The corneal endothelium is a cell monolayer located at the cornea posterior, separating the stroma from the fluid aqueous humor of the anterior chamber. Its major function is to regulate stroma hydration, thereby maintaining corneal thickness and transparency [1
]. Studies have indicated that the mature human corneal endothelial cells in vivo
are arrested in the G1 phase of the cell cycle and, thus, do not replicate to replace dead or injured cells [2
]. Instead, wound healing occurs through the enlargement and migration of adjacent healthy cells [2
]. When excessive cell loss due to accidental or surgical trauma occurs, this lack of proliferative response may result in endothelial dysfunction, leading to the development of cornea edema and the eventual loss of visual acuity [2
]. Current available treatments of endothelial dysfunction include penetrating keratoplasty (full thickness corneal transplantation) and endothelial keratoplasty. However, these surgical options are limited by a shortage of donor corneas and failure due to immune-mediated rejections [3
]. The reconstruction of the cornea endothelium from autologous cells as a tissue engineered replacement is a promising alternative for treatment and also has potential for use as a cell model in in vitro
ocular toxicology testing [4
Current research focuses on the cultivation of primary human cornea endothelial cells (HCECs). The capacity of HCEC to be cultivated in vitro
has been demonstrated, and cell layers have been cultured on temperature responsive culture dishes [5
], human amniotic membranes [6
] and surfaces coated with extracellular matrix (ECM) components, such as bovine corneal endothelial cell-derived ECM [7
] and laminin-5 [8
]. Proof-of-concept studies in animal models have shown the potential of in vitro
cultured cornea endothelial cells for clinical use [6
]. However, the use of these primary cells is limited by donor availability and their low proliferative capacity. It is difficult to establish a lasting HCEC culture, and aged cells tend to display abnormal morphologies [12
]. Variability exists across different donors, and isolated cells can also be contaminated by stromal keratocytes, which will overtake the culture, due to their higher proliferation rate [12
Therefore, it is necessary to consider alternative cell sources. Umbilical cord mesenchymal stem cells (UMSCs) have been transplanted in the corneas of lumican null mice with improved clarity and increased stromal thickness [13
]. More recently, it has been shown that UMSCs are able to specifically attach to wounded areas of the corneal endothelium and, thus, can potentially be used for the healing of the injured corneal endothelium [14
In this study, human microvascular endothelial cells (HMVECs) are considered and cultured as an alternative cell source for corneal endothelium replacement, because of the similarity in the microvascular and the intra-ocular pressure. In vivo
, normal ocular pressure ranges from 10 mmHg to 20 mmHg [15
], while most segments of the microcirculation have mean vascular pressures of 20 mmHg [16
]. The transplantation of vascular endothelial cells as a corneal endothelium replacement in animal models has been previously reported with optimistic results, indicating their potential for use in clinical applications [17
]. Similar to the CECs, vascular endothelial cells regulate fluid exchange. These cells form tight junctions [19
] and have also been reported to possess Na+
adenine triphosphatase (ATPase), which is essential for the ion transport regulation function for the corneal endothelium. However, the vascular endothelial cells in the previously mentioned studies were not characterized or tested for their resemblance to native corneal endothelium.
The basal side of the native corneal endothelium rest on the Descemet’s membrane, the basement membrane secreted by the endothelial cells. The Descemet’s membrane is a three-dimensional network of nanoscale architecture with fibers and pores [20
]. Studies have shown that the use of topographies on synthetic substrates to mimic the native extracellular matrix can influence and direct cell growth and function [21
]. The differential response of endothelial cells to various synthetic topographical structures has also been previously reported [23
We hypothesized that micro- and nano-topographies of substrates can induce vascular endothelial cells to become corneal endothelial cell-like and could potentially be used for in vitro drug testing and the therapy of the corneal endothelium. HMVECs were grown on poly(dimethylsiloxane) (PDMS) substrates with pillar and well topographies of micro- and nano-meter sizes. The experiments were conducted in two different types of medium to test the interaction between the biochemical environment and the underlying substrate topography. The reconstructed monolayers were evaluated for cell morphology, proliferation, expression of corneal endothelial functional markers and microvilli formation.
