Production of Homogeneous Cultured Human Corneal Endothelial Cells Indispensable for Innovative Cell Therapy

Abstract

Purpose: Cultured human corneal endothelial cells (cHCECs) are anticipated to become an alternative to donor corneas for the treatment of corneal endothelial dysfunction. The aim of this study was to establish proper culture protocols to successfully obtain a reproducibly homogeneous subpopulation (SP) with matured cHCEC functions and devoid of cell-state transition suitable for cell-injection therapy.

Methods: The presence of SPs in cHCECs was investigated in terms of surface cluster-of-differentiation (CD) marker expression level by flow cytometry, as combined analysis of CD markers can definitively specify the SP (effector cells) conceivably the most suitable for cell therapy among diverse SPs. The culture conditions were evaluated by flow cytometry in terms of the proportion (E-ratio) of effector cells designated by CD markers.

Results: Flow cytometry analysis identifying CD44⁻CD166⁺CD133⁻CD105⁻CD24⁻CD26⁻ effector cells proved convenient and reliable for standardizing the culture procedures. To ascertain the reproducible production of cHCECs with E-ratios of more than 90% and with no karyotype abnormality, the preferred donor age was younger than 29 years. The continuous presence of Rho-associated protein kinase (ROCK)-inhibitor Y-27632 greatly increased the E-ratios, whereas the presence of transforming growth factor-beta/Smad-inhibitor SB431542 greatly reduced the number of recovered cHCECs. The seeding cell density during culture passages proved vital for maintaining a high E-ratio for extended passages. The continuous presence of ROCK-inhibitor Y-27632 throughout the cultures greatly improved the E-ratio.

Conclusions: Our findings elucidated the culture conditions needed to obtain reproducible cHCECs with high E-ratios, thus ensuring homogeneous cHCECs with matured functions for the treatment of corneal endothelial dysfunction.

The proliferative potential of human corneal endothelial cells (HCECs) is limited,^1–3 and the pathologically irreversible damage to the corneal endothelium leads to corneal endothelial dysfunction and the loss of corneal transparency.^4–6 A recent study⁷ reported that maintaining cultured HCECs (cHCECs) ex vivo for a long period is extremely difficult, and that the repeated passages of cHCECs were frequently accompanied with cell-state transition (CST) into a senescence phenotype, epithelial-mesenchymal transition (EMT), and fibroblastic cell morphology. Hence, corneal transplantation is the only available therapeutic pathway in such cases. From this aspect, it is important to keep in mind the pioneering previous studies by colleagues of Joyce et al.,^8–15 in which the presence of heterogeneity in HCEC cultures was reported. 

The current lack of a standardized culture method for ex vivo expansion of HCECs certainly represents a hurdle in cell therapy for corneal endothelial dysfunction. Many research groups, including Okumura et al.,¹⁶ have put enormous effort into the topic of cHCECs for regenerative medicine and have reported some interesting findings. As was pointed out in their study, a serious issue related to tissue-engineering therapy for corneal endothelial dysfunction is the complete lack of a concept to signify, biochemically, the most homogeneous cHCECs with sufficient qualifications of their composites (i.e., no protocol to date has been established for culturing HCECs with regard to homogeneity for clinical use). 

Hence, in light of the absence of any type of index of culture assessment and the current inadequate insights on the presence of heterogeneous subpopulations (SPs) in cHCECs, the subsequent conclusions stated in recent studies may have been reached via confusion with regard to the findings and now may be in need of scientific reappraisal. 

In a series of recent studies,^17–23 we reported the existence of diverse SPs in cHCECs and identified a cluster of differentiation (CD)166⁺/CD44⁻/CD105⁻/CD26⁻/CD24⁻/CD133⁻ SP as the most suitable “effector” cell population for clinical applications in cell infusion in the anterior chamber. In addition, we developed a method to quantify the proportion of effector cells in cultures, the so-called “E-ratio,” on the basis of the above-described cell surface markers, and defined donor age and other parameters that either up- or downregulate the E-ratio. However, to date, no standardized and reproducible cultivation procedures have been reported. 

