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(Investigative Ophthalmology and Visual Science. 2007;48:4321-4327.)
© 2007 by The Association for Research in Vision and Ophthalmology, Inc.
DOI:  10.1167/iovs.06-1015
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Endogenous Bone Marrow–Derived Cells Express Retinal Pigment Epithelium Cell Markers and Migrate to Focal Areas of RPE Damage

Yang Li,1 Pelin Atmaca-Sonmez,1 Carrie L. Schanie,2 Suzanne T. Ildstad,2 Henry J. Kaplan,1 and Volker Enzmann1,3

1From the Department of Ophthalmology and Visual Sciences and the 2Institute for Cellular Therapeutics, University of Louisville, Louisville, Kentucky; and the 3Department of Ophthalmology, Inselspital, University of Bern, Bern, Switzerland.


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
PURPOSE. The aim of the present study was to investigate whether bone marrow–derived cells (BMCs) can be induced to express retinal pigment epithelial (RPE) cell markers in vitro and can home to the site of RPE damage after mobilization and express markers of RPE lineage in vivo.

METHODS. Adult RPE cells were cocultured with green fluorescence protein (GFP)-labeled stem cell antigen-1 positive (Sca-1+) BMCs for 1, 2, and 3 weeks. Cell morphology and expression of RPE-specific markers and markers for other retinal cell types were studied. Using an animal model of sodium iodate (NaIO3)-induced RPE degeneration, BMCs were mobilized into the peripheral circulation by granulocyte-colony stimulating factor, flt3 ligand, or both. Immunocytochemistry was used to identify and characterize BMCs in the subretinal space in C57BL/6 wild-type (wt) mice and GFP chimeric mice.

RESULTS. In vitro, BMCs changed from round to flattened, polygonal cells and expressed cytokeratin, RPE65, and microphthalmia transcription factor (MITF) when cocultured in direct cell–cell contact with RPE. In vivo, BMCs were identified in the subretinal space as Sca-1+ or c-kit+ cells. They were also double labeled for GFP and RPE65 or MITF. These cells formed a monolayer on the Bruch membrane in focal areas of RPE damage.

CONCLUSIONS. Thus, it appears that BMCs, when mobilized into the peripheral circulation, can home to focal areas of RPE damage and express cell markers of RPE lineage. The use of endogenous BMCs to replace damaged retinal tissue opens new possibilities for cell replacement therapy in ophthalmology.

Age-related macular degeneration (AMD) is the leading cause of blindness in persons older than 65 in developed countries and results in severe and irreversible vision loss for more than 2 million Americans.1 2 Two distinct types of AMD are recognized, dry (nonexudative) and wet (exudative). In dry AMD, the more common form, progressive atrophy of the retinal pigment epithelium (RPE) occurs with subsequent loss of the choriocapillaris and photoreceptors.3 Wet AMD is frequently characterized by the formation of choroidal neovascularization (CNV) and is the cause of severe vision loss.4 No treatment is available to reproducibly reverse the degenerative process and restore vision in AMD. RPE transplantation, which seeks to replace RPE that has undergone geographic atrophy in dry AMD or that has been removed during submacular surgery for CNV in wet AMD, has not been successful to date.

During the past several years, stem cells have been investigated for their potential use in regenerative medicine.5 Adult stem cells, particularly those from the bone marrow (BM), have the capacity to transdifferentiate in response to in vivo and in vitro stimuli.6 Numerous studies have described the ability of bone marrow–derived cells (BMCs), including hematopoietic stem cells (HSCs), to cross lineage boundaries and to express tissue-specific proteins in different organs, including the liver,7 heart,8 brain,9 small intestine,10 skeletal muscle,11 bone,12 and vascular endothelium.13 Intravascular BMC transplantation leads to structural and functional repair in animal models of human diseases, such as hereditary liver disease,14 myocardial infarction,8 and kidney ischemia.15 Furthermore, several successful attempts to deliver exogenous BMCs directly to the site of injury have been reported.16 17 Therefore, it was initially presumed the repair seen in damaged host tissues after stem cell injection resulted from the incorporation and transdifferentiation of stem cells (SCs) at the sites of damage. This was supported in humans by the observations that transplantation of SCs from mobilized peripheral blood (PB) expressing the early hematopoietic CD34+ antigen led to the appearance of donor-derived hepatocytes, epithelial cells, and neurons.18 Similarly, human SCs from the BM contributed to the regeneration of infarcted myocardium.19 20 However, a number of studies have challenged this concept, providing evidence that SCs may instead incorporate into host tissues through fusion with host cells.21 22

