How Do Stem Cells Work? How do Stem Cells Heal? All the Ways Stem Cells Heal Tissue

Stem cells are known to work via a variety of methods to heal tissue depending on where the stem cells come from and into which tissue it is being placed. 

Here is a general list of all the Ways Stem Cells heal tissues or create new tissue. 


1. Stem cells have long been known to have the ability to differentiate (ie become/transform) into “good/normal” tissue. Embryonic stem cells have been fraught with difficulties in coaxing them into “good/normal” tissue and there are major ethical issues.
Adult stem cells turn out to be a more robust and reliable source of cell to heal damaged cells. These adult stem cells, such as from fat (adipose derived) or bone marrow, have the ability to transform into different cells. This ability to transform into another type of cell depends on where the cell is placed and what other growth factors are around. 


2. Stem cells secrete growth factors which can help heal damage cells.


3. New research shows that the mere fact that a stem cell touches or even pass by another cell, stimulates the other cell or damaged cell to behave better. 


More information below. 
SLC


Human adipose-derived stem cells support the growth of limbal stem/progenitor cells

Abstract

The most efficient method to expand limbal stem cells (LSCs) in vitro for clinical transplantation is to culture single LSCs directly on growth-arrested mouse fibroblast 3T3 cells. To reduce possible xenobiotic contamination from 3T3s, primary human adipose-derived stem cells (ASCs) were examined as feeder cells to support the expansion of LSCs in vitro. To optimize the ASC-supported culture, freshly isolated limbal epithelial cells in the form of single cells (SC-ASC) or cell clusters (CC-ASC) were cultured using three different methods: LSCs seeded directly on feeder cells, a 3-dimensional (3D) culture system and a 3D culture system with fibrin (fibrin 3D). The expanded LSCs were examined at the end of a 2-week culture. The standard 3T3 culture served as control. Expansion of SC-ASC showed limited proliferation and exhibited differentiated morphology. CC-ASC generated epithelial cells with undifferentiated morphology in all culture methods, among which CC-ASC in 3D culture supported the highest cell doubling (cells doubled 9.0 times compared to cells doubled 4.9 times in control) while maintained the percentage of putative limbal stem/progenitor cells compared to the control. There were few cell-cell contacts between cultured LSCs and ASCs in 3D CC-ASC. In conclusion, ASCs support the growth of LSCs in the form of cell clusters but not in single cells. 3D CC-ASC could serve as a substitute for the standard 3T3 culture to expand LSCs.
Citation: Mei H, González S, Nakatsu MN, Baclagon ER, Chen FV, Deng SX (2017) Human adipose-derived stem cells support the growth of limbal stem/progenitor cells. PLoS ONE 12(10): e0186238. https://doi.org/10.1371/journal.pone.0186238
Editor: Irina Kerkis, Instituto Butantan, BRAZIL
Received: March 21, 2017; Accepted: September 27, 2017; Published: October 11, 2017
This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This work was supported by National Eye Institute grants (5P30EY000331 and 1R01EY021797), by a California Institute for Regenerative Medicine Early Translational Award (TR2-01768), and an unrestricted grant from Research to Prevent Blindness.
Competing interests: A patent application (“Novel methods to regenerate human limbal stem cells”, UC-2012-743-2-LA; US PCT/US13/44375) has been filed. This does not alter our adherence to PLOS ONE policies on sharing data and materials.

Introduction

The integrity of human corneal epithelial cells is maintained by limbal stem cells (LSCs) [1–5], which are located at the basal limbal epithelium and surrounded by niche cells including limbal mesenchymal cells [6–8], melanocytes [9], and N-cadherin-expressing cells [10]. Loss of LSCs or their dysfunction may lead to limbal stem cell deficiency (LSCD) which present with corneal opacity, vascularization and conjunctivalization. Transplantation of ex vivo expanded LSCs to the LSCD eye has been reported as a successful therapy to treat LSCD [5, 11, 12]. A comprehensive review showed that the overall success rate is 76% from 583 patients [13].