3. Experimental Section
3.1. Preparation of Polydimethylsiloxane Substrates
Soft lithography was used to fabricate polydimethylsiloxane substrates with micro- and nano-topographies as previously described [23
]. Patterned master molds were commercially purchased in silicon wafer format. Briefly, poly(methyl methacrylate) (PMMA) (Microresist, MW 35000 g/mol) was first spin-coated on a clean silicon substrate to form a thin PMMA film. The purchased master mold was placed on top of the spin-coated surface, and the imprinting was carried out at 150 °C under a pressure of 60 bar for 10 min. Subsequently, the system was cooled before demolding the silicon master from the imprinted PMMA polymer layer. The PMMA mold was then used for soft lithography. The master molds were cleaned with nitrogen gas and fluorinated with (tridecafluoro-1,1,2,2-tetrahydrooctyl)-1-trichlorosilane (United Chemical Technologies, Pennsylvania, USA). The molds were then washed with 0.01% Triton X (Biorad, Singapore) and blown dry with nitrogen gas. PDMS base and curing agent (Sylgard 184 Silicone Elastomer Kit, Dow Corning, Singapore) were mixed with a 10:1 ratio and degassed in a desiccator for 30 min. The mixture was poured over the PDMS molds, degassed in a desiccator for another 2 h and cured at 60 °C for 12 h. Upon cooling to room temperature, the PDMS substrates were gently peeled off from the master molds.
To verify the surface topographies and to ensure the fidelity of the replication process, the PDMS substrates were sputter coated with gold (JEOL, Japan, JFC Fine Gold Coater) and examined with a scanning electron microscope (SEM, FEI, Japan, Quanta FEG 200 and JEOL, Japan, JSM-5600LV Scanning Microscope, Japan).
Prior to cell seeding, the PDMS were air plasma treated with low radio frequency (RF) power for 15 sec (Harrick Scientific Corporation, New York, USA, PDC-002), cleaned with 70% ethanol and sterilized under UV for 30 min. The substrates were also precoated with 10µg/mL of laminin (Invitrogen, Singapore) overnight.
3.2. Vascular Endothelial Cell Culture on PDMS Substrates
Human neonatal dermal microvascular endothelial cells (HMVEC, Lonza, Singapore) were expanded in endothelial cell growth medium, EGM-2MV (Lonza, Singapore) in standard tissue culture flasks. HMVEC of passage 7 to 11 were used for experimentation after their morphology was inspected (Supplementary Figure 2A
All experiments detailed in this report, with the exception of the BrdU proliferation assay, were carried out in two different types of medium separately. The first is the endothelial cell culture medium EGM-2MV (Lonza, Singapore), while the second consists of 75% of EGM-2MV and 25% of medium, which had been used for the culture if HCEC primary cells [27
]. The HCEC medium was composed of Dulbecco’s Modified Eagle Medium (DMEM, Biological Industries, Bio-Rev Singapore) supplemented with 15% fetal bovine serum (FBS, Gibco, Singapore), 1% penicillin/streptomycin (Gibco, Singapore) and 2ng/mL basic fibroblast growth factor (bFGF, Invitrogen, Singapore). The cells were seeded on the pre-prepared plasma-treated patterned PDMS substrates and unpatterned PDMS controls at densities of 2500 cells/cm2
for the BrdU cell proliferation assay in EGM-2MV, 40,000 cells/cm2
for other experiments in EGM-2MV and 45,000 cells/cm2
for other experiments in medium B. The HMVECs proliferate slower when grown in medium B. Therefore, a higher seeding density was used for experiments in medium B to ensure that the cells were exposed to the substrates for the same culture period. The cells were suspended in 1 mL of culture medium in 24 well plates and incubated at 37 °C and 5% CO2
. The medium was changed on alternate days, and each experiment was ended and analyzed after 7 days.
