Protective Effects of Human iPS-Derived Retinal Pigment
Epithelium Cell Transplantation in the Retinal Dystrophic
Amanda-Jayne Carr1.*, Anthony A. Vugler1., Sherry T. Hikita2., Jean M. Lawrence1., Carlos Gias1, Li Li
Chen1, David E. Buchholz2, Ahmad Ahmado1, Ma’ayan Semo1, Matthew J. K. Smart1, Shazeen Hasan1,
Lyndon da Cruz4, Lincoln V. Johnson2,3, Dennis O. Clegg2,3, Pete J. Coffey1
1Department of Ocular Biology and Therapeutics, Institute of Ophthalmology, University College London, London, United Kingdom, 2Center for Stem Cell Biology and
Engineering, Department of Molecular, Cellular and Developmental Biology, University of California Santa Barbara, Santa Barbara, California, United States of America,
3Center for the Study of Macular Degeneration, University of California Santa Barbara, Santa Barbara, California, United States of America, 4Department of Vitreoretinal
Surgery, Moorfields Eye Hospital, London, United Kingdom
Transformation of somatic cells with a set of embryonic transcription factors produces cells with the pluripotent properties
of embryonic stem cells (ESCs). These induced pluripotent stem (iPS) cells have the potential to differentiate into any cell
type, making them a potential source from which to produce cells as a therapeutic platform for the treatment of a wide
range of diseases. In many forms of human retinal disease, including age-related macular degeneration (AMD), the
underlying pathogenesis resides within the support cells of the retina, the retinal pigment epithelium (RPE). As a monolayer
of cells critical to photoreceptor function and survival, the RPE is an ideally accessible target for cellular therapy. Here we
report the differentiation of human iPS cells into RPE. We found that differentiated iPS-RPE cells were morphologically
similar to, and expressed numerous markers of developing and mature RPE cells. iPS-RPE are capable of phagocytosing
photoreceptor material, in vitro and in vivo following transplantation into the Royal College of Surgeons (RCS) dystrophic
rat. Our results demonstrate that iPS cells can be differentiated into functional iPS-RPE and that transplantation of these cells
can facilitate the short-term maintenance of photoreceptors through phagocytosis of photoreceptor outer segments. Long-
term visual function is maintained in this model of retinal disease even though the xenografted cells are eventually lost,
suggesting a secondary protective host cellular response. These findings have identified an alternative source of
replacement tissue for use in human retinal cellular therapies, and provide a new in vitro cellular model system in which to
study RPE diseases affecting human patients.
Citation: Carr A-J, Vugler AA, Hikita ST, Lawrence JM, Gias C, et al. (2009) Protective Effects of Human iPS-Derived Retinal Pigment Epithelium Cell Transplantation
in the Retinal Dystrophic Rat. PLoS ONE 4(12): e8152. doi:10.1371/journal.pone.0008152
Editor: Karl-Wilhelm Koch, University of Oldenburg, Germany
Received September 2, 2009; Accepted November 6, 2009; Published December 3, 2009
Copyright: ? 2009 Carr et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted
use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by funding from the London Project to Cure Blindness and The California Institute for Regenerative Medicine. The funders
had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: firstname.lastname@example.org
. These authors contributed equally to this work.
The retinal pigment epithelium (RPE) is a monolayer of cells,
residing at the back of the eye between Bruch’s membrane and
the retina, which is essential for photoreceptor function and
survival. Dysfunction and death of RPE has been observed in
various human degenerative diseases that lead to blindness,
including one of the leading causes of blindness in the western
world, aged-related macular degeneration (AMD). The limited
benefit of existing clinical and surgical interventions for these
diseases has led to increased interest in the development of a
cell-based transplantation therapy. Various cell types have been
examined for use in RPE cell replacement including immortal-
ized cell lines, such as the human RPE cell line, ARPE19,
sheets of adult RPE, foetal RPE, RPE derived from human
embryonic stem cells (HESC-RPE)[5–10] and many non-RPE
Current methods, producing stem cells from adult somatic cells,
offer an alternative cell source for transplantation. Induced
pluripotent stem (iPS) cells are morphologically identical to
embryonic stem cells, display similar gene expression profiles
and epigenetic status, and have the potential to form any cell in the
body [16–18]. iPS cells have been employed to generate cells for
the treatment of various diseases including diabetes, cardiovascular
disease, sickle cell anaemia, Parkinson’s disease and haemophilia
[19–23]. Meyer et al 2009 have recently shown that iPS cells
can be differentiated towards retinal cell types whilst a paper by
Buchholz et al 2009 has shown that human iPS cells can be
differentiated into retinal pigment epithelial cells which display
functionality in vitro. As yet it is unknown whether iPS cells can
differentiate into functional replacement cells for use in the
treatment of progressive diseases specific to the visual system. Here
we examine the potential of human iPS cells to differentiate into
fully characterized RPE cells (iPS-RPE). Furthermore we analyse
PLoS ONE | www.plosone.org1December 2009 | Volume 4 | Issue 12 | e8152
their functionality in vitro, and in vivo after transplantation of iPS-
RPE into the dystrophic RCS rat: a model of retinal dystrophy
where the primary defect, originating in RPE cells , leads to
blindness as a consequence of rod and cone photoreceptor
Materials and Methods
Derivation of iPS-RPE Cells and Cell Culture
The human induced pluripotent stem cell clone, iPS(IMR90)-
3, was passaged onto Mitomycin-C inactivated mouse
embryonic feeder cells with DMEM/F12 culture medium
containing 20% Knock-Out Serum Replacement, 0.1 mM non-
essential amino acids, 0.1 mM b-mercaptoethanol and 100 ng/ml
zebrafish basic fibroblast growth factor (zfbFGF) on a 6-well plate.