The standard method to culture LSCs on 3T3 feeder cells that have been used in clinical study is cultivating single LSC directly on top of the growth-arrested 3T3 feeder cells [14]. Once sufficient amount of LSCs is achieved, the cultivated LSCs are transplanted onto the patient’s cornea after removing the abnormal epithelium and pannus. Although 3T3 fibroblast cells are growth-arrested and theoretically are not populated in patients, there are concerns about the mouse origin of the 3T3 feeder cells in clinical applications including contamination from xenogenic molecules, immuno-rejection, and potential interspecies viral transmission. It has been reported that human embryonic stem cells co-cultured with animal-derived serum and feeder cells express immunogenic nonhuman sialic acid [15]. Retinal pigment epithelial cells and iris pigment epithelial cells co-cultured on mitomycin C-treated 3T3 fibroblasts were found to express mouse collagen type I [16]. 3T3 cells have an endogenous retrovirus containing a 3600-bp region of xenotropic murine leukemia virus-related virus (XMRV) which are associated with human prostate cancer and chronic fatigue syndrome [17].

To replace the mouse fibroblast feeder cells, human amniotic membrane and human-derived feeder cells have been examined for their potential to support the growth of LSCs in vitro. Both intact and denuded amniotic membrane have been shown to support the growth of LSCs either in the form of tissue explants or cell suspension [13, 18–24], although donor variation exists [25, 26]. Human amniotic epithelial cells might support the growth of LSCs that contained uniformly p63-positive epithelial progenitor cells [27]. Human limbal mesenchymal cells and limbal melanocytes [28, 29], and bone marrow-derived mesenchymal stem cells (BM-MSCs) [9], have been reported to serve as feeder cells to culture LSCs in vitro.

Human adipose-derived stem cells (ASCs) are an easily accessible autologous stem cell source, have a higher frequency of mesenchymal stem cells than BM-MSCs [30], and have been shown to support the growth of many types of stem cells including human embryonic stem cells [31, 32], induced pluripotent stem cells [31], and LSCs [33]. ASCs could support the in vitro expansion of LSCs with a lower clonogenic capacity than 3T3 and the expanded LSCs express some putative limbal stem/progenitor cell markers [33]. However, the comparison between the ASC and 3T3 is limited to the colony-forming efficiency (CFE) and there is limited comparison on the stem cell phenotypes of cultured LSCs, which is crucial for pre-clinical development. In addition, only direct co-culture method was used and ASCs do not show superior capacity in supporting the growth of LSCs than 3T3 [33]. We previously reported that a 3 dimensional (3D) culture system, in which the LSCs and the 3T3 feeder cells were cultured on the opposite sides of a porous membrane, supported the growth of LSCs and significantly increased the cell proliferation of LSC cultured in the form of cell clusters [34]. Whether the 3D culture system can facilitate the ASC-supported culture was examined in this study. Fibrin gel, which has been used as a carrier for epithelial cell propagation in vitro and human transplantation [14, 35], was coated on the porous membrane. The cultured LSCs on fibrin could be directly transplanted into patients’ eyes without extra retrieving steps from culture surface. In this study, the potency that ASCs support the growth of LSCs was compared to the standard culture on 3T3 cells, including cell doubling, expressions of putative stem cell markers including ATP-binding cassette sub-family G member 2 (ABCG2) [36], N-terminally truncated transcripts of p63 (∆Np63) [14, 37], N-cadherin [10] and cytokeratin (K) 14 [38], maturation marker K12 [39], and proliferation marker Ki67 [40]. Different forms of seeded LSCs and different culture methods were examined using ASC feeder cells to investigate which approach was the most optimal. The culture method using 3T3s that has been successfully used in clinical study, which is single LSCs cultured directly on 3T3 feeder cells, served as the control in all experiments.

Materials and methods

Human sclerocorneal tissue

Human sclerocorneal tissue was from the Lions Eye Institute for Transplant and Research (Tampa, FL) and the Illinois Eye Bank (Watson Gailey, Bloomington, IL). Tissue donors were aged from 20 to 65 years old. Experimentation on human tissue adhered to the tenets of the Declaration of Helsinki. The experimental protocol was evaluated and exempted by the University of California, Los Angeles Institutional Review Boards. The donors from whom the tissues were used in this study provided informed consent to being included of the study.

The tissues were preserved in Optisol (Chiron Ophthalmics, Inc., Irvine, CA), and the death-to-preservation time was less than 8 hours.

