Cell Homogeneity Indispensable for Regenerative Medicine by Cultured Human Corneal Endothelial Cells

Abstract

Purpose: To identify the subpopulation (SP) among heterogeneous cultured human corneal endothelial cells (cHCECs) devoid of cell-state transition applicable for cell-based therapy.

Methods: Subpopulation presence in cHCECs was confirmed via surface CD-marker expression level by flow cytometry. CD markers effective for distinguishing distinct SPs were selected by analyzing those on established cHCECs with a small cell area and high cell density. Contrasting features among three typical cHCEC SPs was confirmed by PCR array for extracellular matrix (ECM). Combined analysis of CD markers was performed to identify the SP (effector cells) applicable for therapy. ZO-1 and Na⁺/K⁺ ATPase, CD200, and HLA expression were compared among heterogeneous SPs.

Results: Flow cytometry analysis identified the effector cell expressing CD166⁺CD105⁻CD44⁻∼^+/−CD26⁻CD24⁻, but CD200⁻, and the presence of other SPs with CD166⁺ CD105⁻CD44⁺⁺⁺ (CD26 and CD24, either + or −) was confirmed. PCR array revealed three distinct ECM expression profiles. Some SPs expressed ZO-1 and Na⁺/K⁺ ATPase at comparable levels with effector cells, while only one SP expressed CD200, but not on effector cells. Human leukocyte antigen expression was most reduced in the effector SP. The proportion of effector cells (E-ratio) inversely paralleled donor age and decreased during prolonged culture passages. The presence of Rho-associated protein kinase (ROCK) inhibitor increased the E-ratio in cHCECs. The average area of effector cells was approximately 200∼220 μm², and the density of cHCECs exceeded 2500 cells/mm².

Conclusions: A specified cultured effector cell population sharing the surface phenotypes with mature HCECs in corneal tissues may serve as an alternative to donor corneas for the treatment of corneal endothelial dysfunction.

Although human corneal endothelial cells (HCECs) are mitotically inactive and are arrested at the G1 phase of the cell cycle in vivo,¹ they retain the capacity to proliferate in vitro.² However, a recent study³ has shown that culturing HCECs for a long period of time is extremely difficult. 

Since the proliferative potential of HCECs is limited,^1,4,5 severe damage to the corneal endothelium due to pathologic conditions leads to corneal endothelial dysfunction and the loss of corneal transparency.^6–8 At present, corneal transplantation is the only available treatment for such patients. Thus, new technology such as tissue engineering is being explored in regard to corneal endothelial transplantation.⁹ 

To date, most researchers conceptualize cultured HCECs (cHCECs) only from the aspect that they are derived from corneal endothelium tissue, and with little attention to details pertaining to the refinement of the biochemical features. In fact, cHCECs are heterogeneous from culture to culture in their morphology and in their surface markers, such as cluster of differentiation (CD) antigens.^10–12 Studies,^13–20 either directly or indirectly, and including those from Joyce et al., have shown that heterogeneity is present in HCEC cultures. Of particular and striking interest is the finding by Miyai et al.¹⁰ of the presence of frequent chromosomal aneuploidy in cHCECs, thus indicating the presence of heterogeneous subpopulations (SPs) with or without aneuploidy. 

Trials pertaining to the in vitro expansion of cHCECs without cell-state transition (CST), such as epithelial-mesenchymal transition (EMT), and without karyotype aneuploidy have been hampered by the limited insights to date on the cellular features of cHCECs due to the limited availability of reproducible cultures, as well as the distinction between cHCEC populations obtained from either young donors or elderly donors. Cultured HCECs have an inclination toward CST into a senescence phenotype, EMT, and fibroblastic cell morphology. Recently, our group has been attempting to identify, via the use of defined cell surface markers, the HCEC SPs that are clinically applicable for the reconstitution of impaired corneal endothelium. However, the analysis of cHCECs, without using immortalized HCECs, has proved frustrating, as the plasticity of the metabolic profiles of cHCECs (Hamuro et al., manuscript under revision) possibly interferes with the correct interpretation of heterogeneous composites in cHCECs. 