Even if transplanted BMCs have the capacity to colonize different tissues, proliferate, and differentiate into cell lineages of the host organ, the cell transplantation procedure is a complex multistep process. Additionally, there is a risk of transmission of infectious agents and development of an allogeneic immune reaction to the transplanted histoincompatible tissue. Therefore, a noninvasive transplantation method is highly desirable. In vivo stem cell mobilization and expansion could supply new autologous cells for tissue repair without the use of invasive procedures. Mobilization of BMCs into the PB can be induced by two hematopoietic growth factors, granulocyte colony stimulating factor (G-CSF) and flt3 ligand (FL). G-CSF has been shown to mobilize HSCs in vivo.23 24 FL enhances the proliferation of HSCs in vitro and mobilizes HSCs in vivo.25 The mobilizing effect of G-CSF, in combination with the proliferation of HSCs induced by FL, has shown significant synergy in HSC mobilization.26 Additionally, studies by Orlic et al.27 have indicated that in mice with acute myocardial infarction, tissue regeneration occurs within 27 days of treatment with hematopoietic growth factors.

We investigated whether BMCs can be induced to adopt an RPE phenotype in vitro and the mechanism underlying this process. We then used C57BL/6 wild-type (wt) mice and C57BL/6 green fluorescence protein (GFP) chimeric mice in a model of NaIO3-induced RPE degeneration to test in vivo the hypothesis that circulating BMCs mobilized by G-CSF and FL would home to the site of RPE damage and express cell markers of RPE lineage. In the NaIO3 model, a direct correlation has been observed among decreased visual function, decreased electrophysiological function, and anatomic cell loss in the rodent RPE (and subsequently in the retina) after NaIO3 injection. Furthermore, the extent of the RPE damage is time and concentration dependent, as we recently demonstrated.28 Therefore, the selective, patchy loss of the RPE monolayer in the rodent after intravenous injection of NaIO3 provides a model for the study of RPE repopulation of bare areas of normal Bruch membrane.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Model of RPE Degeneration
Animals used in our study were maintained under standard conditions and treated according to the regulations in the ARVO Statement for the Use of Animal in Ophthalmic and Vision Research after approval by the University of Louisville Institutional Animal Care and Use Committee. Four- to 6-week-old male C57BL/6 wt mice weighing between 20 and 25 g were purchased from Harlan Sprague-Dawley Inc. (Indianapolis, IN). To easily identify the BMCs in the subretinal space, we also used GFP chimeric mice (kindly provided by Suzanne T. Ildstad, Institute for Cellular Therapeutics, University of Louisville), which show green fluorescence in cells of BM origin. Briefly, adult C57BL/6 recipient mice were lethally irradiated (950 cGy) and given whole BM transplantation from C57BL/6 mice transgenic for the chicken ß-actin promoter-GFP and cytomegalovirus enhancer. The BM was allowed to reconstitute for 2 months before the animals were used in the experiments. Average GFP chimerism of the BM established was 84.7% ± 4.7%.

For induction of RPE degeneration, mice were briefly restrained (TV-150; Braintree Scientific, Braintree, MA), and a single injection of sterile 1% solution of sodium iodate (NaIO3; Sigma, St. Louis, MO) in saline (0.9% NaCl) was administered into the tail vein of each mouse (50 mg/kg body weight). Saline-only injected animals served as controls.