Isolation, culture, and characterization of the primary ASCs

ASCs at passage 1 and 2 were a generous gift from Prof. Bruno Peault (Professor of Orthopedic Surgery, University of California, Los Angeles). The protocol of ASC isolation was described previously [41]. In brief, the lipoaspirate were incubated in RPMI 1640 (Cellgro, Corning, NY) containing 3.5% bovine serum albumin (Sigma-Aldrich, St. Louis, MO) and 1 mg/ml collagenase type II (Sigma-Aldrich) for 30 min at 37°C. After centrifugation, the adipocytes were discarded and the pellet was further incubated in red blood cell lysis buffer (eBioscience, San Diego, CA) to remove erythrocytes. The remaining cells were cultured in MEM-α (Gibco, Grand Island, NY) supplemented with 10% FBS (Invitrogen, Carlsbad, CA) and penicillin/streptomycin (Invitrogen), which are primary ASCs at passage 0. ASCs were subcultured at 1:6 ratio at around 80% confluence. ASCs at passage 4–6 were used for experiments.

Primary ASCs were characterized by the positive expression of MSC markers CD90 [42–44] and CD105 [42–44], negative expression of endothelial and hematopoietic stem/progenitor cell markers CD31 [45, 46] and CD34 [42–44] and negative expression of differentiation markers adiponectin [44, 47] (a marker of adipogenesis) and osteocalcin [44, 48] (a marker of osteogenesis).

Isolation of limbal epithelial cells

LECs were isolated from corneoscleral rims as previously described [49]. In brief, the iris, endothelium, residual blood vessels, Tenon’s capsules, and conjunctiva were removed mechanically, followed by the digestion in 2.4 U/ml Dispase II (Roche, Indianapolis, IN) in SHEM5 growth medium (DMEM/F12 medium) (Gibco) supplemented with N-2 (Gibco), 2 ng/ml epidermal growth factor (EGF; Gibco), 8.4 ng/ml cholera toxin (Sigma-Aldrich), 0.5 μg/ml hydrocortisone (Sigma-Aldrich), 0.5% dimethyl sulfoxide (DMSO; Sigma-Aldrich), 5% fetal bovine serum (FBS, Invitrogen), penicillin/streptomycin (Invitrogen) and gentamicin/amphotericin B (Invitrogen) for 2 hours at 37°C. Epithelial cell sheets were mechanically scraped from the limbus and pipetted for several times to break the cell sheets into smaller cell clusters, which were usually a mixture of mainly single cells and some small cell clusters (around 2 to 20 cells/cluster, as shown in S1 Fig). Some cell clusters were further digested with 0.25% trypsin and 1 mM EDTA (Gibco) for 10 min at 37°C to obtain single-cell suspension. LECs, either in the form of cell clusters (composed of both single cells and small cell clusters) or single-cell suspension, were seeded at a density of 300 cells/cm2.

Cell culture of LSCs

The direct and 3D culture methods were following the protocol as previously described [34]. In brief, in the direct culture method, the LECs were seeded and cultured directly on the feeder cells. In the 3D culture method, the LECs were cultured on the top side of a 1μm pore polyethylene terephthalate (PET) membrane and the feeder cells were cultured on the bottom side of the membrane [34]. In the fibrin 3D culture method, a 1 to 2mm-thick layer of fibrin gel (Baxter, Deerfield, IL) was coated on top of the porous membrane; the feeder cells were seeded at the bottom side of the membrane and the LECs were seeded on top of the fibrin gel. Subconfluent murine 3T3 cells (from Howard Green, Harvard Medical School, Boston, MA, USA) were growth-arrested with 4 μg/ml of mitomycin C (Sigma-Aldrich) for 2 h, and plated at 3 x 104 cells/cm2 as feeder cells. Subconfluent ASCs were treated with 4 μg/ml of mitomycin C (Sigma-Aldrich) for 2 h, and plated at 5 x 103 cells/cm2 (an optimized density to support epithelial growth) as feeder cells. The LECs from the same donor were used for culture in different conditions for each experiment to minimize donor variation. The cells were cultured in SHEM5 growth medium for 14 days before harvest. The growth medium was refreshed every 2–3 days. The cell doubling was calculated as log2(number of epithelial cells harvested at day 14/ number of cells seeded).