Cells were cultured at 37uC in 5% CO2for 6 days after which
zfbFGF was omitted to facilitate spontaneous iPS cell differenti-
ation. Pigmented colonies were observed within 4 weeks and
allowed to expand for a further 14 weeks, with media changes
every 2–3 days. Pigmented cells were enriched by manual
dissection of expanded colonies followed by dissociation in
0.05% Trypsin-EDTA. Cells were seeded at a density of
1.26104cells/cm2onto gelatin-coated plates with Human foetal
RPE medium containing a-MEM, 1 x N1 supplement, 1 x
Non-essential amino acid solution, 250 mg/ml taurine, 13 ng/ml
Triiodo thyronin (Sigma-Aldrich, Gillingham, UK), 20 ng/ml
Hydrocortisone (Sigma), 2mM L-glutamine (Invitrogen, Paisley,
UK), and 15% Hyclone heat-inactivated foetal bovine serum
(Thermo Scientific, Northumberland, UK), which was replaced
daily. Upon cells reaching confluency the serum concentration of
the medium was reduced to 5% and the media replenished twice
weekly. Subsequent passages were performed using Trypsin-
EDTA dissociation. Cells were then plated onto gelatin-coated
flasks, dishes and transwell membranes.
Characterization of Cells
were fixed for 30 min with 4% paraformaldehyde in 0.1 M
phosphate buffer. Sheets of cells were washed and scraped off the
dish using a cell scraper, cryoprotected in 30% sucrose in PBS and
rapidly frozen in OCT (Tissue Tec H). Sections (14 mm) were cut
onto charged glass slides (VWR). Cells on dishes, or sections were
blocked and incubated with appropriate combinations of primary
antibodies as described previously. The following primary
mouse monoclonal antibodies were used: MITF (1:30, Abeam),
RLBP1 (CRALBP, 1:1000, Affinity Bio reagents); BEST1 (1:1000,
Millipore); RPE65 (1:500, Millipore); PMEL17 (1:100 Dako);
KRT8 (1:2000, Millipore); Na+/K+ATPase (ATP1B1 - 1:100,
Abcam). Primary rabbit polyclonals used were OTX1/2 (1:500,
Millipore); PAX6 (1:300, Covance); Ki67 (1:2000, Vector labs);
ZO1 (1:50 Zymed) and Type IV Collagen (1:100, Morwell
Diagnostic Biosciences). Secondary antibodies were donkey anti-
ImmunoResearch Labs Inc.). Nuclei were visualised with DAPI
(1:5000, 496-diamindino-2-phenylindole dihydrochloride, Sigma).
Specificity of staining was tested by omission of the primary
antibodies. Images were obtained using a Zeiss confocal
microscope with Nomarski optics and analysed with LSM Image
ImageJ processing program to produce movies.
iPS-RPE cells were grown in gelatin-
coated transwells and then fixed in Karnovsky’s fixative (1%
paraformaldehyde and 3% glutaraldehyde in 0.1 M cacodylate
buffer). Cells plus membrane were excised from the transwell and
iPS-RPE cells or sheets of cells
or TRITC (1:200,Jackson
post-fixed in 1% osmium tetroxide. After dehydration through a
graded series of alcohols and epoxypropane, cells were transferred
to resin (Araldite Cy212, Agar Scientific) and polymerized at
60uC. Ultrathin sections were stained with uranyl acetate and lead
citrate prior to examination in a Jeol 1010 electron microscope.
Reverse transcription-Polymerase Chain Reaction (RT-
RNA was extracted from iPS-RPE cells using TriZol
reagent (Invitrogen) and DNA removed using RQ1-RNase Free
DNase. cDNA was then synthesized from 3 mg of RNA using
Superscript III Reverse transcriptase (Invitrogen). Gene expression
was analysed by amplifying 1 ml of cDNA synthesis product in a
PCR MastercyclerH (Eppendorf, Cambridge, UK) using Go Taq
Polymerase (Promega) in a reaction containing 0.2 mM of gene-
specific primers (Eurofins MWG Operon, Ersberg, Germany). For
primer sequence and annealing temperature (Ta) see Table S1.
The PCR cycle parameters consisted of an initial denaturation at
95uC for 2 min followed by 35 cycles of denaturation at 95uC for
30 s, annealing at Ta uC for 30 s, and elongation at 72uC for 30 s.
PCR was completed with a final elongation step at 72uC for
5 min. PCR products were resolved on a 2% agarose gel alongside
a 100 bp DNA ladder (Promega).
Total RNA was isolated from iPS, iPS-
RPE and foetal RPE cell cultures using the Qiagen RNeasy Mini
Kit (Qiagen, Valencia, CA). Contaminating genomic DNA was
digested using RQ1 RNase-free DNase (Promega, Madison, WI)
and the RNA purified again with the RNeasy Kit. cDNA was
synthesized from 1 mg of RNA using the iScript cDNA Synthesis
Kit (Bio-Rad, Hercules, CA). Quantitative real-time PCR was
carried out on a Bio-Rad MyIQ Single Color Real-Time PCR
Detection System using the SYBR Green method. Triplicate 20 ml
reactions were run in a 96 well plate with half of the cDNA
synthesis reaction used per plate. Primer specificity was assessed by
melting temperature analysis, gel electrophoresis and direct
sequencing (Iowa State DNA Facility, Ames, IA). Data was
normalized to the geometric mean of glyceraldehyde phosphate
dehydrogenase (GAPDH), peptidylprolyl isomerase A (PPIA),
hydroxymethylbilane synthase (HMBS) and glucose phosphate
isomerase (GPI). For primer sequence and annealing temperature
(Ta) see Table S1.