In consideration of the reported karyotype aneuploidy observed in cHCECs, as well as the plasticity of their metabolic profiles, we selected several CD markers to define HCEC SPs, namely, CD166, CD133, CD105, CD90, CD73, CD49e, CD44, CD26, and CD24, all of which have a linkage to mesenchymal stem cells (MSCs), cancer stem cells (CSCs), or to phenotypic conversion during CST.^21–25 The selection was auxiliary, yet based on the reasoning that normal stem cells are the longest lived cells in tissues and are more likely to accumulate mutations over time, and that CSCs may arise from transit-amplifying cells (i.e., one-step differentiated cells form stem cells and have the property of proliferating rapidly, albeit transiently, with a limited life span).^26,27 

To date, no HCEC-specific cell surface markers have been described. Glypican-4 and CD200 have been proposed as HCEC markers to distinguish HCECs from corneal stromal fibroblasts.¹² However, the practical problem associated with HCEC culture is the potential contamination with vulnerable transformed cHCECs.¹¹ 

Flow cytometry analysis demonstrated the presence of several cHCEC SPs and showed that one specific cHCEC SP, with surface expression of CD166⁺, CD105⁻, CD44⁻, CD26⁻, and CD24⁻, is the SP without CST. This is the first relevant finding making it possible to adapt the SP in the clinical setting; moreover, the combination of CD markers defined in this study was found to be the most appropriate for quality control to ensure the functional characteristics of cHCECs for clinical application. 

Materials and Methods

Human Corneal Endothelial Cell Donors

The human tissue used in this study was handled in accordance with the tenets set forth in the Declaration of Helsinki. Human corneal endothelial cells were obtained from human donor corneas obtained from SightLife (Seattle, WA, USA) eye bank and were cultured before performing karyotyping analysis. Informed written consent for eye donation for research was obtained from the next of kin of all deceased donors. All tissues were recovered under the tenets of the Uniform Anatomical Gift Act of the particular state in which the donor consent was obtained, and the tissue was recovered. 

All donor corneas were preserved in Optisol GS (Chiron Vision, Inc., Irvine, CA, USA) and imported via international air transport for research purposes. Donor information accompanying the donor corneas showed that they were all considered healthy and absent of any corneal disease, and that all donors had no past history of chromosomal abnormality. 

Cell Cultures of HCECs

Unless otherwise stated, the HCECs were cultured according to published protocols, with some modifications.¹¹ Briefly, the Descemet membranes with the CECs were stripped from donor corneas and digested at 37°C with 1 mg/mL collagenase A (Roche Applied Science, Penzberg, Germany) for 2 hours. The HCECs obtained from a single donor cornea were seeded in one well of a Type-I collagen–coated six-well plate (Corning, Inc., Corning, NY, USA). The culture medium was prepared according to published protocols. Briefly, basal medium was prepared with OptiMEM I (Life Technologies Corporation, Carlsbad, CA, USA), 8% fetal bovine serum, 5 ng/mL epidermal growth factor (Life Technologies), 20 μg/mL ascorbic acid (Sigma-Aldrich Corporation, St. Louis, MO, USA), 200 mg/L calcium chloride (Sigma-Aldrich), 0.08% chondroitin sulfate (Wako Pure Chemical Industries, Ltd., Osaka, Japan), and 50 μg/mL gentamicin. Mesenchymal stem cell–conditioned medium was prepared as previously described.²⁸ The HCECs were cultured by using MSC-conditioned medium at 37°C in a humidified atmosphere containing 5% CO₂, and the culture medium was changed twice per week. The HCECs were passaged at ratios of 1:3 by using 10x TrypLE Select (Life Technologies) at 37°C for 12 minutes when they reached confluence. The HCECs at passages 2 through 5 were used for all experiments. 

Phase Contrast Microscopy

Phase contrast images were obtained by use of an inverted microscope system (CKX41; Olympus Corporation, Tokyo, Japan). For the area distribution analysis, the cHCECs were washed three times with phosphate-buffered saline (PBS)(-) (Ca⁺⁺ and Mg⁺⁺), and phase contrast images were then acquired by use of a BZ X-700 Microscope System (Keyence Corporation, Osaka, Japan). The area distributions were quantified by BZ-H3C Hybrid Cell Count Software (Keyence). 