Collection of expanded LSCs at the end of 14-day culture

For 3T3 SC control, SC-ASC, and CC-ASC culture, in which LSCs and feeder cells were grown on the same side of plates/plate inserts, the LSCs and the feeder cells were incubated in Versene (Gibco) for 3 min; then the feeder cells were washed away by pipetting and the epithelial colonies remained attached to the plates/plate inserts. For 3D SC-ASC, 3D CC-ASC, and fibrin 3D CC-ASC culture, in which LSCs and feeder cells were grown on the opposite sides of plate inserts, the feeder cells were removed by mechanical scraping with a cell scraper (Fisher Scientific) and the epithelial colonies remained attached on the top side of plate inserts. The remaining epithelial cells were incubated in 0.25% trypsin and 1 mM EDTA (Gibco) for 5 to 8 min at 37°C and collected for further analysis.

Success rate of culture

The success rate of culture, which was to examine whether the culture supported a consistent growth of LSCs from different donors, was presented as the number of donors showing LSC growth out of the total number of donors examined.

RNA isolation, reverse transcription and quantitative real-time PCR

RNA was extracted from harvested LECs (RNeasy Mini Kit, Qiagen, Valencia, CA), treated with DNase (DNA-free kit, Ambion, Austin, TX), and reverse-transcribed into cDNA (SuperScript II, Invitrogen) according to the manufacturers’ protocols. mRNA transcripts were quantified by using the Kapa Sybr Fast qPCR kit (Kapa Biosystems, Woburn, MA). Cycle conditions were as follows: the reactant was denatured for 20 s at 95°C; amplified for 40 cycles (temperatures in each cycle were 95°C for 3 s, 60°C for 20 s, and 72°C for 8 s); and subjected to a melting curve program to obtain the dissociation curves. The primers used in quantitative real-time PCR (qRT-PCR) were listed in S1 Table.

Immunocytochemistry and quantitation

Cells were cytospined onto slides by a cytocentrifuge (Cytofuge; Fisher Scientific, Hampton, NH) and stored at -80°C until use. Cytospin slides were fixed with 4% paraformaldehyde at room temperature for 10 min and washed 3 times with phosphate-buffered saline (PBS). Samples were blocked and permeabilized in PBS containing 1% BSA and 0.5% Triton X-100 (Sigma-Aldrich) for 30 min at room temperature and incubated with the primary antibody diluted in PBS containing 1% bovine serum albumin (BSA) and 0.1% Triton X-100 overnight at 4°C in a moist chamber. Slides were washed 3 times with PBS, incubated with the secondary antibody diluted in PBS containing 1% BSA and 0.1% Triton X-100 at room temperature for 1 h, and washed with PBS for three times. Nuclei were labeled with Hoechst 33342 (4 μg/ml; Invitrogen) at room temperature for 15 min, washed 5 times with PBS, and mounted in Fluoromount medium (Sigma). The primary antibodies and their dilution ratios are listed in S2 Table.

Images were taken by a confocal microscope (Confocal Laser Scanning Microscopy; Olympus, San Jose, CA) and an image capture system (Fluoview FV10-ASW 3.1 Viewer; Olympus). The nuclear intensity of p63α was analyzed by the Definiens Tissue Studio software (Larchmont, NY).

Analysis of cell-cell contact in the 3D CC-ASC culture by high resolution light microscopy and electron microscopy

At the end of the 3D CC-ASC culture, membranes with cells were carefully removed, fixed with 2% glutaraldehyde (Electron microscope Sciences, Hatfield, PA) and 2% paraformaldehyde (Electron microscope Sciences) in 0.1 M cacodylate buffer, washed with 0.1 M cacodylate buffer, osmicated for 1 h, washed extensively and embedded in Epon resin (Momentive Specialty Chemicals, Houston, TX). Tissue was sectioned at 1 μm and stained with toluidine blue (Sigma) for light microscopy. The number of the pores showing no visible cell extension, showing cell extension from epithelial cells (non-contacting extension), showing cell extension from ASC feeder cells (non-contacting extension), or showing extension throughout pores connecting epithelial cells to ASC feeder cells (contacting extension) were manually counted. For electron microscopy, tissue was sectioned at 60 nm and stained in 8% uranyl acetate (Ted Pella Inc., Redding, CA) for 15 min followed by a solution of 0.4% lead citrate (Ted Pella Inc.) and 0.4% sodium hydroxide (Fisher Scientific) for 2 min. The images were taken by a JEOL JEM1200-EX transmission electron microscope (JEOL, Peabody, MA).