iPS-RPE cells were collected on ice in lysis
buffer (10 mM HEPES, 1% Triton, 150 mM KCl, 1 mM PMSF,
10 ng/ml leupeptin, 1 mM DTT, 50 ng/ml aprotinin, 10 mM
NaF, and 100 mM sodium vanadate) and incubated for 30 min on
a tube rotator at 4uC. Cell debris was removed from the sample by
centrifugation at 13,000 rpm for 30 min at 4uC and the
supernatant diluted in sample buffer. Proteins were denatured at
95uC for 5 min, separated on a SDS-PAGE gel alongside a Dual
Color Precision Plus Protein Standard (Biorad) and transferred to
Hybond PVDF membrane (GE Healthcare Life Sciences,
Buckinghamshire UK). Membranes were blocked for 2 hours in
10% milk in PBS-0.05% Tween-20 and incubated overnight in
10% milk containing primary antibodies raised in Mouse: RPE65
BioReagents, CO, USA); KRT8 (1:2000, Chemicon); PEDF
(1:500, Chemicon); PMEL17 (1:100, Dako); Rabbit: MERTK
(1:500, Abcam); FAK (0.1 mg/ml, Stratech) and MITF (1:500,
Chemicon). Membranes were washed in PBS-0.05% Tween-20,
incubated for an hour with secondary HRP-conjugated antibodies,
then washed and incubated in LumiLight western blotting solution
(Roche Products Ltd., Welwyn Garden City, UK). Proteins were
detected by exposure to autoradiographic film.
In vitro phagocytosis assay.
(POS) were isolated from freshly slaughtered porcine eyes using a
continuous sucrose gradient as previously described[8,32]. POS
Photoreceptor outer segments
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were then labelled with AlexaFluor H 488 Dye (Invitrogen) in 0.1
M sodium bicarbonate/5% sucrose in a light-tight microcentrifuge
tube for 1 h at room temperature. Labelled outer segments were
washed, resuspended in human foetal RPE medium and seeded
onto iPS-RPE cells cultured on gelatin-coated 35 mm dishes. Cells
were incubated at 37uC in 5% CO2 for 20 h. External
fluorescence was removed by treatment with trypan blue for
10 min. Cells were then washed in PBS and processed for
immunocytochemistry. iPS-RPE were also incubated with porcine
retina explants in an in vitro preparation previously described.
Briefly, iPS-RPE were cultured on gelatin-coated culture plate
inserts. Fresh porcine retinal tissue was dissected, orientated on a
0.45 mm filter with the photoreceptor cell surface uppermost and
placed on top of the iPS-RPE monolayer so that outer segments
were adjacent to the RPE. The explant model was incubated in
Human foetal RPE medium at 37uC in 5% CO2for 3 or 12 hours.
The iPS-RPE plus retina was fixed and processed for electron
microscopy as above.
dystrophic (rdy2/p+) and non-dystrophic control, pigmented
(rdy+/p+) Royal College of Surgeons (RCS) rats, which were
maintained in a 12 h light/dark cycle. Food and water were given
ad libitum and all procedures were carried out in accordance with
the UK Home Office regulations under the Animals (Scientific
procedures) Act 1986. The water contained 210 mg/l ciclosporin
(Sandoz, Camberley, UK) and was given to all animals
anaesthetized with a mixture of medetomidine hydrochloride
and ketamine. iPS-RPE cells were suspended in a-MEM (Sigma)
at a density of 56104cells per microlitre and 2 ml were injected
into the subretinal space of the left eye (N=10) between the host
RPE and photoreceptor cells, using a 30-gauge needle attached to
a Hamilton syringe. Control sham injections containing an
equivalent volume of a-MEM medium only were injected into
the dorsal subretinal space in an identical manner (N=6). Four
further dystrophic and 4 non-dystrophic (normal control) rats
remained as unoperated controls. Animals were tested for
visual function 13 weeks post-transplantation (16 weeks of age),
a time point where sham surgery no longer exerts an effect
on photoreceptor rescue. Termination was then achieved
by anaesthetic overdose and transcardiac perfusion of PBS
followed by 4% paraformaldehyde. Two of the iPS-RPE grafted
animals were sacrificed 8 days post-surgery for intermediate
anatomical assessment. Following perfusions, eyes were either
immunocytochemistry above or the retina was removed intact as
a whole-mount preparation.
Functional assessment and anatomy.
was assessed in two ways: firstly by optokinetic testing whereby an
animal that can see will involuntarily move its head in response to
a moving stimulus, and secondly, by examining light-induced c-
Fos activation in the host retina. The induction of c-Fos expression
in the inner nuclear layer (INL) and ganglion cell layer (GCL) of
the retina in response to light is an indication of functioning retinal
circuitry. The location of transplanted human cells, their
expression profile and ability to phagocytose rod photoreceptor
material was examined in vivo using immunohistochemistry.
We used the head-tracking response to
moving vertical lines to examine visual acuity in the RCS rat 13
weeks following surgery. The method for assessing head-tracking
(optokinetic response) has been reported previously. Briefly,
animals were placed into a circular container and exposed to a
rotating stimulus consisting of vertical black and white lines of
varying widths, subtending the spatial frequencies of 0.312, 0.25,
and 0.5 cycles/degree which rotate clockwise (to test acuity in the
left eye) or anticlockwise (to test acuity in the right eye). Each eye
was tested twice over a 60 second period. Animals that could see
made a well-defined ‘‘head-tracking’’ movement, following the
moving vertical lines. Testing was performed over three
consecutive days and the results were videotaped. The time
spent head-tracking over each 60 second period was measured
blind off-line by a single observer.