Flow Cytometry Analysis of cHCECs

Human corneal endothelial cells were collected from the culture dish by TrypLE Select treatment as described above and suspended at a concentration of 4 × 10⁶ cells/mL in FACS buffer (PBS containing 1% bovine serum albumin [BSA] and 0.05% NaN₃). Next, an equal volume of antibody solution was added and incubated at 4°C for 2 hours. The antibody solutions were prepared by appropriately combining the following antibodies: FITC-conjugated anti-human CD26 mAb, PE-conjugated anti-human CD166 mAb, PerCP-Cy 5.5–conjugated anti-human CD24 mAb, PE-Cy 7–conjugated anti-human CD44 (all from BD Biosciences, San Jose, CA, USA), APC-conjugated anti-human CD105 (eBioscience, Inc., San Diego, CA, USA), DyLight 488–conjugated anti human LGR5 antibody (LifeSpan Biosciences, Inc., Seattle, WA, USA), and Alexa Fluor 647–conjugated anti-HLA class I mAb (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA). After washing with FACS buffer, the HCECs were analyzed by use of a BD FACSCanto II Flow Cytometry System (BD Biosciences). 

Immunocytochemical Staining

For immunocytochemical staining, cHCECs were fixed with ice-cold methanol for 10 minutes, and then permeabilized with PBS(-) containing 0.1 % Triton X-100 at room temperature (RT) for 15 minutes. After blocking of nonspecific reactivity with 1% BSA in PBS(-) at RT for 1 hour, the samples were incubated at 4°C overnight with antibodies against Na⁺/K⁺-ATPase (EMD Millipore Corporation, Temecula, CA, USA), ZO-1 (Life Technologies), followed by N-Histofine MAX-PO (MULTI) (Nichirei Biosciences, Inc., Tokyo, Japan) detection reagent. After washing with PBS(-) containing 0.1 % Triton X-100, cells were developed with N-Histofine Simple Stain DAB Solution (Nichirei Biosciences) and counterstained with hematoxylin (Merck KGaA, Darmstadt, Germany). Finally, cells were mounted with N-Histofine Aqueous Mounting Medium (Nichirei Biosciences) and observed under a bright-field microscope. 

Immunohistochemical Staining

Descemet membranes with the CECs were stripped from donor corneas and placed on silanized slides (DAKO Denmark A/S, Glostrup, Denmark), air dried, and refixed in 4% paraformaldehyde for 10 minutes. After washing in PBS(-) containing 0.1% Triton X-100 at RT for 15 minutes, sections were incubated with 1% BSA (Nacalai Tesque, Inc., Kyoto, Japan) at RT for 1 hour to block nonspecific binding. The sections were then incubated with primary antibody at 4°C overnight. The primary antibodies used were as follows: anti-LGR5 antibody (GenTex, Inc., Irvine, CA, USA), anti-CD24 antibody, anti-CD26 antibody, anti-CD166 antibody (all from BD Biosciences), and anti-CD44 antibody (Trans Genic, Inc., Kobe, Japan) and Na⁺/K⁺-ATPase (EMD Millipore Corporation). After washing with PBS(-), the sections were then incubated with the secondary antibodies Alexa Fluor 594–conjugated anti-mouse IgG Ab and Alexa Fluor 488–conjugated anti-rabbit IgG Ab (Life Technologies) at RT for 1 hour. After washing with PBS(-), the sections were then incubated with the secondary antibodies Alexa Fluor 594–conjugated anti-mouse IgG Ab and Alexa Fluor 488–conjugated anti-rabbit IgG Ab (Life Technologies) at RT for 1 hour. After washing with PBS(-), the sections were then coverslipped by using VECTASHIELD Antifade Mounting Medium with DAPI (Vector Laboratories, Inc., Burlingame, CA, USA) and examined under a fluorescence microscope (BZ-9000; Keyence). 

Preparation of Noncultured HCECs

For the preparation of noncultured HCECs, Descemet membranes with the CECs were stripped from donor corneas and soaked at 4°C overnight in 1000 PU/mL dispase (Dispase I; Sanko Junyaku, Co., Ltd., Tokyo, Japan). Next, these were resuspended in 1x TrypLE Select and incubated at 37°C for 5 minutes. The dissociated HCECs were then washed with FACS wash buffer and subjected to FACS analysis. 