Statistical analysis

Student’s t-test was performed to analyze the data. Error bar represent the standard error of the mean (SEM) from 3–5 experiments. P values ≤ 0.05 were considered statistical significant.

Results

Characterization of the ASCs

Human primary ASCs expressed the mesenchymal stem cell (MSC) markers including Cluster of Differentiation (CD) 90 and CD105 (Fig 1). The cells showed negative expression of endothelial and hematopoietic stem/progenitor cell markers, CD31 and CD34, respectively (Fig 1). There was no detectable expression of adiponectin, a differentiation marker of adipogenesis or osteocalcin, a differentiation marker of osteogenesis (Fig 1).

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Fig 1. Characterization of primary ASCs.
The expression of CD90, CD105, CD31, CD34, adiponectin, and osteocalcin was examined by immunocytochemistry in the primary ASCs at passage 4–6. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g001

Optimization of the cell density of ASCs as feeder cells

LSCs in single cell suspension were cultured directly on gradient densities of ASCs, i.e. 2.5 x 103, 5 x 103, 1 x 104, and 2 x 104 cells/cm2, for 2 weeks. The densities of 5 x 103 and 1 x 104ASCs/cm2 supported epithelial growth (Fig 2B), although the cultured LSCs were not as cuboidal and compact as the LSCs cultured on 3T3 cells (Fig 2A). The lower density of 2.5 x 103 ASCs/cm2 and the higher density of 2 x 104 ASCs/cm2 failed to support epithelial expansion and favored the growth of fibroblast-like cells instead (Fig 2B). Between the two densities, 5 x 103 and 1 x 104 ASCs/cm2, the former density seemed to support slightly better epithelial growth and was chosen for the following experiments.

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Fig 2. Optimization of the cell density of ASCs as feeder cells.
(A) Representative morphology of LSCs cultured on 3 x 104 3T3 cells/cm2 for 2 weeks. (B) Representative morphology of LSCs cultured on gradient densities of ASCs for 2 weeks. The density of feeder cells was labeled at the top right corner of each image. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g002

ASCs do not support the growth of LSCs in single cell suspension

Single cell suspension of LSCs were cultured on ASCs using the direct (SC-ASC) and 3D methods (3D SC-ASC) for 2 weeks. Single LECs cultured directly on 3T3 feeder cells (SC-3T3) served as the control. The LECs in the control group had a consistent growth (100%, 3 out of 3 donors) and a compact and cuboidal undifferentiated epithelial morphology (Fig 3B). SC-ASC and 3D SC-ASC cultures did not grow consistently that 67% of the culture (2 out of 3 donors) failed to grow and only 33% culture (1 out of 3 donors) showed epithelial growth. The expanded epithelial cells from the 33% culture formed small colonies (Fig 3A) and showed an uneven, flattened, and differentiated morphology (Fig 3B). Cell doubling in the SC-ASC and 3D SC-ASC cultures were significantly lower than the control (Fig 3C). There was no difference in cell doubling between the SC-ASC and 3D SC-ASC cultures. Due to the inconsistent growth and differentiated morphology, single LSCs cultured on ASCs were abandoned from further investigation.

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Fig 3. Single-cell suspension of limbal epithelial cells cultured on ASCs.
(A) Cell morphology of cultured LSCs. The images for SC-ASC and 3D SC-ASC were from the 33% culture which supported epithelial expansion. (B) Cell morphology of cultured LSCs. The images for SC-ASC and 3D SC-ASC were from the 33% culture which supported epithelial expansion. (C) Cell doubling of limbal epithelial cells. *: p<0.05 in comparison with results of control. Ctl: control. SC-ASC: single cell suspension of LECs cultured directly on ASCs. 3D SC-ASC: single cell suspension of LECs cultured on ASCs using the 3D method. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g003