Head-tracking data was analysed using
SPSS, version 12.0.1. Two-way repeated-measures ANOVA was
performed to compare the optokinetic response between the
clockwise and counterclockwise directions as a function of spatial
frequency. Post hoc t-tests using a Bonferroni correction were used
to establish the spatial frequencies at which the difference between
both directions was significantly different. The effect of cell
transplantation treatment on the optokinetic response was
compared as a function of spatial frequency by using two-way
ANOVA with treatment as an independent-measures factor and
spatial frequency as a repeated-measures factor. Data are shown as
mean 6 SEM and P,0.05 was considered to be statistically
The expression of the immediate early
gene c-Fos was used to assess integrity of retinal function following
animals were dark-adapted overnight and then either sacrificed
in darkness (using a dim red light) or following 90 minutes of
constant illumination (white light at 250 mW/cm2). Retinal tissue
(whole-mounts or sections) were stained with rabbit anti-c-Fos
Immunohistochemistry to identify transplanted human
The antibodies listed above in the immunocytochemistry
section were also used on eye sections together with a cocktail of
antibodies generated against human-specific markers (HSM) to
identify human cells (the Oka blood group antigen, mouse TRA-1-
85, 1:10 (a kind gift from Peter Andrews, University of Sheffield,
Sheffield, UK) together with mouse human nuclear antigen,
1:1000, Millipore). Additionally, human-specific mouse anti-Ki67
(1:50, Dako) revealed any human cells in active phases of the cell
cycle. Where mouse monoclonal antibodies were used to probe for
RPE specific markers in iPS-RPE cells in vivo (e.g. anti-RLBP1 and
anti-PMEL17), the human identity of target cells was confirmed by
subsequent re-probing of sections with human specific markers
(HSM) as described previously. Immunocyctochemistry was
also used to assess phagocytosis of host outer segments in vivo.
Phagocytosis of photoreceptor outer segments was defined by the
inclusion of rhodopsin-labelled material within the TRA-1-85-
bordered intracellular compartment of grafted human iPS-RPE
Differentiation of iPS into RPE Cells
iPS cells, derived from the IMR-90 human foetal lung
fibroblast cell line, were cultured in stem cell medium lacking
bFGF to encourage the spontaneous differentiation of cells
(Fig. 1A). This led to the appearance of pigmented colonies
within 4 weeks. Colonies were allowed to expand for a further 14
weeks before we enriched for pigmented cells by manual
dissection of expanded pigmented colonies followed by cell
dissociation. Cells were cultured in human foetal RPE medi-
um where, within 13 days, they formed a pigmented
monolayer with characteristic RPE cobblestone appearance
similar to that observed in human post-mortem RPE sheets and
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HESC-derived RPE cells (Fig. 1B). The enrichment method
employed produced an almost homogeneous population of iPS-
RPE cells at passage 2 in a 25 cm2tissue culture flask (Fig. 1C)
with no evidence of cell multi-layering. The appearance of dome-
shaped blisters suggested that fluid transport from the apical to
basal surface of cells might be occurring.
Characterisation of iPS-RPE Cells
The appearance of RPE cell morphology was associated with
the expression of a panel of classic RPE genes and proteins
required for retinoid recycling (RPE65, LRAT, RLBP1), phago-
cytosis (FAK and MERTK) and melanogenesis (Tyrosinase,
PMEL17 and MITF). Cells also expressed the anti-neovascular
agent/neurotrophic factor PEDF and KRT8, an epithelial keratin
associated with RPE cell proliferation (Fig. 1D and 1E). The
increase of RPE cell markers in iPS-RPE was accompanied by the
down-regulation of the iPS reprogramming molecules OCT4,
SOX2 and NANOG expression (Fig. 1F and 1G), indicative of
differentiation away from the iPS cell phenotype.
Ultrastructurally, melanosomes, responsible for the pigmen-
tation normally observed in RPE cells, were clearly observed
within iPS-RPE (Fig. 2A and 2B). Akin to human RPE, the cells
were highly polarized with basal nuclei, apical microvilli and
adherens junctions (Fig. 2C and 2D). Coated pits, associated
with endocytosis, were also present on iPS-RPE cells (Fig. 2E).
Cells had secreted their own basal lamina (Fig. 2F), which was
positive for the extracellular matrix protein, Collagen IV
(COL4) (Fig. 3). In sections through the iPS-RPE monolayer
we observed apical expression of Na+/K+ATPase (ATP1B1).
PAX6, OTX2 and MITF, transcription factors involved in RPE
cell development, were localised in the nucleus, whilst RLBP1,
PMEL17, and BEST1 were expressed cytoplasmically. Impor-
Figure 1. Human induced pluripotent stem cells differentiate into retinal pigment epithelial cells. (A) Photomicrographs showing
undifferentiated (left) and differentiating iPS cells (right). (B) iPS-RPE cells form a pigmented monolayer in culture with typical RPE cell cobblestone
appearance. (C) Comparison of pigmentation observed in T25 flasks of confluent ARPE-19 cells and iPS-RPE cells. (D) PCR amplification of classic RPE
cell markers in iPS-RPE cells. A 100 bp DNA ladder was applied to the gel as an amplicon size reference. (E) Western blot analysis of iPS-RPE protein
expression using antibodies against a panel of RPE cell markers. A protein standard was used on each Western blot to determine the correct
molecular weight of proteins. (F) Quantitative PCR analysis of RPE and iPS gene expression in iPS, iPS-RPE and foetal RPE cDNAs. (G) iPS
reprogramming proteins are reduced after differentiation to iPS-RPE. OCT4 and SOX2 expression is undetectable in iPS-RPE cells. Combined confocal
and Nomarski image of iPS-RPE cells are shown on the left with confocal channels to the right. OCT4 and SOX2 (red channel) and DAPI stained nuclei
are blue. Scale bars: A, 20 mm; B and G, 50 mm.
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tantly, differentiated cells were negative for RPE cell de-
differentiation marker, KRT8 and a marker of the active
phase of cell cycle, Ki67, indicating cells were no longer
proliferative (Fig. 3).
Functional Assessment of iPS-RPE In Vitro
We used phagocytosis assays to assess the functional potential
of cells. iPS-RPE were able to phagocytose fluorescently labelled
porcine photoreceptor outer segments (POS) in co-culture.
