In most experiments, activin a was added for two weeks (weeks 3-4) after the clusters had differentiated for 2 weeks, as we found that application at this time was optimal for enhancing RPE differentiation. Considering the observations that expression of markers of early eye development also started after 2 weeks of differentiation (fig. 9A-9L), it appears that activator proteins enhance the process of eye and RPE development. Furthermore, since the percentage of pigmented cells increased 5-6 fold in the presence of activin A, and the expression of markers of mature RPE cells such as wilting protein increased about 10 fold, it appears that activin A has an effect on the maturation and phenotype of the cells in addition to its inducing effect. This is supported by the morphological appearance of the pigmented cells obtained in the presence of activin a, which are darker with clear boundaries with the surrounding cells.

Q-PCR time course analysis of the effect of 2-week activin a treatment on gene expression showed that activin a significantly increased expression of retinal progenitor markers Rx1 and Chx10, as well as the RPE marker, wilsonin. Activin A treatment also increased the expression level of the total MiTF isoforms at 4 weeks of differentiation (FIGS. 11M-11P).

The effect of activin a on the induction of retinal and RPE differentiation was also observed with other members of the TGF β superfamily. Treatment of the differentiating clusters with TGF β 3 significantly enhanced the expression levels of transcripts of MiTF-a, which plays a key role in RPE development in vivo (fig. 5F). Furthermore, treatment with TGF β 1, another member of the TGF β superfamily, also significantly increased the appearance of clusters with pigmented cells (fig. 11D and 11H). In contrast, differentiation in the presence of NA and basic fgf (bfgf), but not factors from the TGF β superfamily, abolished the appearance of pigmented cells (fig. 11F).

Furthermore, it was shown that Bone Morphogenetic Proteins (BMP) belonging to the second subfamily of the TGF β superfamily play a role in RPE differentiation of hescs. As shown in the dark field micrograph, the differentiation of hESC clusters into pigmented RPE cells was blocked in the presence of the BMP antagonist noggin (fig. 5D). Even when the medium was also supplemented with NA, differentiation into pigmented cells was blocked by addition of noggin (fig. 5C). At the RNA level, real-time RT-PCR demonstrated that noggin significantly reduced MiTF-a expression both in the presence and absence of NA in the culture medium (fig. 5E). Thus, BMP plays a role in inducing differentiation of hescs into RPE cells.

Differentiated RPE cells derived from hESCs can be used for intraocular transplantation

An enriched population of hESC-derived RPE cells engineered to express eGFP were first transplanted into the vitreous and subretinal space of albino rats to facilitate localization of pigmented cells. Following intraocular transplantation, the transplanted pigmented cells can be readily identified in vivo (FIGS. 6A and 6B). After enucleation of the eye, removal of the cornea and lens, and separation of the retina shown in fig. 6B, the retina shows the primary graft and other scattered pigmented cells (fig. 6C). In tissue sections, grafts were present that included live pigmented transplanted cells that also expressed GFP (FIGS. 6D-6G). Transplanted cells can be found in the vitreous space, in the retina, along the injected channel, and in the subretinal space (fig. 6H, 6I, 6O, and 6P). Transplanted RPE cells also migrated from the subretinal graft and integrated into the RPE layer of the host rat (fig. 6J). In the grafts, tight junctions were formed that are characteristic of RPE cells, as shown by the expression of the tight junction marker ZO-1 in transplanted eGFP positive cells (fig. 6K-6N). Cells within the graft also maintained expression of RPE65, wilting protein and MiTF-a.

hESC-derived RPE cells survive, integrate and maintain the characteristics of differentiated RPE after transplantation into the subretinal space of RCS rats

The subjects of the transplantation experiments were performed in RCS rats, which revealed RPE and retinal degeneration due to mutations in the mertk gene, to examine whether delivery of hESC-derived RPE cells modulates the progression of the disease.

The transplanted pigmented cells can be readily identified in vivo in the eyes of RCS rats using standard fundus imaging systems (FIGS. 12A-12C). hESC-derived GFP expressing cells can be seen to emit fluorescence upon fluorescence excitation and use of an emission filter (fig. 12C). In the preparation of cups imaged ex vivo in a fluorescence microscope (fig. 12D-12E), large clusters of GFP positive cells under the retina (fig. 12D) and multiple discrete smaller clusters (fig. 12E) were visible.

Histological and immunohistochemical assessments confirmed macroscopic observations in vivo and ex vivo. Transplanted cells survived and integrated into the subretinal space, maintaining expression of proteins that were characteristic and often specific for mature RPE (fig. 13 and 14). It is important to note that there was no significant inflammatory or immune response and no tumor or teratoma was observed in more than 100 transplanted eyes. Hematoxylin and eosin (fig. 13A and 13B) stained sections show transplanted hESC-derived pigmented cells in subretinal and occasionally intraretinal locations, appearing in clusters or as isolated cells (arrows). Immunostaining with GFP (FIGS. 13C-13F) confirmed that these cells were indeed hESC-derived. The grafts were often quite large and scattered (FIGS. 13C and 13E), and pigmented cells co-expressing GFP were evident (FIGS. 13D and 13F).