RPE Preparation
For the preparation of mouse RPE cells, 10-day-old C57BL/6 wt mice were euthanatized, and the eyes were enucleated. Connective tissue was removed, and eyes were washed twice in PBS and incubated in 2% neutral protease (Dispase; Invitrogen, Carlsbad, CA) in Dulbecco modified Eagle medium (DMEM) at 37°C for 45 minutes. After removal of the anterior segment and the neurosensory retina, RPE cells were gently peeled mechanically with a rounded glass stick. The cells were then centrifuged at 1200 rpm for 5 minutes, resuspended, and triturated gently 10 times with DMEM plus 10% fetal bovine serum (FBS) using a glass Pasteur pipette. Cells were centrifuged again and resuspended with DMEM plus 10% FBS. The cells were then plated onto fibronectin-coated (10 µg/mL in PBS) culture dishes and cultured in DMEM, 10% FBS, and gentamicin. Cells were routinely stained for the RPE markers cytokeratin, RPE65, and microphthalmia transcription factor (MITF) so that their origins could be determined.

Isolation of GFP+ Sca-1+ BMCs
GFP+ whole BM was kindly provided by Suzanne T. Ildstad (Institute for Cellular Therapeutics). Briefly, BM was obtained from adult male GFP+ mice (C57BL/6-Tg [UBC-GFP]; Jackson Laboratory, Bar Harbor, ME). The femur and tibia were dissected and placed in RPMI 1640 culture medium. BM was obtained by slowly flushing medium inside the diaphyseal channel with a syringe through a 25-gauge needle. BM was then homogenized through a 20-gauge needle and filtrated with a nylon filter (70 µM; BD Biosciences, Bedford, MA). Cells were collected and Sca-1+ cells were isolated with the use of paramagnetic minibeads (Miltenyi Biotec, Auburn, CA) according to the manufacturer's protocol.

Coculture of GFP+ Sca-1+ BMCs with RPE
For direct coculture, RPE cells (35,000 cells/well) of passages 2 to 3 were plated on eight-well chamber slides coated with fibronectin (10 µg/mL in PBS) and cultured in DMEM, 10% FBS, and gentamicin for 4 hours to attach. Attached RPE cells were then treated with 100 µL mitomycin C solution (50 µg/mL; Sigma) for 20 minutes at 37°C to block proliferation. After the cells were washed twice with PBS, GFP+ Sca-1+ BMCs (35,000 cells/well) were added to the RPE cells in a ratio of 1:1. Cultures were maintained for 1, 2, and 3 weeks in Iscove modified Dulbecco medium (Invitrogen) supplemented with 20% horse serum, 10–6 M hydrocortisone, 10–4 M ß-mercaptoethanol, 2 mM L-glutamine, and 25 mM NaHCO3. Sca-1+ BMCs were also cocultured in the same ratio with a monolayer of mouse fibroblasts (3T3; CRL-1658; American Type Culture Collection, Manassas, VA). Additionally, cell survival was determined with a viability assay (Live/Dead; Molecular Probes, Eugene, OR).

For separated coculture, RPE cells (35,000 cells/well) of passages 2 to 3 were plated on fibronectin (10 µg/mL in PBS)-coated 24-well culture plates and treated with mitomycin C, as described. GFP+ Sca-1+ BMCs (17,500 cells/insert) were then added on membrane inserts with a pore size of 0.4 µm (Becton Dickinson, Franklin Lakes, NJ) placed in the well. This allowed the cells to share the culture medium but kept them physically separated by the membrane. Cell cultures were maintained as described.

BMC Mobilization
Mice were mobilized by daily subcutaneous injections of 10 µg FL from days 1 to 10 and of 7.5 µg G-CSF from days 1 to 6 (both generously provided by Amgen Inc., Thousand Oaks, CA). Mobilization started 4 hours after NaIO3 injection. Growth factors were diluted in saline before each injection to a total volume of 100 µL. Control animals were injected with saline only.