Confocal microscopy through cells labelled with the apical
marker ATP1B1 demonstrated that POS are internalized by
iPS-RPE (Fig. 4A and Movie S1). iPS-RPE cells were also able to
ingest and digest photoreceptor material from porcine retinal
explants. The apical surface of cells envelop the POS (Fig. 4Bi)
and internalized coated pits are seen after 3 hours co-culture
(Fig. 4Bii), with end-stage lipid deposits observed after 12 hours
In Vivo Assessment of iPS-RPE Cell Morphology and
Phagocytosis After Transplantation into the RCS Rat
In order to test the therapeutic efficacy of iPS-RPE, we
transplanted cells into the subretinal space of RCS dystrophic
rats (Fig. 5A). After 20 h we observed a layer of pigmented cells
within the subretinal space of the RCS rat, which were Ki67
negative. The origin of the cells was confirmed by staining with
HSM (Fig. 5B). We compared the expression of RPE cell
markers in transplanted cells in vivo against cells re-plated and
cultured in parallel. After 8 days in culture, the re-plated iPS-
RPE cells were fixed and stained for RPE-associated markers
(Fig. 5C). Cells maintained the expression profile they had
displayed prior to grafting and had largely exited active phases
of the cell cycle (Ki67 negative). At the comparable time point in
vivo, iPS-RPE cells were observed in the subretinal space and
could be clearly identified with the aid of HSM staining
Figure 2. Human iPS-RPE cells are polarized and display classic
RPE cell morphology. (A) Electron micrograph of an iPS-RPE cell
monolayer. Human iPS-RPE are pigmented cuboidal epithelial cells with
cytoplasmic polarization. Indicated are apical microvilli (AMv), melanin-
containing melanosomes (red arrows), the basal nucleus (N), desmo-
somes (white arrows) and basal lamina (black arrows). (B) Densely
packed melanosomes: stages II, III and IV of melanosome maturation
are labelled. (C) Microvilli (AMv) extend out from the apical surface of
iPS-RPE. (D) Adherens junctions (white arrows) between cells are in the
apical portion of the cell, whilst the nucleus (N) is basal. (E) Coated pits
(asterisk) are found throughout the plasma membrane. (F) iPS-RPE
produce their own basal lamina (indicated by black arrows). Scale bars:
A, 2 mm; B-F, 1 mm.
Figure 3. Immunocytochemical localization of RPE cell specific
proteins in sectioned sheets of iPS-RPE cells. The left column
shows combined Nomarski and confocal images of the pigmented iPS-
RPE cell sheet sections; adjacent are images of immunolabelling only.
Protein staining is indicated by the colour of the text (red or green) and
DAPI stained nuclei are blue. Scale bars: All 50 mm.
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(Fig. 5D). At 8 days iPS-RPE were no longer present as a layer,
and instead formed cell boluses. However, the transplanted cells
maintained expression of RPE markers such as RLBP1 and
OTX2, with only occasional cells positive for Ki67. The in vivo
expression of RPE65 was not detectable by immunocytochem-
istry. Grafted human cells expressed PMEL17 in vivo and could
down-regulate Pax6 in the subretinal space, a hallmark of
terminally differentiated RPE (Fig. 5D). The pattern of staining
displayed by transplanted iPS-RPE cells in vivo was similar to
that reported recently for HESC-RPE grafted into the
dystrophic RCS rat retina. Importantly, akin to HESC-
RPE in vivo, cells at the outside edge of the iPS-RPE cell bolus
could phagocytose photoreceptor outer segments from the RCS
rat,as indicated by the presence of rhodopsin-positive
photoreceptor material within the cellular membrane of iPS-
RPE labelled with HSM (Fig. 5E).
Figure 4. iPS-RPE cells phagocytose photoreceptor outer
segment material in vitro. (A) Confocal images showing phagocy-
tosis of isolated FITC-labelled porcine photoreceptor outer segments
(POS – green) by iPS-RPE in culture. The nuclei are stained with DAPI
(blue). Internalization of POS is observed in a single optical y-axis
projection (,1 mm) of pigmented iPS-RPE cells labelled with the apical
cell surface marker ATP1B1 (red). (B- i) Electron microscopy image of a
porcine photoreceptor outer segment (POS) adjacent to an iPS-RPE cell
following 3 hours co-culture with a porcine retina explant. iPS-RPE
apical microvilli (red arrows) and coated vesicles (white arrows) are
observed proximal to the porcine POS. (ii) Internalized coated pits
(black arrow and enlarged inset in blue box) are seen within the
cytoplasm of iPS-RPE cells co-cultured with POS. (iii) Lipid deposits (L), a
sign of late stage POS phagocytosis, are observed within the iPS-RPE
cytoplasm after 12 h co-culture. Scale bars: A, Upper panel 20 mm and
lower panel 10 mm; i, iii, 2 mm and ii, 1 mm.
Figure 5. iPS-RPE maintain RPE cell markers and phagocytose
host photoreceptor outer segment material following trans-
plantation into the subretinal space of dystrophic RCS rats. (A)
A schematic of retinal cell organisation. The nuclear layers are
indicated. iPS-RPE cells were injected into the subretinal space
between the host RPE and photoreceptor cells (B) A layer of iPS-RPE
cells in the subretinal space of the dystrophic RCS rat 20 hours
following transplantation. (C) Retention of RPE markers by iPS-RPE cells
in vitro after dissociation, re-plating and culturing for 8 days. Protein
staining is indicated by the text colour. (D) Expression of the same RPE
cell markers is maintained in vivo by iPS-RPE (white arrows) 8 days
following transplantation into the subretinal space of the RCS rat. (E)
Rhodopsin-positive material (red) is present within the cell membrane
of human specific marker (HSM)-labelled iPS-RPE (green) 8 days post-
transplantation and in the tips of the host outer segment (OS) layer.
DAPI (blue) stains nuclei. Indicated are the outer and inner nuclear
layer (ONL and INL respectively) of the retina. Scale bars: B-D, 50 mm; E,
50 mm and 20 mm in magnified view.