Thus, the purpose of this present study was to describe detailed culture conditions to reproducibly culture HCECs with high E-ratios, thereby enabling obtainment of the utmost-homogeneous cHCECs with matured functions to serve as a novel tool for the treatment of corneal endothelial dysfunction. In this present study, we also present a protocol for the standardized and reproducible cultivation of endothelial “effector” cells for clinical application. Therefore, this study is of major clinical relevance and practical importance for future treatment strategies. 

Materials and Methods

Reagents

Rho-associated protein kinase (ROCK)-inhibitor Y-27632 was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan) and SB431542 was obtained from Tocris Bioscience (Bristol, UK). 

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 from 20 human donor corneas obtained from SightLife Surgical (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. 

The 29 male and 12 female donors ranged in age from 14 to 71 years. 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 history of chromosomal abnormality. 

Cell Cultures of HCECs

Human corneal endothelial cells obtained from a total of 41 human donor corneas at distinct ages were cultured according to published protocols, with some modifications.¹⁶ Briefly, the Descemet's membranes with the corneal endothelial cells 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 (FBS), 5 ng/mL epidermal growth factor (Life Technologies), 20 μg/mL ascorbic acid (Sigma-Aldrich Corp., St. Louis, MO, USA), 200 mg/L calcium chloride (Sigma-Aldrich Corp.), 0.08% chondroitin sulfate (Wako Pure Chemical Industries, Ltd., Osaka, Japan), and 50 μg/mL gentamicin. Mesenchymal stem cell (MSC)-conditioned medium was prepared as previously described.²⁴ The HCECs were cultured using MSC-conditioned medium at 37°C in a humidified atmosphere containing 5% CO₂, and the culture medium was changed twice a week. The HCECs were passaged at ratios of 1:3 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 PBS(−), 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). 

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 (CKX41; Olympus Corporation, Tokyo, Japan). 

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% 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 as follows: FITC-conjugated anti-human CD26 mAb, phycoerythrin (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), and allophycocyanin APC-conjugated anti-human CD105 (eBioscience, Inc., San Diego, CA, USA). After washing with FACS buffer, the HCECs were analyzed by use of a BD FACSCanto II Flow Cytometry System (BD Biosciences). 

Multiplex Analysis of Cytokines in MSC-Conditioned Medium

The cytokine levels of the MSC-conditioned mediums were analyzed by use of x-MAP Technology (Luminex Corporation, Austin, TX, USA) incorporated into the Bio-Plex 200 (Bio-Rad Laboratories, Inc., Hercules, CA, USA) suspension array system with the Bio-Plex Human 27-plex panel kit (Bio-Rad Laboratories) according to the manufacturer's instructions. The measured cytokines were as follows: interleukin (IL)-1β, IL-1ra, IL-2, IL-4, IL-5, IL-6, IL-7, IL-8, IL-9, IL-10, IL-12p70, IL-13, IL-15, and IL-17A, BFGF, eotaxin, granulocyte colony-stimulating factor, granulocyte macrophage colony-stimulating factor, IFN-γ, IFN-induced protein-10 (IP-10), monocyte chemotactic protein-1 (MCP-1), macrophage inflammatory protein-1α (MIP-1α), MIP-1β, platelet-derived growth factor-BB, regulated-upon-activation normal T-cell expressed and secreted (RANTES), TNF-α, and VEGF. Standard curves for each cytokine (in duplicate) were generated using the reference cytokine concentrations supplied in the above-detailed kit and were used to calculate the cytokine concentrations in the MSC-conditioned mediums. 