Fluorescence-Activated Cell Sorter
Aliquots of 100 µL PB were incubated with anti–Sca-1 phycoerythrin (PE), c-kit allophycocyanin (APC), CD8 FITC, Mac-1 FITC, B220 FITC, Gr-1 FITC, and T-cell receptor (TCR) FITC monoclonal antibodies (mAbs; BD PharMingen, San Diego, CA) for 30 minutes on ice. Cells were washed twice in FACS medium (PBS, 1% BSA, 0.1% NaN3). Red blood cells were lysed with ammonium chloride lysis buffer (0.83%) for 6 minutes at room temperature (RT). The remaining PB mononuclear cells (PBMNCs) were then washed twice and fixed in 1% paraformaldehyde (PFA), and flow cytometric analysis was performed (FACSCalibur; Becton Dickinson Biosciences, San Jose, CA). For enumeration of BMCs, cells negative for lineage markers (Lin–) and positive for Sca-1 (Ly-6A/E) were gated. Gated cells were then analyzed for their expression of c-kit (CD117). Lin–/Sca-1+/c-kit+ cells were defined as bone marrow–derived stem cells (BMSCs). Statistical analysis of flow data was performed (CellQuest software, version 3.0.1; Becton Dickinson). The percentage of BMCs of total PBMNCs was determined, and the absolute number per microliter of blood was calculated based on individual PBMNC counts.

Immunocytochemistry
In Vitro Experiments.
After 1 week, 2 weeks, and 3 weeks of coculture, cells were washed in PBS and fixed in 4% PFA at RT for 15 minutes. Subsequently, nonspecific staining was blocked with normal sheep serum (Serotec, Raleigh, NC) at RT for 1 hour. Cells were incubated with anti-mouse cytokeratin (1:100; Research Diagnostics, Flanders, NJ), anti-mouse RPE65 (1:200; Chemicon, Temecula, CA), anti-mouse MITF (1:75; Abcam, Cambridge, CA), anti-mouse glial fibrillary acidic protein (GFAP; 1:100; Sigma), or anti-mouse opsin (1:10,000; Sigma) primary antibodies at 4°C overnight. Control reagent (Universal Negative Control Reagent; Dako, Carpinteria, CA) for mouse antibodies was used for negative control. Cells were then washed three times for 5 minutes each in PBS and incubated with Cy3-conjugated sheep anti-mouse secondary antibody for 1 hour. Cells were washed three times in PBS for 5 minutes and mounted with 4',6'-diamino-2-phenylindole (DAPI) mounting medium (Vector, Burlingame, CA) to stain the cell nuclei. The number of cells was counted at 1000x magnification in four random visual fields with an area per field of 0.45 mm2, and the number of positive cells was used to calculate the percentage of positive cells. For coculture using membrane inserts, after fixation with 4% PFA, membranes with Sca-1+ cells on the top were carefully cut along the edges of the inserts and then stained for RPE markers as described.

In Vivo Experiments.
Mice were euthanatized at 2 and 4 weeks after NaIO3 injection, and eyes were enucleated and fixed in 4% PFA at 4°C overnight.

Whole eye flat mounts (FMs) were prepared by removing the anterior segment and the neurosensory retina and making four radial relaxing incisions in the remaining sclera-choroid-RPE complex. Nonspecific binding was blocked with 3% normal goat serum (Serotec) at RT for 60 minutes. The specimens were washed three times for 5 minutes in PBS and incubated overnight at 4°C with anti-mouse c-kit (1:100; R&D Systems, Minneapolis, MN), anti-mouse F4/80 (1:100; Serotec), or anti-mouse CD11b (1:100; Serotec) primary antibodies. After three 5-minute washes in PBS, specimens were incubated with Cy3-conjugated goat anti-rat secondary antibody at RT for 1 hour. The specimens were then washed three times for 5 minutes each in PBS and mounted with DAPI mounting medium.