Immunostaining showed that a large number of transplanted cells within the graft expressed proteins that characterized mature, differentiated RPE cells (fig. 14). This includes the expression of the RPE-specific markers RPE65 (FIGS. 14A-14E) and wilting protein (FIGS. 14F-14J). The cells are also able to form tight junctions (fig. 14K-14O), which are important functions of RPE cells and are critical for maintaining the blood-retinal barrier. The left-most column in each row shows a low magnification fluorescence image of a graft co-expressing GFP and associated markers. High magnification confocal images at each row show pigment (by Nomarski optics) at the single cell level as well as co-expression of GFP and different markers.

hESC-derived RPE cells provide functional and structural retinal rescue in dystrophic RCS rats

In the RCS rat model of retinal degeneration, retinal function is often severely impaired by 2-3 months of age. Structurally, a corresponding loss and thinning of the outer nuclear layer of the retina (ONL) occurs, which often drops below the 1-2 rows of photoreceptor nuclei at this age. In eyes transplanted with hESC-derived RPE cells, the video network membrane image recordings showed significant relative preservation of retinal function compared to control untreated or medium injected eyes (fig. 7 and 15).

Representative ERG responses to a series of increasing intensity white flashes under Dark Adaptation (DA) conditions are shown in the transplanted eye (FIG. 15A) and its cognate control eye (FIG. 15B). Figure 15C shows a significant difference in mean amplitude between transplanted eyes and control eyes of different groups. At the highest intensity, the mean DAb amplitude was 283.3 ± 37.5 (mean ± SEM; n ═ 13) in RPE transplanted eyes, compared to 158.5 ± 18.1(n ═ 13, p <0.01) in the untreated control eyes and 89.9 ± 14.4(n ═ 5, p <0.01) in media injected eyes of the same species. It is important to note that there was a tendency for better preservation of retinal function following transplantation of activin a-treated RPE cells (figure 15) compared to the rescue effect achieved following transplantation of RPE cells derived in the absence of activin a (figure 7).

Qualitative and quantitative assessment of retinal structure confirmed the functional findings (figure 16). Relative preservation of the photoreceptor (ONL) layer and internal and external photoreceptor segments (IS + OS) was observed near the subretinal RPE graft as compared to regions distant from the graft (both examples are shown in fig. 16A, 16B). Overall retinal thickness (fig. 16C) and ONL and IS + OS thickness (fig. 16D) were significantly increased near the hESC-derived RPE graft compared to regions distant from the graft (grey bars) (black bars, mean ± SEM, n ═ 7). This type of structural rescue was only observed at the subretinal and deep intraretinal grafts, but not when the graft was only intravitreal (not shown).

Uptake of rhodopsin in vivo by transplanted hESC-derived activin A-treated RPE cells

One key function of healthy RPE is to ingest and recycle shed photoreceptor outer fragments as part of the photoreceptor renewal process. Confocal images of subretinal transplanted RPE cells showed co-localization of pigment, GFP, RPE65 and rhodopsin within the same single cell. This indicates that the transplanted cells have the phenotype of mature RPE and are capable of uptake of the shed outer fragment (which contains rhodopsin). Note that native RPE cells of RCS rats expressed RPE65 (fig. 17C, arrows), but did not express GFP (fig. 17D, arrows) and contained minimal amounts of rhodopsin (fig. 17B, 17E).

The above results thus provide evidence that the obtained hSC-derived RPE cells can be used for transplantation in vivo to provide RPE cells that are essentially fully functional in the eye by culturing in a culture system comprising a member of the TGF β superfamily, and preferably in the presence of NA.

All references to "human embryonic stem cells" in this specification refer to commercially available human embryonic stem cells.

Claims (7)

1. A method of promoting directed differentiation of commercially available human embryonic stem cells into Retinal Pigment Epithelium (RPE) fates, the method comprising:

a. culturing commercially available human embryonic stem cells in medium supplemented with Nicotinamide (NA) to produce differentiating cells; and then

b. Culturing said differentiating cells in a medium supplemented with activin A under conditions that induce said differentiating cells to differentiate into RPE cells.

2. The method of claim 1, wherein said culturing in said medium supplemented with nicotinamide is performed for at least 2 days, followed by culturing in said medium supplemented with activin A.

3. The method of claim 1, wherein the culturing is performed as a suspension culture.

4. The method of claim 1, wherein said differentiating cells are cultured in said medium supplemented with nicotinamide for 2 weeks.

5. The method of claim 1, wherein said medium supplemented with activin A is further supplemented with nicotinamide.

6. The method of claim 1, wherein the concentration of nicotinamide is 10 mM.

7. The method of claim 1, wherein the concentration of activin A is 20-180 ng/ml.