ASCs support the growth of LECs in cell clusters

We have previously shown that LSCs grew better in the form of cell clusters than single cells in 3T3-supported culture [34]. Cell clusters of LSCs were then cultured on ASCs using the direct (CC-ASC), 3D (3D CC-ASC) and fibrin 3D (fibrin 3D CC-ASC) methods. CC-ASC and 3D CC-ASC supported a consistent (100%, 5 out of 5 donors) cell growth and produced a compact and cuboidal epithelial morphology, which was comparable to that in the control (SC-3T3 culture) (Fig 4A). However, the cells cultured in fibrin 3D CC-ASC did not always proliferate well. 33% culture (1 out of 3 donors) of fibrin 3D CC-ASC grew well and 67% culture (2 out of 3 donors) barely had proliferation. 3D CC-ASC had the consistently highest cell doubling, in which cell were doubled 9.0 times, compared to cells doubled 4.9 times in control (p<0.05), 4.9 times in CC-ASC (p<0.05), and 3.9 times in fibrin 3D CC-ASC (p<0.05) (Fig 4B). There was no difference on cell doubling among CC-ASC, fibrin 3D CC-ASC, and fibrin 3D CC-ASC (Fig 4B).

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Fig 4. Cell clusters of limbal epithelial cells cultured on ASCs.
(A) Cell morphology of cultured LSCs. The image for fibrin 3D CC-ASC was from the 33% culture which supported epithelial expansion. (B) Cell doubling of limbal epithelial cells. *: p<0.05 in comparison with results of control, CC-ASC, and fibrin 3D CC-ASC cultures. Ctl: control. CC-ASC: cell clusters of LECs cultured directly on ASCs. 3D CC-ASC: cell clusters of LECs cultured on ASCs using the 3D method. Fibrin 3D CC-ASC: cell clusters of LECs cultured on ASCs using the fibrin 3D method. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g004

The stem cell phenotype of cultured LECs was characterized by qRT-PCR. Compared to the control (SC-3T3 culture), LECs cultured on ASCs expressed a similar mRNA level of ABCG2, ΔNp63 and N-cadherin (putative LSC markers) in all three culture methods, a significantly lower level of K14 (a putative LSC marker) in 3D and fibrin 3D methods (decreased by 54% and 72%, respectively, p<0.05) and a significantly lower level of K12 (a differentiation marker) in the direct, 3D and fibrin 3D methods (decreased by 65%, 85% and 90%, respectively, p<0.05) (Fig 5). There was no difference on the mRNA level of Ki67 (a proliferation marker) between the control and the three culture methods using ASCs as feeder cells (all p>0.05, Fig 5).

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Fig 5. Relative mRNA levels of the putative stem cell markers and maturation marker in cultured LEC cell clusters with ASCs as evaluated by qRT-PCR.
The expression of each marker was normalized to the expression of housekeeping gene GAPDH and the value of the control group was designated 1. *: p<0.05 in comparison with results of the control group. Ctl: control. CC-ASC: cell clusters of LECs cultured directly on ASCs. 3D CC-ASC: cell clusters of LECs cultured on ASCs using the 3D method. Fibrin 3D CC-ASC: cell clusters of LECs cultured on ASCs using the fibrin 3D method.
https://doi.org/10.1371/journal.pone.0186238.g005

The phenotype of cultured LSCs was further examined by the protein expression using immunocytochemistry. LSC cell clusters cultured on ASCs had comparable percentages of p63α-bright and K14+ progenitor cells in all three culture methods compared to the control (Figs 6B, 7B and 7E). Percentage of p63α-bright cells was shown to correlate with clinical success rate [14]. Therefore, we used percentage of p63α-bright cells to evaluate the quality of LSCs expanded. Because 3D CC-ASC had the highest cell doubling as shown in Fig 3B, it generated significantly higher absolute numbers of p63α-bright (4.4-folds higher, p<0.05) and K14+ cells (13-folds higher, p<0.05) than the control (Figs 6C and 7C). There was no significant difference in the absolute number of K12+ cells between the control and all ASCs cultures (Fig 7F).