For cross-sections, mouse eyes were embedded in paraffin, and consecutive sections (5 µm) were double-stained for GFP and RPE markers. Briefly, the specimens were deparaffinized and microwaved in citrate buffer (pH 6.0) at 350 W for 10 minutes. Sections were then blocked with 2% normal goat serum and 2% normal rabbit serum (both Serotec) at RT for 60 minutes. This was followed by incubation with chicken anti-mouse GFP (1:100) and mouse anti-mouse RPE65 (1:200; both Chemicon) or mouse anti-mouse MITF (1:75; Abcam, Cambridge, CA) primary antibodies at 4°C overnight. After three 5-minute washes in PBS, specimens were incubated with fluorescent dye (Alexa 647; Invitrogen, Carlsbad, CA)-conjugated goat anti-mouse and FITC-conjugated rabbit anti-chicken secondary antibodies at RT for 1 hour. The specimens were mounted with DAPI mounting medium after final washing steps in PBS.

Statistical Analysis
All experiments were independently performed three times, and five animals per group were used for the in vivo experiments. Data were presented as mean ± SD or SEM, depending on experimental design. Statistical significance was evaluated with an unpaired Student's t-test, and P < 0.05 was considered significant.


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
In Vitro Experiments
Isolated BMCs were placed in culture and evaluated for morphology and expression of RPE-specific markers. The morphology of GFP+ BMCs changed from round to flattened, polygonal cells after 7 days of coculture in direct contact with RPE cells (Fig. 1A) . Immunohistochemical staining showed that by day 14, BMCs started to express markers of RPE lineage—89% ± 6% of GFP+ cells expressed cytokeratin, 80% ± 11% expressed MITF, and 81% ± 13% expressed RPE65. Figure 1B depicts a representative example of positive RPE65 staining. By day 21, 93% ± 8% of cells expressed RPE65 and 88% ± 8% expressed MITF. These effects were abolished when the BMCs were separated by a semipermeable membrane and were not in direct cell–cell contact with the RPE. The astrocyte marker GFAP (Fig. 2A) and the photoreceptor marker opsin (Fig. 2B) were not expressed by BMCs in coculture with adult RPE. Furthermore, BMCs cocultured with fibroblasts showed no morphologic changes and were negative for the studied RPE markers (Fig. 2C ; MITF). RPE and BMC cells without coculture stained positively for the respective lineage markers (RPE65, Sca-1) during the investigated culture period. Survival rates for BMCs, RPE cells, and fibroblasts after 3 weeks in coculture were 94.7%, 95.1%, and 93.1%, respectively. Thus, the proportion of each cell type over time was stable.


Figure 1
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  		FIGURE 1. Morphology and phenotype of in vitro cocultured BMCs. (A) Comparison of the morphology of GFP+ BMCs cocultured with RPE cells (left) or fibroblasts (right; green). Original magnification, 630x. In contrast to fibroblast coculture, BMCs changed from round to flattened, polygonal cells after 7 days. (B) Coculture of RPE and GFP+ BMCs showed double-positive staining for the RPE-specific marker RPE65 (upper left; Cy2-red) and GFP (upper right; Cy2-green), indicating RPE-specific differentiation at 2 weeks (lower right; merged fluorescence, orange). All cell nuclei were stained with DAPI (lower left; blue) and the same individual cells in each panel are marked with arrows (BMCs) or arrowheads (host RPE). The prominent nuclear staining in the GFP and Cy3 images represent contamination from oversaturation of the DAPI signal.

 

Figure 2
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  		FIGURE 2. Coculture of GFP+ BMCs (green) and RPE shows no positive staining for (A) glia-specific GFAP and (B) photoreceptor-specific opsin, indicating cell type-specific differentiation at 2 weeks. Immunocytochemical staining also showed that GFP+ BMCs (green) cocultured in direct contact with mouse fibroblasts were negative for the RPE marker RPE65 (C). All cell nuclei were stained with DAPI (blue). Our positive controls for GFAP and opsin consisted of immunohistologic sections of neural retina.