BD Lyoplate Screening

Screening of cell surface markers was performed by assessing the expression of those markers through a BD Lyoplate Human Cell Surface Marker Screening Panel (BD Biosciences) according to the manufacturer's protocol. Briefly, cultured HCECs were incubated with primary 242 antibodies and isotype IgGs (BD Biosciences) at the dilution indicated by the manufacturer's protocol at 4°C for 30 minutes. The cells were washed with PBS containing 1% BSA and 5 mM EDTA, and then incubated with Alexa Fluor 647–conjugated secondary antibodies (1:200; BD Biosciences) at 4°C for 30 minutes. The cells were washed again with PBS containing 1% BSA and 5 mM EDTA, and then analyzed by use of the FACSCanto II Flow Cytometry System and CellQuest Pro software (BD Biosciences). 

Cell Sorting

For cell sorting experiments, HCECs were collected and stained with FITC-conjugated anti-human CD24 mAb and PE-Cy 7–conjugated anti-human CD44 mAb (BD Biosciences) as described above. After washing with FACS buffer, the cells were resuspended in FACS buffer. The CD24⁻/CD44⁺ and CD24⁻/CD44⁻ cells were sorted by use of a BD FACSJazz cell sorter (BD Biosciences) and seeded at a density of 4.2 × 10⁴ cells on a 24-well cell culture plate for subsequent analysis. 

Polymerase Chain Reaction (PCR) Array

Total RNA was extracted from the cHCECs by use of the miRNeasy Mini kit (Qiagen, Hilden, Germany). Complementary DNA synthesis was performed with 100 ng total RNA for a 96-well plate format by use of an RT² First Strand kit (Qiagen). Expression of endothelial mRNAs was investigated by use of an RT² Profiler PCR Array Human Extracellular Matrix and Adhesion Molecules (Qiagen), according to the manufacturer's recommendations, and then analyzed by use of an RT² Profiler PCR Array Data Analysis Tool version 3.5 (Qiagen). 

Results

Phenotypic Variations Among Cultures

Human corneal endothelial cells obtained from the corneas of five aged donors, one young donor (the age range of the aged and young donors was above 50 years and between 7 and 29 years, respectively), and two newborn donors were cultured according to the method described by Okumura et al.,¹¹ and the surface expression of CD166, CD105, CD44, and CD24 was then characterized (Table 1). The other set of data for cHCECs from other donor corneas for CD166, CD105, CD44, and LGR5, instead of CD24, is summarized in Table 1. Depending on the different donors, a great distinction was found in the proportions of SPs defined with the surface markers. The highest proportion resided in the SP with CD166⁺CD105⁺CD44⁺ CD24⁻, whereas the highest SP in cHCECs from elderly donors No. 1 and No. 2 was the SP with CD166⁺CD105⁺CD44⁺CD24⁻. The highest SP with CD166⁺CD105⁺CD44⁺LGR5⁻ is shown in Table 1. Interestingly, even primary cHCECs exhibited various phenotypes during cultures under the same culture protocol. 

Table 1

 

CD166, CD105, CD44, CD24, and LGR5 Expression of the Cultured HCECs

 

To confirm the heterogeneity depending on the passages, many cultures were monitored by flow cytometry for the changes of SP composition. The representative FACS analysis is summarized in Figure 1. Some of the cHCECs are shown in Figure 2, together with phase contrast microscopy; Figure 2A corresponds to Figure 1A, and Figure 2B corresponds to Figure 1B. In summary, both the number of culture passages and the difference of donors exhibited great variation in the proportion of SPs. This finding immediately pointed to the necessity of finding the appropriate method of defining cHCECs in biochemical terms, instead of using the well-established term “cultured corneal endothelial cells,” in order to avoid the discourse in this field becoming a “catch-22.” 

Figure 1

 

Varied expression of CD166, CD24, CD44, and CD105 in cHCECs over passages. The expression of CD166, CD24, CD44, and CD105 was analyzed by use of a FACSCanto II Flow Cytometry System. After gating for CD166⁺CD24⁻ (R1) or CD166⁺CD24⁺ (R2), the following five SPs were defined: CD166⁺CD24⁻CD44^−/+CD105^+/− SP (gate [G] 1), CD166⁺CD24⁻CD44⁺⁺CD105⁺ SP (G2), CD166⁺CD24⁻CD44⁺⁺⁺CD105⁺ SP (G3), CD166⁺CD24⁺CD44⁺CD105⁺ (G4), and CD166⁺CD24⁺CD44⁺⁺CD105⁺ SP (G5). (A) Human corneal endothelial cells from a 53-year-old donor. (B) Human corneal endothelial cells from a 71-year-old donor.