3.3. Immunofluorescence Staining of ZO-1 and Na+/K+-ATPase, BrdU Proliferation Assay
The samples were fixed and stained following standard immunofluorescence staining protocol after 7 days of culture. Generally, the cells were fixed with 4% paraformaldehyde (Sigma Aldrich, Singapore) and permeabilized with 0.1% Triton X-100. They were subsequently blocked with 1% bovine serum albumin (BSA) and 10% goat serum for 1 hour at room temperature. Then, the samples were incubated with primary antibodies at 4 °C overnight, followed by an hour incubation with the secondary antibody, anti-mouse IgG Alexa Fluor 546 (Invitrogen, Singapore) diluted 1:750 at room temperature. The cells were labeled for ZO-1 or Na+/K+-ATPase, while the nucleus was counterstained with 4’,6-diamidino-2-phenylindole (DAPI). All samples were mounted onto coverslips using ProLong Gold Antifade mounting medium (Invitrogen, Singapore). The stained samples were viewed with an epifluorescence microscope (Leica, Singapore, DM IRB) and analyzed using ImageJ (National Institute of Health, Bethesda, MD, USA).
The primary antibody used for ZO-1 staining was the Mouse IgG Anti-ZO-1 antibody (ZYMED Laboratories, Invitrogen, Singapore), diluted 1:30. The images obtained from ZO-1 staining were used for the measurement of cell area, cell area coefficient of variance (CV), cell circularity and the hexagonal shape factor (HSF). CV of the cell area is defined as the ratio of standard variation to the mean and is presented as a percentage. Cell circularity is defined as 4π × (area/perimeter2). A perfect circle will return a value of 1.0. As the number approaches 0.0, the shape becomes increasingly elongated. HSF is defined as the absolute value of (perimeter2/area-13.856), where 13.856 is the shape factor of a regular hexagon. For each pattern, at least 250 cells were analyzed for each replica and each pattern type has 3 replicas.
Samples that were stained for the Na+/K+-ATPase pump followed the standard protocol, as described previously, but without the permeabilization step. The primary antibody used is the mouse IgG anti-Na+/K+-ATPase (Santa Cruz Biotechnology, Texas, USA) diluted 1:40. Random areas of each sample were photographed for analysis.
For cell proliferation studies, the cells were seeded at a low density to reduce cell-cell contact, which could inhibit cell proliferative ability. At the termination of the experiment, the samples were incubated with BrdU labeling reagent (Sigma Aldrich, Singapore) for 4 h before fixing with 4% paraformaldehyde and permeabilized with 0.1% Triton X-100 for 15 min. Next, incubation in 4N hydrochloric acid was carried out for 10 min at room temperature. The samples were stained according to the previously described standard immunofluorescence staining protocol. The primary antibody used is the mouse anti-BrdU (Developmental Studies Hybridoma Bank, Iowa, USA) diluted 1:20,000. Images of random regions of each sample were photographed and analyzed with ImageJ to obtain the number of BrdU incorporated nuclei and the total number of nuclei. BrdU incorporation percentage was then calculated as the percentage of BrdU incorporating nuclei with respect to the total number of nuclei. A minimum of 500 cells were analyzed per replica, and each sample has 3 replicas.
3.4. Scanning Electron Microscopy of HMVECS on Different Topographies
On the 7th day of culture, the samples were fixed with 2.5% glutaraldehyde (Fluka, Singapore). The cells were then dehydrated through graded ethanol solutions. A final dehydration in 100% ethanol solution was carried out three times for 5 min each before the samples were subjected to critical point drying (Balzers, Hudson, NH, USA, Critical Point Dryer 030). The samples were then gold coated by ion sputtering (JEOL, Japan, JFC 1600 Fine 270 Gold Coater, 90 s, 10 mA) before SEM examination (SEM, FEI, Japan, Quanta FEG 200, HV mode) at an accelerating voltage of 10 kV.
3.5. Data Analysis
For every experiment that was conducted, there were 3 replicas for each type of substrate topography. All data are presented as the mean ± standard deviation (SD), unless otherwise specified. One factor analysis of variance (ANOVA) with repeated measures and Bonferroni post-test correction were used to analyze the statistical significance where indicated. Two-way ANOVA tests with repeated measures were used to analyze the interaction effects between samples grown in medium B and samples grown in EGM-2MV. The significance level was set to be p = 0.05.