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Fig 6. Expression of p63α in cultured LEC cell clusters with ASCs evaluated by immunocytochemistry.
(A) Representative images showing the expression of p63α in cultured LECs. (B) The percentages of p63α-bright cells in cultured LECs. (C) The absolute numbers of p63α-bright cells in cultured LECs. The absolute number of p63α-bright cells = the percentage of p63α-bright cells x (number of cells harvested/number of cells seeded). *: p<0.05 in comparison with results of control. Ctl: control. CC-ASC: cell clusters of LECs cultured directly on ASCs. 3D CC-ASC: cell clusters of LECs cultured on ASCs using the 3D method. Fibrin 3D CC-ASC: cell clusters of LECs cultured on ASCs using the fibrin 3D method. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g006

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Fig 7. Expression of K14 and K12 in cultured LEC cell clusters with ASCs evaluated by immunocytochemistry.
(A) Representative images showing the expression of K14 in cultured LECs. (B) The percentages of K14+ cells in cultured LECs. (C) The absolute numbers of K14+ cells in cultured LECs. (D) Representative images showing the expression of K12 in cultured LECs. (E) The percentages of K12+ cells in cultured LECs. (F) The absolute numbers of K12+ cells in cultured LECs. The absolute number of K14+ or K12+ cells = the percentage of K14+ or K12+ cells x (number of cells harvested/number of cells seeded). *: p<0.05 in comparison with results of control. Ctl: control. CC-ASC: cell clusters of LECs cultured directly on ASCs. 3D CC-ASC: cell clusters of LECs cultured on ASCs using the 3D method. Fibrin 3D CC-ASC: cell clusters of LECs cultured on ASCs using the fibrin 3D method. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.g007

Few direct cell-cell contacts between the LSCs and the ASC feeder cells in 3D CC-ASC culture

To study whether there was any direct cell-cell contact between the LSCs and the ASC feeder cells, 3D CC-ASCs were fixed, sectioned across the pores in the PET membrane, stained and counted. Over 400 pores were examined and 0.5% of the pores showed direct cell-cell contact (Fig 8A, 8B and 8C). Majority (81.5%) of the pores showed no visible cell extension from either the epithelial or the ASC feeder cells. There were 18% pores containing cell extension either from the epithelial cells (11.7%) or from the ASC feeder cells (6.3%) (Fig 8C); however, the extensions failed to reach the cells on the other side of the membrane (Fig 8A).

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Fig 8. Few cell-cell contacts between the epithelial cells and the ASC feeder cells in 3D CC-ASC culture.
(A) Representative images showing an empty pore, a pore with possible cell-cell contact, a pore with no cell-cell contact but with cell extension from the epithelial cells, and a pore with no cell-cell contact but with cell extension from the ASC feeder cells, respectively. (B) An image showing the cell-cell contact between LSCs and ASC feeder cells using electron microscopy. White arrow indicates the interface of cell-cell contact. (C) Percentages of pores with empty content, cell-cell contact, no contact but with cell extension from epithelial cells, and no contact but with cell extension from ASC feeder cells in the 3D CC-ASC culture. Epi: cultured epithelial cells. Scale bar = 3 μm.
https://doi.org/10.1371/journal.pone.0186238.g008

Discussion

ASCs can be easily harvested in large quantity from white adipose tissue through minimally invasive liposuction procedure [50, 51] and are well characterized [52–54]. ASCs could serve as feeder cells for many cell types including neuronal cells [55], melanocytes [56], and various types of epithelial stem cells including LSCs [33]. Our results showed that ASCs did not support the growth of LSCs in the form of single cell suspension as evident by the differentiated morphology and poor and inconsistent cell expansion. However, ASCs supported the growth of LSCs in the form of cell clusters. To access the quality of our culture, cell morphology, cell doubling, and expression of putative stem cell markers and maturation marker were examined. CFE was not quantitatively analyzed in this study because holoclones formed from limbal stem cells could not be distinguished from meroclones by colony size or shape [57]. Instead, the expression of p63α, which has been well characterized to predict holoclones [37] and shown to correlate with clinical success [14], was examined. LSC cell clusters cultured with ASCs were compact and small in size, and contained comparable percentages of p63α-bright, K14+, and K12+ cells in all three culture methods compared to the 3T3 culture (control) although there was a reduced expression of K14 and K12 in mRNA levels. The discrepancies between the data from immunocytochemistry and those from qRT-PCR on K12 and K14 expression indicate a poor correlation between the mRNA and protein expressions. The poor correlation is observed genome-widely in both bacteria and eukaryotes. Approximately 60% of the variation in protein concentration cannot correlate to the mRNA abundance [58]. The weak correlation between mRNA and protein levels reflects the precise control on gene expression at multiple levels, including transcript stability, post-translational modification, intracellular trafficking and packaging, and/or pathway-specific degradation. The transcriptional profile, translational profile, and even the discrepancies between them, may distinguish a type of cells from other populations and serve as cell signatures.