Figure 2

 

Morphologic and SP composite variances in cHCECs over culture passages. Phase contrast images were taken and the changes in the ratios of SPs defined in Figure 1 were analyzed (•: G1, ▪: G2, ▴: G3, □: G4, ▵: G5). (A) Cultured HCECs from a 53-year-old donor (corresponding to Fig. 1A). (B) Cultured HCECs from a 10-year-old donor. (C) Cultured HCECs from a 71-year-old donor (corresponding to Fig. 1B). (D) Cultured HCECs from a 75-year-old donor.

Homogeneity Not Sufficed by ZO-1 and Na⁺/K⁺ ATPase Expression

Both ZO-1 and Na⁺/K⁺ ATPase, well known as the markers of HCECs, were stained for CD44^+/− SPs, but also for CD44⁺⁺CD24⁻ and CD44⁺⁺CD24⁺ SPs with stratified and fibroblastic morphology (Fig. 3). One CD44⁺⁺⁺CD24⁺ SP with typical stratified and fibroblastic morphology was devoid of these expressions (Fig. 3). 

Figure 3

 

The expression of Na⁺/K⁺ ATPase and ZO-1 in HCEC SPs. The HCECs from four different donors were cultured and the expression of Na⁺/K⁺ ATPase and ZO-1 was detected. The ratios of SPs in each group of cHCECs were also determined as in Figure 1.

CD200 and HLA Class I Expression Among SPs

The expression of CD200, previously reported as a marker of HCECs, was surprisingly restricted to only one CD44⁺⁺ SP with vulnerable CST⁺ cHCECs (Fig. 4C). Importantly, CD44⁻ cells did not show any sign of CD200 expression (Fig. 4A), and even more interestingly, some SPs with high CD44 positivity did not express CD200. Regarding the immunogenicity of cHCECs that can be transplanted into allogeneic hosts, our findings showed that SPs differed in the expression of surface HLA class I antigen, thus illustrating a parallel decrease with the decrease of CD44 and CD26 expression (Fig. 5). 

Figure 4

 

The expression of CD200 on cHCECs. The cHCECs were analyzed for the expression of CD44 and CD200 by use of the FACSCanto II Flow Cytometry System. (A) Human corneal endothelial cells from a 4-year-old donor (passage 3). (B) Human corneal endothelial cells from an 8-year-old donor (third passage). (C) Human corneal endothelial cells from a 3-month-old donor (passage 3). (D) Human corneal endothelial cells from a 59-year old donor (passage 3).

Figure 5

 

Different expressions of HLA class I on the cHCEC SPs. The cHCECs from four different donors were analyzed for the expression of CD26, CD44, and HLA class I by use of a FACSCanto II Flow Cytometry System. After gating for CD26⁻CD44^−/+ (blue), CD26⁻CD44⁺⁺ (green), CD26⁻CD44⁺⁺⁺ (orange), or CD26⁺CD44⁺⁺⁺ (purple), HLA class I expression of each gate was plotted in a histogram.

Flow cytometry analysis was further extensively performed to define the SP adaptable for cell injection into the anterior chamber. The phenotyping of cHCECs was routinely performed with CD166, CD105, CD44, CD26, CD24, HLA-I, HLA-DR/DP/DQ, and PD-L1. Simultaneously, we tested the expression of CD markers in freshly excised corneal tissues and found that they were negative for CD105, CD44, CD26, and CD24 (Fig. 6A). Based on this immunohistochemical study, we defined SPs with CD105, CD44, CD26, and CD24 negativity and CD166 positivity as effector cells that ensure application to cell injection therapy. The representative immunohistochemical staining is shown in Figure 6B. In our preliminary yet extensive experiments, the fresh human corneal endothelium showed no sign of the presence of CD24, CD26, and CD44, yet did manifest a strong expression for CD166. This indicates that the phenotypes of the cHCEC SP specified here, with complete absence of aneuploidy, are very consistent with those of HCECs in fresh corneal tissue. 

Figure 6

 

Expression of CD24, CD26, CD44, CD105, CD166, and LGR5 on the fresh corneal endothelial cells. (A) Corneal endothelial cells were collected from a 71-year-old donor cornea (female, endothelial cell density [ECD] = 3008 cells/mm²) and analyzed for the expression of CD166, CD24, CD44, and CD105. (B) Immunohistochemical staining of the cornea from a 65-year-old donor (female, ECD = 3311/3476 cells/mm²) was performed before cell dissociation (please refer to Materials and Methods in the main text).