 
In Vivo Experiments
Mobilization.
We evaluated whether RPE damage by NaIO3 induced the mobilization of BMCs into the peripheral circulation. Mice were injected intravenously with NaIO3 or saline; this was followed 3 and 5 days later by flow cytometry examination of the PB for the presence of Lin– Sca-1+ c-kit+ BMSCs. Circulating Lin– Sca-1+ c-kit+ cells were not detected in significantly higher numbers in NaIO3 injected mice than in saline controls (Fig. 3A) , indicating that NaIO3 alone did not increase the mobilization of BMCs into the PB.


Figure 3
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  		FIGURE 3. Influence of NaIO3 on BM mobilization into the PB. Values are given as mean ± SEM of three independent experiments. (A) Number of Lin– Sca-1+ c-kit+ BMSCs in peripheral blood in NaIO3 or saline-injected (control) mice on days 3 and 5. Representative FACS blots of day 5 are shown below. (B) Number of Lin Sca-1+ c-kit+ BMSCs in PB on day 11 in NaIO3-treated mice after injection with saline, G-CSF, FL, or a combination. Representative FACS blots of control and combination groups are shown below. The increase of BMCs in PB after mobilization was significant (*P < 0.01).

 
We assessed the potential of growth factor administration to mobilize BMCs in our NaIO3 model by determining the absolute numbers of BMSCs in the PB after treatment with FL alone, G-CSF alone, or a combination of FL and G-CSF. As shown in Figure 3B , animals treated with G-CSF and FL alone showed a significant 4-fold and 48-fold increase of BMCs (P < 0.01), respectively. Combined administration of G-CSF and FL showed a synergistic effect, with a 77-fold increase of BMCs on day 11 after mobilization (P < 0.01).

Homing.
To determine whether mobilized circulating BMCs had the ability to home to the damaged tissue within the subretinal space, C57BL/6 wt mice were studied after mobilization with G-CSF and FL. Fourteen days after NaIO3 injection, c-kit+ cells were observed in the subretinal space in C57BL/6 wt mice (Fig. 4A) but not in control mice, in which no positive staining was detected. However, the number of BMCs in the subretinal space was small (1.2 ± 0.2 cells/microscopic visual field).


Figure 4
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  		FIGURE 4. Immunocytochemical staining of flat mounts of RPE and choroid 2 weeks after NaIO3 treatment and BMC mobilization; c-kit+ cells (Cy3-red) were observed in wt (A) and chimeric (B) mice. Merged green (GFP) and red (Cy3) fluorescence stained yellow to orange, depending on the intensity of green fluorescence. F4/80+ (Cy3-red) macrophages (C) and CD11b+ (Cy3-red) leukocytes (D) were also found in the subretinal spaces of chimeras.

 
We then investigated the ability of BMCs to home to the subretinal space of GFP chimeric mice after mobilization with G-CSF and FL. GFP+ BMCs of different sizes were visible within the subretinal space in control and NaIO3-treated mice 14 days after injection. GFP+ cells were most abundant within and adjacent to the areas of RPE damage. With the use of immunocytochemistry, we investigated the colocalization of GFP with markers for BMCs (c-kit), macrophages (F4/80), and leukocytes (CD11b) and found that 21% ± 8% expressed c-kit compared with 9% ± 4% in controls (P < 0.05; Fig. 4B ). F4/80 (Fig. 4C) and CD11b (Fig. 4D) cells were also observed, indicating that BM-derived macrophages and leukocytes had also been recruited to the subretinal space of the chimeric mice. Most of the cells positively stained for RPE or leukocyte/macrophage makers were found in the subretinal space in areas devoid of RPE. In the wild-type (i.e., non-NaIO3–treated) controls, the cells were randomly distributed over the flat mounts.