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Assessment of Visual Function in the Dystrophic RCS Rat
Following iPS-RPE Transplant
We used the head-tracking response to assess the visual function
of RCS dystrophic rats 13 weeks after receiving a subretinal iPS-
RPE transplantation in one eye only (Fig. 6 and Movies S2 and
S3). Preservation of the higher spatial frequency (0.5 c/d)
monocular optokinetic head-tracking response was associated with
the iPS-RPE transplanted eye when compared with the sham-
injected eye (ANOVA, F (1,14)=1.753, P,0.05; Post hoc test at
0.5 c/d p=0.01) and non-transplanted (ANOVA, F(1,9)=48.8,
P,0.001. Post hoc t-test; t(9)=1.61, p,0.05 for all spatial
frequencies). Conservation of visual acuity in the iPS-RPE
transplanted eyes was associated with the preservation of
photoreceptors in the host outer nuclear layer (ONL – Fig. 7A),
identified by the expression of rhodopsin in the outer segments of
photoreceptors (Fig. 7A inset, Dystrophic+transplant). At this age,
in the RCS dystrophic rat, the ONL is normally reduced to a
single layer of cells with an autofluorescent debris zone (Fig. 7A
inset, Dystrophic) [9,33].
Survival of iPS-RPE In Vivo After Transplantation into the
Dystrophic RCS Rat
Even though the ONL was preserved 13 week post-transplan-
tation, there was little evidence of surviving iPS-RPE cells within
the subretinal space by this time. Occasional HSM-positive
material could be detected in the subretinal space (Fig. 7B), but
the lack of a defined membrane and the absence of DAPI staining
suggest that these cells were not viable. However, we did find host
cells positive for the monocyte/macrophage marker CD68 within
the sub-retinal graft site, which did not stain for HSM. At 8 days
these cells had a clear appearance (Fig. 8A), and rhodopsin-
positive material was found within the CD68-expressing cells
(Fig. 8B). At 13 weeks, two distinct CD68-positive cell populations
were observed, one type was small and non-pigmented (Fig. 8C)
and the other consisted of large highly pigmented cells (Fig. 8 D).
Both these cell types contained rhodopsin-positive material
To assess the topography of retinal function in grafted versus
non-grafted eyes we employed functional anatomy to look at the
activation of inter-neurons of the inner nuclear layer (INL). This
was achieved by examining the light-induced expression of c-Fos,
a member of the immediate early gene family. Light is known to
induce nuclear c-Fos expression in the INL of the mouse retina,
however this response is reduced in a mouse model of
photoreceptor degeneration. In the normal, non-dystrophic
rat retina, the ONL was present and there was clear activation of
c-Fos in the INL in response to light, with a complete absence of
staining in the dark (Fig. 9A). Only occasional foci of c-Fos positive
cells were detectable in the INL of the non-transplanted dystrophic
retina where the ONL consisted of a single layer. Following
transplantation of iPS-RPE cells, there was a clear preservation of
light-induced c-Fos activation in the INL beneath regions of
photoreceptor preservation and in the ganglion cell layer (GCL),
the output layer of the retina. The distribution of occasional light-
activated c-Fos cells observed in the GCL of non-transplanted
dystrophic rats matches that reported for intrinsically light-
responsive melanopsin ganglion cells. Importantly, in grafted
Figure 6. Preservation of visual function following iPS-RPE
transplantation into the RCS rat. (A) Preservation of optokinetic
head-tracking response to a rotating vertical stimulus in 16-week-old
RCS dystrophic rats following transplantation of iPS-RPE. Mean visual
acuity (6S.E.M.) of the transplanted eye versus control sham-injected
eye and non-transplanted dystrophic eye. Spatial frequency is indicated
in cycles per degree (c/d).
Figure 7. Preservation of the photoreceptor cell layers after
transplantation of iPS-RPE. (A) Extensive preservation of the nuclear
photoreceptor layers in the dorsal retina of the dystrophic RCS rat 13
weeks following transplantation of iPS-RPE cells (DAPI stained nuclei).
Inset shows higher resolution confocal images of photoreceptor cell
nuclear layers (DAPI blue) and rhodopsin expression (red) in the
dystrophic control (left inset) and dystrophic with iPS-RPE transplant
(right inset) RCS rat. (B) Confocal images showing HSM-positive staining
(green, indicated with white arrow) within the subretinal space 13
weeks post-transplantation. Scale bars: A 500 mm; inset, 50 mm; B
Transplantation of iPS-RPE
PLoS ONE | www.plosone.org7 December 2009 | Volume 4 | Issue 12 | e8152
animals, preservation of the ONL and light-induced c-Fos
expression in the INL and GCL was restricted to the dorsal
retina, the region where iPS-RPE cells had been injected (Fig. 9B).
The successful differentiation of iPS cells into RPE represents a
significant advance in the search for a potential cell source for the
treatment of human neural retinal diseases. iPS(IMR90)-3 cells
readily differentiate into RPE cells and the differentiation protocol
used in this paper is a highly efficient method of producing
multiple confluent flasks of highly enriched pigmented cells. An
efficient initial RPE differentiation protocol is essential for the
production of cells for use in any therapeutic application since
repeated passaging of RPE results in phenotypical and morpho-
logical changes associated with dedifferentiation[36,37]. The RPE
cells derived from iPS(IMR90)-3 cells have been well-character-
ized here and satisfy many of the known criteria of RPE cells,
including protein expression, cellular pigmentation and polariza-
tion. These properties are similar to those observed in cultured
HESC-derived RPE[5–7,9]. iPS-RPE also down-regulate the
embryonic transcription factors originally used to induce pluripo-
Figure 8. Macrophages are present in the subretinal space of the dystrophic RCS following transplantation of iPS-RPE cells. (A)
Coronal confocal projection showing confocal channels and Nomarski images of iPS-RPE transplanted eye containing non-pigmented CD68-positive
cells (green) 8 days following transplantation. The white box contains a CD68-positive cell with rhodopsin-positive cytoplasmic inclusions (red) which
is magnified and shown in serial confocal slices in B. (C) Co-labelling of non-pigmented CD68 and rhodopsin-positive cells at 13 weeks post-graft.