Results

Ascertainment of Cell-Culture Quality by Composite SP Variances in cHCECs

Human corneal endothelial cells from donors of different ages were cultured according to the method of Okumura et al.¹⁵ in the presence of TGF-β signaling-blockade SB431542 in MSC-conditioned medium, and the surface expression of CD44, CD166, CD24, CD26, and CD105 were characterized (Fig. 1B). The analysis was also performed for cHCECs prepared in a cell-processing center. Representatives are summarized in Figure 1 (C1–C4) and Table 1 (C1–C7), together with the phase-contrast microscopy (Fig. 1C [C2, C4]). At first observation, it quickly became evident that the cHCECs contained a variety of SPs, as we reported in a previous study.²² It is also of note that those cHCECs elicited a strikingly distinct proportion of SPs defined by cell-surface CD markers, even when cultured along the lines of the protocol proposed by Okumura et al.¹⁵ The E-ratio defined by the proportion of SPs with CD44⁻CD166⁺CD24⁻CD26⁻CD105⁻ varied from 0.2% (C5, G1) to 31.2% (C7, G1 in Table 1). As described elsewhere, CD44 expression decreased in accordance with the differentiation of cHCECs to matured cHCECs.²² The presence of SPs expressing either CD24 or CD26 in some cultures quickly raised the alarm of the presence of some SPs with remarkable karyotype abnormality, such as sex chromosome loss, trisomy, or translocation, because we confirmed the presence of karyotype abnormality in cHCEC SPs expressing CD24 or CD26.¹⁸ Thus, most of those cells are not suitable for transplantation in the clinical setting. The proportion of CD24⁺ cells and CD26⁺ cells ranged from 0.3% up to 96.6% (Fig. 1 [BR2; C2 vs. C4]) and that of CD26⁺ cells from 0.3% up to 20.7% (Table 1 [Gd; C2 vs. C3]), respectively. The most striking variation was observed in the expression of CD44⁺⁺⁺ cells showing the highest ratios of more than 80% (Fig. 1B [C2, G2+G3]), although phase-contrast microscopy revealed that the visible phenotypes were nonfibroblastic with the characteristic polygonal contact-inhibited shape and monolayer (Fig. 1A [C1]). Surprisingly, both ZO-1 and Na⁺/K⁺-ATPase, well-known markers of HCECs, were stained for CD24⁺, CD26⁺, or CD44⁺⁺⁺ SPs (Fig. 1C [C4]). This raised the alarm that the morphologic judgment only does not suffice in the distinction of heterogeneous SPs present in cHCECs. This immediately forced us to precisely reassess the culture protocols and processes from the aspect of the proportion of effector-cell SP (E-ratio) to avoid the conversation in this field becoming controversial. 

Figure 1

 

(A) Phase-contrast images of the cHCECs: C1, HCECs from a 50-year-old donor; C2, HCECs from a 57-year-old donor; C3, HCECs from a 29-year-old donor; and C4, HCECs from a 58-year-old donor. (B) Representative results of the expression of CD166, CD24, CD44, CD105, and CD26. The expression of CD166, CD24, CD44, CD105, and CD26 was analyzed by a FACSCanto II Flow Cytometry System (BD Biosciences). After gating for CD166⁺CD24⁻ (R1) or CD166⁺CD24⁺ (R2), the following five SPs were defined: (1) CD166⁺CD24⁻CD44^−/+CD105^+/− SP (gate [G] 1), (2) CD166⁺CD24⁻CD44⁺⁺CD105⁺ SP (G2), (3) CD166⁺CD24⁻CD44⁺⁺⁺CD105⁺ SP (G3), (4) CD166⁺CD24⁺CD44⁺CD105⁺ (G4), and (5) CD166⁺CD24⁺CD44⁺⁺CD105⁺ SP (G5). CD26⁺CD44⁺⁺ SP (Gd) were also defined with the CD24 and CD44 plot. Note: C2 and C4 both correspond to the numbers shown in (A). (C) The expression of Na⁺/K⁺ ATPase, ZO-1, and Claudin-10 on cHCECs. Cultured human corneal endothelial cells C2 and C4 in (A) were stained with antibodies against Na⁺/K⁺ ATPase, ZO-1, Claudin-10, or CD26, as described in Materials and Methods.