RPE Lineage Markers.
Furthermore, we double stained GFP+ BMCs with RPE-specific markers RPE65 and MITF at 1 and 2 months after NaIO3 injection to determine whether these cells in the subretinal space were induced to express RPE cell markers. RPE65 (not shown) and MITF (Fig. 5) were both expressed by GFP+ cells. The cells formed a monolayer in focal areas of RPE damage in the subretinal space. These results indicated that BMCs had acquired phenotypic markers of RPE cells within this milieu. Occasionally, the vertical sections also showed GFP+ cells in the choroid, but these cells did not express RPE cell markers.


Figure 5
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  		FIGURE 5. Immunocytochemical staining of vertical sections of a GFP chimeric mouse eye 4 weeks after NaIO3 treatment and BMC mobilization. GFP+ BMCs were stained with anti-GFP (Cy2-green) and anti-MITF (Cy3-red). The merged Nomarski image shows a monolayer of MITF+ GFP+ BMCs (yellow) on Bruch membrane. The monolayer of BMCs can be seen adjacent to pigmented host RPE cells (arrows) outlined with white dashes.

 

   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
We have demonstrated that BMCs can be induced to assume an RPE morphology in vitro and to express cell markers of RPE lineage in vitro and in vivo. Most BMCs adopted a flat, epithelial cell-like morphology in culture and stained for the RPE-specific markers cytokeratin, RPE65, and MITF. Coculture with separation of the BMCs from the RPE by a semipermeable membrane and coculture on a layer of fibroblasts did not result in a change in either morphology or lineage marker expression, indicating that direct cell–cell contact between these cells is required. On the other hand, our in vivo results suggested the same requirement because GFP+ BMCs in the choroid of NaIO3-injected mice did not express RPE-specific markers. However, some of the RPE markers (protein, mRNA) might have been transferred to the BMCs by cell-derived microparticles because mitomycin C induces vesiculation of cells in culture.29

We also found that mobilization of BMCs into the peripheral circulation, combined with RPE damage, led to homing of BMCs to the subretinal space and the expression of RPE lineage markers by these cells. The ability of exogenous BMCs to home to damaged tissue and to differentiate into cells of the host organ has been shown.8 14 15 Numerous circulating stem cells and organ damage are two determinants identified as critical for homing and differentiation of BMCs in a variety of tissues.14 27 30 31 32 In the present study, we could maximize the number of BMCs in the peripheral circulation by using endogenous stem cell mobilization to increase the probability of their homing to the site of RPE damage without invasive procedures. The use of endogenous BMC clearly avoids the risks associated with the transplantation procedure.

In our experiments, we used C57BL/6 wt mice and C57BL/6 GFP chimeric mice to study the homing of stem cells to the subretinal space. Chimeric mice must undergo irradiation to permit acceptance of the transplanted BM and create the chimera. We observed more c-kit+ cells in the subretinal space in chimeric mice than in wt mice. Control C57BL/6 wt mice did not have c-kit+ cells in the subretinal space, whereas these cells were observed in control GFP chimeric mice, though the number was significantly less than in NaIO3 chimeric mice. These results suggested that the process of creating chimerism might have enhanced mobilization of BMCs from the BM so that these cells were in the subretinal spaces of chimeric mice even though there was no retinal damage or the radiation used to condition chimeric mice for BM transplantation resulted in mild damage to the retina. We occasionally found mild cataract in the chimeric mice, consistent with reports that the radiation damage to the eye is most often localized to the lens.33 34 NaIO3 does cause primary injury to the RPE but does not interfere with the BM mobilization induced by G-CSF and FL in wt mice as comparison with data by Neipp et al. shows.35 Therefore, the combination of whole body irradiation, growth factor mobilization, and NaIO3 toxicity might have optimized the conditions for BMCs to home to the eye.