The Sorted CD44⁻SP Elicited the Phenotype Without CST

CD44 is known to actively contribute to the maintenance of stem-cell features, and the functional contribution of CD44 relies on its particular ability to communicate with neighboring molecules, adjacent cells, and the surrounding matrix.²⁹ Accordingly, CD44⁻ (gate I in Fig. 7) and CD44⁺ (gate II in Fig. 7) SPs were sorted by use of the BD FACSJazz cell sorter, and purified SPs were cultured in 24-well plates followed by 17-days further culture. Subpopulation isolated as CD44⁺ SP proliferated rapidly with spindle-like morphology and it showed weak irregular staining of Na⁺/K⁺ ATPase, whereas the CD44⁻ SP showed relatively slow growth and evident expression of Na⁺/K⁺ ATPase (Fig. 7). These results further support the newly introduced concept of effector cells in SPs suitable for cell-based therapy. 

Figure 7

 

The expression of Na⁺/K⁺ ATPase on CD44⁺ and CD44⁻ HCECs. Cultured HCECs from a 73-year-old female donor (ECD = 3543/3406 cells/mm²) (A) were sorted by the expression of CD24 and CD44 by use of a FACSJazz cell sorter (B). The sorted cells were plated at the density of 220 cells/mm², and phase contrast images were then captured at 3, 10, and 17 days of cultivation (C). After 17-days cultivation, the cells were stained with anti–Na⁺/K⁺ ATPase antibodies.

Discrimination of SPs by PCR Array for Extracellular Matrix and Adhesion (EMA)

Investigation of the expression of endothelial mRNAs from effector cells and the other two SPs with CST by use of the RT² Profiler PCR Array Human Extracellular Matrix and Adhesion Molecules confirmed contrasting features among three typical cHCEC SPs. As shown in Figure 8, cluster (heat map) analysis demonstrated a clear distinction among those three SPs, therefore indicating, once again, the presence of an effector SP distinct in its EMA gene signatures. 

Figure 8

 

Functional discrimination of SPs by PCR array for ECM. The gene signatures were compared by PCR arrays between CD44⁻ SPs and SPs with CST by using the RT² Profiler PCR Array Human Extracellular Matrix. Messenger RNA was extracted from cHCECs (No. 66 [P5, donor age: 23 years, ECD = 3504 cells/mm²], No. 67 [P5, donor age: 67 years, ECD = 2717 cells/mm²], and C11 [P2, donor age: 26 years, ECD = 3321 cells/mm²]). The hierarchical clustering of gene signatures was determined by using RT² Profiler PCR Array of EMT and was illustrated as heat maps.

Subpopulation Proven as Effector Cells in the Context of CD Marker

Cell surface markers for CD44⁻ and CD44⁺ cHCECs were evaluated by screening for the expression of 242 cell surface antigens by flow cytometry (Lyoplate); the expression profiles of the CD markers are shown in Figure 9 (just some examples) and Table 3. SP contents of the cHCECs used in Figure 9 and Table 3 are shown in Table 2. The markers that exhibited the low-level expression in both groups are not shown. Protein expression of CD73, CD26, and CD105 was present in CD44⁺ SPs, yet completely absent in CD44⁻ SPs. On the other hand, CD166, which was used as a representative marker for HCEC-derived cells, was observed in nearly all SPs, irrespective of the expression and the intensity of CD44. This finding is consistent with that of Okumura et al.¹¹ who found that CD166 was expressed in both normal cells and in fibroblastic cells. CD markers effective for distinguishing distinct SPs were selected by analyzing those on established cHCECs with a small cell area and a high cell density. The combined analysis of CD markers clearly identified the SP (effector cells) applicable for therapy. 

Figure 9

 

Representative results of BD Lyoplate screenings. Cell-surface markers of cHCECs listed in Table 2 were evaluated by BD Lyoplate screening, and the representative results are shown.