Table 1

 

Subpopulation Contents of the cHCECs

 

No Critical Role of MSC-Conditioned Medium in Improving the E-Ratio

The findings in the report by Nakahara et al.²⁴ prompted us to apply human bone marrow–derived MSCs (MSC-CM) for the maintenance of the cHCECs. In that study, the authors claimed that MSC-CM not only stimulated the proliferation of cHCECs by regulating the G1 proteins of the cell cycle, but also maintained the differentiated phenotypes necessary for the endothelial functions of cHCECs. Considering the variations in SP composites elucidated above, we were forced to reassess that groups reported conclusion in the context of the changes of E-ratios and the composition of SPs between the cultures with or without MSC-CM. 

As shown in Table 2, the MSC-CM produced varied greatly from lot to lot with regard to their content of cytokines, such as senescence-associated secretory pathway (SASP)-related IL-6, IL-8, MCP-1, and VEGF. Especially those derived from distinct bone marrow donors showed a great difference in the content of these SASPs (Table 2). It should be kept in mind that no parallel was found with regard to the levels of those four cytokines. 

Table 2

 

Cytokine Contents of the MSC-Conditioned Medium

 

In addition to the above-described differences in their quality, different lots of MSC-CM manifested distinct effects on E-ratios and the composition of SPs (Table 3). The E-ratios ranged from 8.7% to 26.4%, whereas the proportion of CD44⁺⁺⁺ (G3) ranged from 44.0% to 22.8%. The proportion of CD26⁺ cells (Gd) ranged from 5.5% to 1.2%. Taken together, these findings show that MSC-CM lot D more heavily spoiled the cHCEC culture than lot A with regard to the presence of SPs other than the effector SP. 

Table 3

 

Contents of the HCEC SPs in HCECs Cultured in the Presence or Absence of MSC-CM From Different Donor MSCs

 

Finally, we investigated whether or not the presence and absence of MCM-CM would influence the E-ratios (Fig. 2). The E-ratios of both groups, either in the presence or absence of MSC-CM did not show any statistically meaningful differences (P = 0.934, Student's t-test). Thus, it can be safely concluded that the addition of MSC-CM did not play a critical role in improving the culture to gain the effector cell SP with the differentiated phenotypes necessary for the endothelial functions of cHCECs. 

Figure 2

 

Human corneal endothelial cells from 10 donors were cultured in MSC conditioned or nonconditioned medium. Gate 1 corresponding to the gated SP presented in Figure 1B (= E-ratio) at P0 were analyzed as in Figure 1B. The mean percentages were 72.7% (MSC conditioned) and 71.8% (nonconditioned), respectively; P = 0.934.

Induction of Morphologically Transformed Cells by TGF-β Signaling Blockades

Cultured HCECs have an inclination toward CST into a senescence phenotype, EMT, and fibroblastic cell morphology. Regarding the pluripotent functions of TGF-β and its existence in aqueous humors, we postulated that TGF-β may function to interfere with the acquisition of an invasive phenotype into the extracellular matrix (ECM). To the contrary, TGF-β reportedly can initiate and maintain EMT in a variety of biological and pathologic systems.^25,26 However, the same growth factor is the key signaling molecule for EMT, and the role of TGF-β as a key molecule in the development and progression of EMT is well studied.^25–28 