Furthermore, we found that 21% of GFP+ BM-derived cells in the damaged subretinal space were immunoreactive for c-kit. After homing to the eye, they started to express markers of RPE lineage and formed a monolayer in focal areas of RPE damage in the subretinal space. We observed that BM-derived leukocytes and macrophages were also recruited to the subretinal space in chimeric mice that received G-CSF and FL. This was documented in control and NaIO3-treated animals, suggesting that the recruitment of these cells to the eye was driven by the irradiation used to condition the chimeras.

Stromal cell-derived factor (SDF)-1, a critical chemokine in the recruitment of stem cells to an area of injury, is increased in animal models of liver, limb, and heart damage.36 37 38 39 Recent work in our laboratory showed that cytokines important for stem cell homing—SDF-1, C3, and HGF—were up-regulated in the subretinal space of mice after NaIO3 treatment.40 The increased local expression of these cytokines may be important in the homing of BMCs to focal areas of RPE damage. However, we did not investigate this possibility in the present study.

The bone marrow harbors several different types of stem cells, including HSCs (Lin–, Sca-1+, c-kit+, CD45+), mesenchymal stem cells (adherent, CD105+), and tissue-committed stem cells (TCSCs; small, highly mobile, Lin–, Sca-1+, CD45–). These cells have different phenotypes and different capacities to differentiate into other cell types. We did not explore the possible mechanisms underlying differentiation within the milieu of the damaged RPE. However, stem cell plasticity, which involves mobilization and homing of CD45+ HSCs from the BM to the site of injury and transdifferentiation into cell lineages of the host organ, is one possibility.41 Spontaneous fusion of the migrated BMCs with host cells also must be considered,42 but the existence of a monolayer of GFP+ cells argues against this explanation. Alternatively, predifferentiated CD45– TCSCs that can be activated by a specific tissue injury and can home to the damaged site for tissue repair may exist.43 44 Recent work in our laboratory indicated that ocular-committed stem cells, characterized by the expression of early ocular markers, may preexist in the BM. These TCSCs were mobilized into the peripheral circulation by G-CSF and FL and migrated, in vitro, along gradients of cytokines secreted by damaged RPE.40 Our hypothesis is that these are the cells that migrated to the subretinal space in our model, that is, the Lin–, Sca-1+, CD45– BMCs.

In conclusion, our findings suggest that BMCs, once mobilized, have the ability to respond to signals from damaged RPE, migrate to the altered subretinal space, and form a monolayer of cells that express markers of RPE lineage. This suggests the potential use of BMCs as therapy for AMD and other hereditary eye diseases. Our study, however, did not show that these cells have the anatomic or functional characteristics of RPE cells. Strategies to optimize the number of BMCs within the subretinal space and the demonstration that these cells are anatomically and functionally differentiated RPE cells is necessary before such an approach should be considered for the treatment of AMD.


   Acknowledgements
 
The authors thank George B. Harding for his assistance with scanning laser microscopy and Franziska Flückiger for her technical assistance with cell culture experiments.


   Footnotes
 
Supported by the National Institutes of Health (Grants 1 R41 EY015336–01A2, R24 EY015636), the Scientific and Technological Research Council of Turkey (TUBITAK), the Research Challenge Trust Fund, and Research to Prevent Blindness.

Submitted for publication August 28, 2006; revised February 1 and May 1, 2007; accepted July 16, 2007.

Disclosure: Y. Li, None; P. Atmaca-Sonmez, None; C.L. Schanie, None; S.T. Ildstad, None; H.J. Kaplan, None; V. Enzmann, None

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Corresponding author: Volker Enzmann, Department of Ophthalmology, Inselspital, University of Bern, Freiburgstrasse 14, 3010 Bern, Switzerland; volker.enzmann{at}insel.ch.


   References
Top
 Abstract
 Materials and Methods
 Results
 Discussion
 References
 

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V. Enzmann, E. Yolcu, H. J. Kaplan, and S. T. Ildstad
Stem Cells as Tools in Regenerative Therapy for Retinal Degeneration
Arch Ophthalmol, April 1, 2009; 127(4): 563 - 571.
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