Table 2

 

Subpopulation Contents of the Cultured HCECs Used in Figure 9 and Table 3

 

Table 3

 

The Expression Profiles of CD Makers

 

Factors Influencing the Proportion of Effector Cells in cHCECs

In the absence of a practical scientific index to define SPs, the only way to qualify variations from culture to culture is via the microscopic observation of morphology. However, we have now developed a method to quantitate the proportion of effector SP in cultures (i.e., the “E-ratio”). This method definitively clarifies the relation between donor ages, donor endothelial cell density, and death-to-preservation time with the effector proportions in cHCECs (Fig. 10). Interestingly, we found that only the donor age has an association with the E-ratio. 

Figure 10

 

Correlation of the proportion of effector SP (E-ratio) with donor cornea conditions. Human corneal endothelial cells from 15 donors (donor age range: 10–71 years, ECD: 2526–3879 cells/mm², death-to-preservation time: 4 hours, 14 minutes–24 hours, 12 minutes) were cultured and the ratios of effector SP (G1) at passage 1 were analyzed as in Figure 1. E-ratios were plotted with donor ECD (A), age (B), or death-to-preservation time (C). The coefficients of determination were calculated (R²).

Limiting Factors That Affect the E-Ratio

As mentioned above, CD44 plays diverse critical roles in CST, the maintenance of stem-cell features, and the induction of CSCs. Hence, it is relevant to investigate the primary factors associated with CD44 expression on cHCECs. During the extended primary culture, CD44 expression gradually decreased (Fig. 11A), thus indicating a link between this reduction and differentiation to a mature type. Even at the sixth passage, the addition of Rho-associated protein kinase (ROCK)–inhibitor Y-27632 throughout a 35-day culture strikingly increased the E-ratio from 1.2% to 52.3%, although no morphologic difference could be recognized. The presence of ROCK-inhibitor Y-27632 during a 47-day culture also increased the E-ratio. These findings were additionally confirmed with the clear narrowing of the distribution of cell areas, the decrease of the average cell sizes from 258 to 216 μm², and the increase of cultured cell density from 2229 to 2582 cells/mm² (Fig. 12). 

Figure 11

 

(A) Increase of E-ratios during cultivation. Human corneal endothelial cells from a 29-year-old donor were plated in five separate wells of a 24-well plate, and collected once per week during cultivation at passage 0. The content of SPs was analyzed as in Figure 1. (B) Increase of E-ratio by addition of ROCK-inhibitor Y-27632. The cHCECs from an 18-year-old donor at passage 5 were seeded to two separate wells, either in the presence or absence of ROCK-inhibitor Y-27632. After 62-days of cultivation, the content of SPs in collected cells was analyzed as in Figure 1.

Figure 12

 

Area distribution analysis of cHCECs. Cultured HCECs from a 29-year-old donor at passage 1 were seeded to two separate wells and cultured either in the presence or absence of ROCK-inhibitor Y-27632. After 47 days of cultivation, the cells were washed with PBS(-) to clarify the cell borders, and then phase contrast images were captured at a 20-fold objective (i.e., nine fields per well). The nine images from each lot were then combined and analyzed; the area of each cell was monitored by BZ-II Hybrid Cell Count Software.

Discussion

Expansion of cHCECs from donor corneal endothelium could provide a practical tool for regenerative medicine, that is, cHCEC therapy.^30–35 Cultured HCECs expanded in an in vitro culture system can be composed of multiple SPs of cHCECs, including those with CST. Cultured cells also tend toward karyotype changes.¹⁰ Thus, the quality of cHCECs should be carefully monitored from the aspect of their use in the clinical setting. 

Not only were there morphologic variations in cultured cells, but also the composition of SPs in cHCECs varied greatly between cultures (Table 1), even under identical culture protocols. One possible reason for those variations might be the differences in donor age,^18,19 while another possible reason might be the difference of culture passages, as is the case for MSC culture in regard to stem cell markers CD29, CD49e, CD73, CD90, CD105, and CD166.²⁶ Cornea donor ages and the frequency of effector cells showed a significant inverse correlation (Fig. 10), in contrast to the positive correlation in karyotype aneuploidy.¹⁰ 

To date, no HCEC-specific cell surface markers have been elucidated, although two groups have reported the expression of CD markers.^11,12 The effector cell did not show any sign of surface CD200. Okumura et al.¹¹ have reported that CD340, CD166, and CD98 are elevated in HCECs of nonfibroblastic phenotype, while CD9, CD49e, CD44, and CD73 are the indices of fibroblastic phenotypes. However, they have not elucidated a combination of CD markers to qualify the SPs. In our aim toward application in the clinical setting, we applied the combination of CD markers with a concern toward CSCs owing to the EMT that frequently occurs in HCEC cultures. 