The continuous addition of SB431542, a selective inhibitor of the TGF-β receptor, at a concentration of 1 μM in the culture of cHCECs revealed a significant decrease of the recovered cell numbers of P0 cultured cells, as shown in Figure 3Aa. The observed decrease of cell numbers may reflect the presence of cells with enlarged phenotypes in the presence of SB431542, as illustrated in the phase-contrast microscopy images in Figures 3B and 3C. Human corneal endothelial cells prepared from another eight donors were cultured without SB431542 at a cell-processing center, and endothelial cell densities (ECDs) were evaluated. The mean cell density was 2246 cells/mm², indicating the high reproducibility of the superiority of the culture in the absence of SB431542. In addition, the E-ratio was lowered from 56.1% to 6.2%, whereas the proportion of CD24⁺ cells increased to 87.0% from 35.5% and that of CD44⁺⁺⁺ to 37.8% from 4.6% (representative experiments shown in Figs. 3B, 3C). The results clearly indicate that the blockade of TGF-β signaling induced morphologically transformed cells in cHCECs. The increase of CD44⁺⁺⁺CD24⁺ SP in cHCECs in the presence of SB431542 means an increase of the SPs with the high risk of karyotype aneuploidy.¹⁸ The changes of gene signatures, followed by RT-PCR of cHCECs shown in Figure 3B, also clarified the upregulation of CD44, CD24, IL-8, and IGFBP3, and the downregulation of collagen 4A1, 4A2, and 8A2 (Asada et al., unpublished data, 2015), thus supporting the results deduced from the SP analysis by flow cytometry. 

Figure 3

 

(Aa) Human corneal endothelial cells from 16 donors were cultured with or without SB431542, and ECDs were calculated on the basis of the number of recovered cells. The means of ECDs were 2432 cells/mm² (with SB) and 975 cells/mm² (without SB), respectively; P < 0.001. (Ab) Human corneal endothelial cells prepared from another eight donors were cultured without SB431542 in a cell-processing center, and the ECDs were evaluated as in Aa. The mean ECD was 2246 cells/mm². (B, C). The E-ratio was reduced by addition of SB431542. Cultured human corneal endothelial cells at passage 0 from a 36-year-old donor were seeded to two separate wells, and cultured either in the presence (B) or absence (C) of SB431542. The expression of CD166, CD24, CD44, CD105, and CD26 was analyzed as in Figure 1B.

Influence of Seeding Cell Density on E-Ratios in cHCECs

Under the culture conditions without SB431542, we were able to obtain stable cHCECs with a more reduced proportion of CD24⁺ cells than in the previously described cultures that included SB431542. Consequently, the quality of cHCECs are able be monitored mostly by the E-ratio and the proportion of CD44⁺⁺⁺ cells. Before finalizing the protocol of culturing HCECs from the aspect of homogeneity for clinical use, we investigated the influence of the seeding cell density on E-ratios and the proportion of CD44⁺⁺⁺ cells (Table 4). To ascertain the correct conclusion, we investigated three lots of cHCECs in which the E-ratios were different; namely 54.0% (culture passage 1), 77.3% (culture passage 0), and 93.4% (culture passage 1). In all three groups, the higher the seeding cell density, the lower the reduction of E-ratio after following passages. However, the group with high E-ratios before culture exhibited the lowest reduction of these ratios. At a seeding cell density between 750 and 1000 cells/mm², the culture of the next passage works well, whereas at a lower seeding cell density, such as 200 cells/mm², most of the cultured cells entered into CST and exhibited a poor proliferative behavior. However, it turned out that this poor proliferative behavior mostly depends on the E-ratios of the seeded cell populations. Specifically, we found that the seeded cells with an E-ratio greater than 90% showed a good increase in cell numbers at the following passages, even at a seeding cell density as low as 200 cells/mm², and reached a cell density comparable to the cultures that began with a seeding cell density between 750 and 1000 cells/mm². Surprisingly, even the group with a seeding density of 200 cells/mm² showed the proportion of CD44⁺⁺⁺ cells to be only 1.7% (G5 in Table 4), compared with that of 0.6% (G5 in Table 4) in the group with a seeding density of 750 cells/mm². However, the shift to increased proportion of CD44⁻ to CD44⁺⁺ was prominent in the groups with the lower seeding cell density. 