Note the presence of an intact outer segment and outer nuclear layer at this stage. (D) Large pigmented cells are observed in the subretinal space at
13 week post-graft. The white box indicates a CD68-positve cell containing rhodopsin-positive inclusions, shown magnified and in serial confocal
slices in (E). Scale bars: All 20 mm.
Transplantation of iPS-RPE
PLoS ONE | www.plosone.org8 December 2009 | Volume 4 | Issue 12 | e8152
tency from the somatic cell, suggesting differentiation away
from the initial iPS phenotype.
Critical functions of RPE cells include the maintenance of
photoreceptor cell integrity by phagocytosing debris shed by the
retina each day and epithelial transport of molecules and
metabolic waste. Functional properties of RPE cells are also
observed in iPS-RPE in vitro, including the transport of fluid across
cells, as indicated by the formation of blister-like domes in the
monolayer. Similar to HESC-derived RPE, iPS-RPE can
phagocytose photoreceptor outer segments from isolated prepara-
tions and porcine retina explants. A recent paper has also
described the in vitro phagocytic properties of iPS-RPE cells
derived using an alternative culture protocol , however, the in
vivo function of these cells has yet to be tested.
After 8 days in the sub-retinal space of the dystrophic RCS rat,
iPS-RPE cells are capable of phagocytosing host photoreceptor
outer segments. This evidence of in vivo phagocytosis is
characterised by the presence of an exclusive outer segment
marker, rhodopsin, within the cytoplasmic compartment of cells
labelled with HSM. These data suggest that human iPS-RPE cells
are able to contribute to host photoreceptor cell integrity by
removing retinal debris at this time-point.
iPS-RPE cells were transplanted into dystrophic RCS rat eyes at
three weeks of age, a time when retinal abnormalities are first
observed. The progression of retinal dystrophy in the RCS rat is
such that by 13 weeks post-graft most of the ONL has disappeared
 and the photoreceptor outer segment layer is reduced to a
debris zone, a finding we observed in dystrophic controls. The
preservation of these layers 13 weeks after iPS-RPE cell injection
suggests that the transplantation of these cells preserves retinal
structure. At this stage we also performed analysis of visual
function. Visual acuity tests showed that the performance of
animals receiving iPS-RPE transplants was significantly better
than sham-operated and control animals. We also identified the
presence of functional neuronal circuitry in the retina after iPS-
RPE transplantation as indicated by the light induced c-Fos
response in the INL and GCL of the neural retina. This is the first
time that that this assay has been used to ascertain retinal function
Figure 9. iPS-RPE transplantation preserves the light induced c-Fos response in the RCS dystrophic rat retina. (A) Age-matched non-
dystrophic control, dystrophic no-transplant control and dystrophic rats with iPS-RPE grafts were dark-adapted overnight and sacrificed in the dark or
after 90 min exposure to white light (250 mW/cm2). Coronal sections through the retina of dystrophic transplanted, dystrophic non-transplanted and
non-dystrophic control RCS rats showing the inner and outer nuclear layers (ONL and INL respectively) and the ganglion cell layer (GCL). Preservation
of ONL 13 weeks after iPS-RPE transplant correlates with preservation of the light-induced c-Fos expression (red) in coronal sections of the retina and
in representative dorsal whole-mount preparations of the inner nuclear and ganglion cell layers. Note absence of activity in darkness and responsivity
to light in the normal and transplanted eyes. c-Fos positive cells in the ganglion cell layer of unoperated RCS eyes match the distribution expected for
intrinsically light-responsive melanopsin-containing ganglion cells. DAPI-stained nuclei are shown in blue in the coronal section and the
autofluorescent debris zone (dz) is indicated. (B) Light-induced c-Fos activation in the transplanted eye of RCS rats is preferentially preserved in the
dorsal retina (the region of the transplant), corresponding with preservation of photoreceptors (ONL) by the iPS-RPE graft. No such preservation is
observed in the ventral retina of the transplanted animal. Scale bars: Coronal sections, 50 mm; whole-mount images of the inner nuclear and retinal
ganglion cell layers, 200 mm.
Transplantation of iPS-RPE
PLoS ONE | www.plosone.org9 December 2009 | Volume 4 | Issue 12 | e8152
after cellular therapy in a retinal degenerate animal. Importantly,
the preservation of light-induced c-Fos expression was restricted to
the dorsal retina, surrounding the region of iPS-RPE cell injection
only, suggesting preservation of localised neural activation
corresponding to the histology of rhodopsin expression. Previous
studies, using electroretinography to assess function following cell
transplantation into the RCS rat have only been able to
demonstrate global activity across the retina [6,41–43].
The absence of iPS-RPE cells in the subretinal space at the time
of functional assessment (13 weeks) indicates that the significant
benefits observed could not be wholly attributed to the donor cells.
Although all animals were maintained on an oral immunosup-
pression drug, ciclosporin, throughout the experiment, this was not
sufficient to sustain iPS-RPE cell survival. These findings are in
agreement with previous studies that show that xenografts can be
compromised[4,44] even after triple immune suppression.
Our analysis suggests that loss of transplanted cells is associated
with infiltration of the subretinal space by macrophages/microglia.
The large pigmented CD68-positive cells observed in the
subretinal space at 13 weeks are likely to be macrophages/
microglia filled with melanin[46,47] from the transplanted human
iPS-RPE cells. The presence of rhodopsin within the macrophag-
es/microglia could explain some of the behavioural and functional
benefits observed, since clearance of outer segment debris by these
cells in the subretinal space could also contribute to photoreceptor
cell survival. This conclusion has been implied previously in a
study which suggests that macrophage infiltration in response to
the trauma of retinal detachment after saline injection, contributes
to extend the longevity of photoreceptor cells in the RCS rat.