A multifunctional CD44 displays diverse functions in many cells by regulating stem cell behavior, including self-renewal and differentiation, and detects changes in ECM in response to changes in cell-to-cell and cell-to-ECM interactions, cell trafficking, and homing and signal transduction events, thus enabling the pliant responses to the tissue environment.^25,36 In this study, the effector cells were found to express CD166⁺CD105⁻CD44^−∼+/− CD26⁻CD24⁻, but not CD200. The presence of other SPs with CD166⁺CD105⁻CD44^+∼+++ (CD26 and CD24 either + or −) was also confirmed. In addition, depending on the culture variations, such as the presence of ROCK-inhibitor Y-27632 and the extension of culture periods, CD44 expression was likely reduced, thus resulting in the increase of E-ratios. Future extensive investigation is vital to determine the role of CD44 in the differentiation pathway to mature HCECs. Downstream factors of CD44 include Ras homolog gene family member, A (RhoA) and matrix metalloproteinase (MMP)-2, which are responsible for the organization of tubulin and actin cytoskeleton and the formation of cellular pseudopodia.³⁷ 

Epithelial-mesenchymal transition is aberrantly induced in many diseases. Cultured HCECs have an inclination toward CST into EMT, and through EMT, cell-to-cell adhesion is reduced. Cells undergoing EMT frequently obtain stem cell–like properties,²³ including that induced by transforming growth factor-β (TGF-β). It is well known that EMT shares close linkages to the generation of cancer stem cells or their niches during phenotypic conversion,²³ and this prompted us to investigate the presence of heterogeneous SPs in terms of CD44. The identification of CSCs is most commonly performed using CD133, CD44, CD24, and CD90.^38–40 CD26 accompanies the increased expression of CD133, featuring CD26 as a CSC marker.⁴¹ 

Considering the phenotypic heterogeneity due to genetic instability of cHCECs, the selected combination of markers, including auxiliary CSC markers, may appropriately qualify the SP most suitable for cell therapy. McGowan et al.¹³ have reported the identification of corneal endothelial stem cells; however, they have never been isolated for culture. 

CD44 is the benchmark for distinguishing differentiated cHCECs from either undifferentiated cHCECs or cHCECs with CST. CD44 plays a critical role as a major adhesion molecule of the ECM and in TGF-β–mediated mesenchymal phenotype induction, and loss of CD44 reportedly arrests those changes.⁴² CD44 ablation increases metabolic flux to mitochondrial respiration and concomitantly inhibits entry into glycolysis. Such metabolic changes, induced by CD44 ablation, reportedly result in marked depletion of cellular reduced glutathione.⁴³ Moreover, HDAC1 reportedly regulates the activation of the miRNA-34a/CD44 axis and the downstream factors of CD44, including RhoA and MMP-2.³⁷ 

The findings of this present study demonstrated that cHCECs are composed of a hierarchy of cells, and that one specific cHCEC SP with surface expression of CD166⁺, CD105⁻, CD44⁻, CD26⁻, and CD24⁻ is able to be reproducibly cultured without CST or EMT. The combination of CD markers described above qualified the SP, showing the inclination toward mitochondria-dependent oxidative phosphorylation but not anaerobic glycolysis. Thus, the identified effector SP proved to have no sign of karyotype abnormality, thereby opening the pathway toward providing cultured cells in a safe and stable manner for therapy, namely, transplanting cHCECs into the anterior chamber in the form of a cell suspension for bullous keratopathy. 

Acknowledgments

The authors thank Michio Hagiya as well as Yuki Hosoda and Shunsuke Watanabe for technical assistance, Yoko Hamuro, Keiko Takada, and Tomoko Fujita for secretarial assistance, and John Bush for his excellent review of the manuscript. Finally, the authors wish to extend their sincere thanks to Naoki Okumura for his helpful discussion. 

Supported by The Highway Program for Realization of Regenerative Medicine from Japan Agency for Medical Research and Development, AMED and JSPS KAKENHI Grant Numbers JP26293376. 

Disclosure: J. Hamuro, None; M. Toda, None; K. Asada, None; A. Hiraga, None; U. Schlötzer-Schrehardt, None; M. Montoya, None; C. Sotozono, M. Ueno, None; S. Kinoshita, None