Table 4

 

Subpopulation Contents of the HCECs Seeded at Various Cell Densities

 

Culture Protocol of HCECs to Semi-Homogeneity for Clinical Use

In consideration of the above-mentioned observations, the culture protocol of HCECs to obtain the utmost homogeneity for clinical use was tentatively fixed as follows: (1) optimal donor age being between 7 and 29 years,²² (2) seeding cell density after passage 1 greater than 750 cells/mm² (note: further detailed investigation is needed to ultimately determine the recommended seeding cell density with regard to the E-ratio of cHCECs of each passage), and (3) no addition of SB431542 (a selective inhibitor of the TGF-β receptor) to use the beneficial effect of TGF-β signaling. In addition, we further revised the protocols proposed by Okumura et al.²⁹ The addition of ROCK-inhibitor Y-27632 was changed from the addition only at the beginning of new passages to every 3 days throughout the entire culture period to produce an efficient differentiation to CD44⁻ SP. Under this culture condition, Y-27632 dramatically induced the differentiation of cHCECs to the matured state with reduced expression of CD44 (Fig. 4). The downstream factors of CD44 reportedly includes RhoA, the target of Y-27632.²⁵ In accordance with this expectation, the continuous addition of Y-27632 dramatically increased the E-ratios and reduced the proportion of CD24⁺ or CD26⁺ SPs, as illustrated in Figure 4. E-ratios were approximately 90% or greater and the contaminating SPs were scarcely found in cHCECs. Under these conditions, we frequently observed high-quality cHCECs even at the fifth passage or from the donors 57 or 71 years of age (Fig. 4). 

Figure 4

 

E-ratios increased by addition of ROCK-inhibitor Y-27632. (A) Cultured human corneal endothelial cells from a 57-year-old donor at passage 0 were seeded to two separate wells, and cultured either in the presence or absence of Y-27632. The expression of CD166, CD24, CD44, CD105, and CD26 was analyzed as in Figure 1B. (B) Cultured human corneal endothelial cells from two different donors (both 22 years old) were cultured in the presence of Y-27632, and the expression of CD166, CD24, CD44, CD105, and CD26 was analyzed as in Figure 1B. (C) Cultured human corneal endothelial cells from a 71-year-old donor were cultured in the presence of Y-27632, and the expression of CD166, CD24, CD44, CD105, and CD26 was analyzed as in Figure 1B.

Discussion

To date, most researchers are used to applying just the rhetorical term “cultured human corneal endothelial cells” merely on the grounds that they are derived from corneal endothelium tissues, regardless of the fact that the biochemical features are undefined. Thus, this situation has caused long-standing confusion among diligent cHCEC researchers, as the biochemical features are an important aspect that needs to be considered. 

In a concomitant study, we defined the cHCEC SP with CD133, CD105, CD90, CD44, CD26, CD24, HLA-DR, and DQ “negative” and CD166, HLA-ABC, and PD-L1 “positive” as effector cells that ensure a safe and stable application to regenerative medicine.²² To ascertain the reproducible production of cHCECs with E-ratios of more than 90% and with no karyotype abnormality, the continuous presence of ROCK-inhibitor Y-27632 throughout the culture period, and in the absence of SB431542 (a selective inhibitor of the TGF-β receptor) was recommended. 