Our finding highlights a key aspect of cellular transplantation not
fully addressed in many short or long-term studies of RPE
transplantation: the host inflammatory/immune response to the
xenograft and its indirect role in the preservation of the retina.
Importantly, we show that the presence of pigmented cells within
the subretinal space does not necessarily reflect survival of
transplanted cells. We suggest correct identification of the origin
of these cells (using human specific markers, which define cell
membranes) is essential in order to distinguish viable donor cells
from host inflammatory cells which have engulfed transplanted
cells. Identification using pigmentation alone is not sufficient.
The increased visual acuity and outer nuclear layer preservation
observed in iPS-RPE injected animals could also be due to a
neuroprotective effect produced by the donor cells. RPE are
known to secrete neurotrophic growth factors such Glial cell
derived neurotrophic factor (GDNF), Brain derived neurotrophic
factor (BDNF), PEDF and bFGF. As these factors can
exert protective effects on retinal neurons[52–56] and neurons in
other neurodegenerative disease models [49,57,58], it seems likely
that these substances may contribute to the latent photoreceptor
cell survival we observe in the dystrophic retina. The fact that the
area of preservation extends beyond the borders of the graft
suggests that a neuroprotective effect may also be involved.
Release of growth factors can occur as a result of surgery but the
effects are of shorter duration.
The major limitation of this study is the rejection of human iPS-
RPE after transplantation into the RCS rat. RPE cells derived
from HESC show long-term viability in vivo, since in a previous
study we have shown that HESC-derived RPE can survive in the
RCS subretinal space for up to 10 weeks, whilst a recent study
has indicated survival of cells for up to 30 weeks. The
embryonic origin of HESC-derived RPE may reflect a more
immune privileged cell type in comparison to iPS-RPE, which
contributes to their longer-term survival after xenografting. HESC
have been shown to have reduced immunogenicity, expressing low
levels of MHC-I and MHC-II in both the undifferentiated and
dedifferentiated state, and to possess an adaptive mechanism to
immune responses[60–62]. As yet the immunogenic profile of iPS
or iPS-derived cells has not been described. Thus, the RCS rat
retina offers a useful model for the short-term analysis of iPS-RPE
cell function in vivo, but the donor cell loss due to host macrophage
infiltration of the xenograft indicates that additional modification
may be necessary to promote long-term donor cell survival in this
animal model system.
The major benefit of using iPS cells to treat AMD is that
developing a patient specific therapy may help to eliminate the
problems associated with immune rejection. Proof of concept for
the therapeutic use of a patient’s own iPS-derived RPE lies in
current clinical treatments for AMD. Although complicated,
surgical procedures such as the autologous transplantation of
peripheral RPE to the macular region[63–66] and macular
translocation, where the neural retina is detached and the fovea
relocated to a less diseased area of RPE[67–69], have been shown
to stabilize visual acuity in AMD patients. Consequently, although
iPS-RPE may still carry the same genetic defect responsible for
AMD in the patient, the fact that these cells have not been
diseased by age, like macular RPE, suggests that they could still be
used as a viable therapeutic. iPS cell therapy might also be useful
in patients with genetic diseases, such as Leber’s congenital
amaurosis where transplantation could be combined with gene
therapy to correct genetic defects inherent to the patients’ own
RPE cells. Alternatively iPS cells derived from a tissue-matched
healthy sibling may also be useful. iPS-RPE may also provide
a useful in vitro model system in which to study the pathogenesis of
human RPE-linked diseases and identify novel molecular/
biochemical therapeutic targets.
While this particular line of iPS-RPE cells could not be used as a
direct therapeutic due to viral insertions of pluripotency genes, the
recent advances in iPS cell reprogramming technology, including
the use of small molecules[71–73], piggyBac transposition[74,75],
non-integrating episomal vectors and manipulation of endog-
enous transcription factors should eliminate the risks associ-
ated with integration of stem cell genes into the genome.
Furthermore, the finding that blood cells can be used to derive
iPS cells may remove the need for invasive patient biopsies
required for the collection of somatic cells and accelerate the
ethical production of stem cell-derived tissue for therapeutic use.
Amplicon size is in base pairs (bp).
Found at: doi:10.1371/journal.pone.0008152.s001 (0.09 MB
Primer pairs used for RT-PCR and quantitative PCR.
outer segments (green) by iPS-RPE cells in vitro. The apical
surface of the RPE cells is identified by ATP1B1 (red).
Found at: doi:10.1371/journal.pone.0008152.s002 (0.50 MB
Phagocytosis of fluorescently labelled photoreceptor
head-tracking response is observed in the 16-week old dystrophic
RCS rat during optokinetic testing of the right, non-transplanted,
Found at: doi:10.1371/journal.pone.0008152.s003 (1.80 MB
Visual acuity testing of the non-transplanted eye. No
A head-tracking response is observed in the dystrophic RCS rat
Visual acuity testing of the iPS-RPE transplanted eye.
Transplantation of iPS-RPE
PLoS ONE | www.plosone.org 10 December 2009 | Volume 4 | Issue 12 | e8152
during optokinetic testing of the left eye, 13 weeks after iPS-RPE
Found at: doi:10.1371/journal.pone.0008152.s004 (1.90 MB
The authors would like to thank James A. Thomson for providing the
human iPS cells used in this study. We would also like to thank Peter
Munro and Robin Howes for helpful advice and technical assistance.
Conceived and designed the experiments: AJFC AV STH JML LdC LJ
DC PC. Performed the experiments: AJFC AV STH JML CG LLC DEB
AA MJKS SH. Analyzed the data: AJFC AV STH JML CG DEB MS DC.
Contributed reagents/materials/analysis tools: AJFC AV STH JML DEB
PC. Wrote the paper: AJFC AV JML.
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Transplantation of iPS-RPE
PLoS ONE | www.plosone.org 12 December 2009 | Volume 4 | Issue 12 | e8152