CD44 actively contributes to the maintenance of stem-cell features, and the functional contribution of CD44 reportedly relies on its particular communication skills with neighboring molecules, adjacent cells, and the surrounding matrix.³⁰ CD44 is the key to distinguishing differentiated cHCECs from either undifferentiated or cHCECs with CST. Thus, it is important to elucidate the factors that control the levels of CD44 expression on cHCECs to fix the culture protocols to gain a final product with an E-ratio of more than 90%. A multifunctional CD44 displays diverse functions in many cells with regard to regulating stem-cell behavior, including self-renewal and differentiation, and detects changes in the ECM in response to changes in cell-to-cell and cell-to-ECM interactions, cell trafficking, homing, and signal-transduction events, thus enabling pliant responses to the tissue environment.^31,32 Downstream factors of CD44 include RhoA and MMP2, both responsible for the organization of tubulin and actin cytoskeleton and the formation of cellular pseudopodia.³³ Loss of CD44 reported abrogates those changes.³⁴ CD44 ablation increased metabolic flux to mitochondrial respiration, and concomitantly inhibited entry into glycolysis. The activation of micro RNA (miR)-34a/CD44 axis reportedly regulates the downstream factors of CD4, such as Ras homolog gene family member A (RhoA) and matrix mealloproteinase-2 (MMP2).³³ This pathway may, albeit in part, participate in the reduction of CD44 expression on cHCECs by ROCK-inhibitor Y-27632. In a previous study, we confirmed the upregulation of miR29 in parallel with that of CD44 expression among distinct SPs, as our findings showed that miR29 was most downregulated in effector cells with a CD44-phenotype among cHCEC SPs.³⁵ Interestingly, miR29 reportedly has an EMT-promoting effect.³⁶ 

Reportedly, TGF-β can initiate and maintain the EMT in various biological and pathologic systems.^26,37 Moreover, the findings in those two studies showed that suppression of TGF-β signaling promoted CST under an inflammatory microenvironment, and that MMP2 was activated in the benign tissues with intact TGF-β signaling.^26,37 Although blocking TGF-β signaling does not cause morphologic changes, indicating that TGFβ signaling is not required for differentiation of normal intestinal stem/progenitor cells,³⁸ the suppression of TGF-β signaling in the injured intestinal mucosa reportedly blocks mucosal regeneration by suppressing differentiation, followed by the expansion of an undifferentiated cell population.³⁹ 

At the beginning of this present study, cHCECs exhibited unstable phenotypes with varying expressions of CD44 and CD24 with CST (Fig. 1). These highly CD44-positive SPs, which show elevated levels of MMP2,¹⁷ were used for the experiments by Koizumi et al.⁴⁰ involving a nonhuman primate model. Thus, it is impossible to know which of the SPs used in their study was responsible for the reported results. 

As mentioned above, CD44 is also linked with the miR34/RhoA/MMP2 pathway. It is noteworthy that the only miR capable of discriminating CD44⁻ effector cells from CD44⁺⁺–CD44⁺⁺⁺ SPs was identified as miR34a.³⁵ Consequently, it is conceivable that in the previous experiment of Koizumi et al.,⁴⁰ either CD44⁺⁺⁺ or CD44⁺⁺ SPs with downregulated miR34a used the RhoA/MMP2 pathway activated in the undifferentiated SPs.³³ If this assumption is correct, SB431542 might function to block the EMT of undifferentiated SPs and Y-27632 worked both for the adhesion to the culture dish and for promoting the proliferation of these SPs. This may result in a failure to elevate the most relevant index of cHCEC homogeneity, (i.e., the E-ratio). These interpretations do not exclude the possible effectiveness of other SPs in cell therapy aimed at restoring corneal endothelial dysfunctions by some other unknown pathway. 

In conclusion, detailed culture protocols to reproducibly produce cHCECs with a high proportion of effector cells were established by using the recently developed E-ratio index, thereby enabling the availability of the utmost-homogeneous cHCECs with matured functions for cell-injection therapy for corneal endothelial tissue damage due to Fuchs' corneal endothelial dystrophy, trauma, or surgical intervention. 

Acknowledgments

The authors thank Michio Hagiya, Yuki Hosoda, and Shunsuke Watanabe for their technical assistance, and Yoko Hamuro, Keiko Takada, and Tomoko Fujita for their secretarial assistance. The authors also express their sincere appreciation to John Bush for his valuable assistance in reviewing the manuscript. Finally, the authors thank to Noriko Koizumi and Naoki Okumura for their instructive and helpful discussion. 

Supported by the Highway Program for Realization of Regenerative Medicine from Japan Agency for Medical Research and Development (AMED) and Japan Society for the Promotion of Science (JPS) KAKENHI Grant Numbers JP26293376. 

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