Compared to the human limbal mesenchymal cell-supported culture and human bone marrow stromal cell-supported culture which generate similar or 3-fold more LSCs respectively than the 3T3 culture at their optimized conditions [59, 60], 3D CC-ASC supported a remarkably high cell doubling of LSCs, in which cells were doubled 9 times compared to the cells doubled 4.9 times in control. In other words, 3D CC-ASC generated 14.3-fold more LSCs than 3T3 control. Meanwhile, 3D CC-ASC maintained a comparable percentage of LSC/progenitor cell population compared to 3T3 control, which led to significantly higher absolute numbers of p63α-bright and K14+ cells in cultured LSCs. The fact that 3D CC-ASC could support a significantly higher cell doubling of LSCs while maintaining the percentage of LSCs/progenitor cells is of great clinical significance, which enables the generation of sufficient amount of expanded LSCs for ransplantation from fewer cells derived from small biopsies. There were few cell-cell contacts between the LSCs and the ASCs in 3D CC-ASC culture thus limiting the contamination from feeder cells. These results suggest that expanding LSC cell clusters on ASCs using the 3D culture method may be an appropriate substitute for the 3T3 culture method.

The mRNA levels of Ki67, a proliferation marker, were comparable between the SC-3T3 control culture and the three culture methods in CC-ASC cultures in Fig 5, which was not consistent with the actual cell doubling in Fig 4B. One possible explanation is that the LSCs in the 3D CC-ASC culture may reach confluence at the end of 2-week culture and may stop the highly proliferative state at the time point of cell harvesting for mRNA quantitation.

The reason why 3T3 cells support the expansion of LSCs in single cell suspension whereas ASCs do not needs to be elucidated. One possible explanation is ASCs secret different molecules. It has been reported that ASCs showed a different mRNA expression pattern of secretary molecules that are known to regulate epithelial stem cells including pleiotrophin, cystatin C, hepatocyte growth factor, keratinocyte growth factor, Insulin-like growth factor 1α compared to 3T3 cells [33].

Another explanation may be the separation of LSCs from their neighboring cells and/or the cleavage of membrane proteins of LSCs by trypsin/EDTA that are important for the survival or attachment of LSCs. Previous reports showed that when single LECs which contained the LSC population and the LSC niche cells were seeded at a high density (5×104 cells/cm2), the reunion of LSCs with their niche cells occurred and promoted the propagation of LSCs in vitrowithout 3T3 feeder cells [8]. However, when single LSCs were seeded at a lower density (300 cells/cm2) and single LSCs were separated sparsely from the niche cells, there was no cell growth without 3T3 feeder cells [61]. Similar to the niche cells, 3T3 cells support the single LSC growth under direct feeder-LSC contact, however they do not support the growth when the LSCs are cultured separately in the overhanging cell culture inserts [62]. These findings support the notion that LSCs need to be in direct contact or in close proximity with the niche cells or 3T3 feeder cells to survive and proliferate in vitro. In other words, niche cells and 3T3 feeder cells provide niche factors that signal through cell-cell contact or within a short distance to support the survival and proliferation of LSCs in vitro. This hypothesis that LSCs need to be in direct contact or in close proximity with the niche cells or feeder cells to survive and proliferate can also be used to explain the inferior performance of fibrin 3D CC-ASC culture. The 1–2 mm thick fibrin destructs the close proximity between ASC feeder cells and LSCs and may also block the cell-cell contact between the ASCs and LSCs, thus less niche factors from feeder cells could travel this distance through the gel and reach the LSCs to support their survival and proliferation. Interestingly, single LSCs were able to grow into transplantable cell sheets on denuded amniotic membrane without feeder cells [63], which suggests that amniotic membrane may contain the essential niche signals for the in vitro survival and proliferation of LSCs. In addition, trypsin, as a serine protease, cleaves outer membrane peptide chains mainly at the carboxyl side of lysine and arginine. The process of trypsinization to make single LSCs suspension may damage some membrane or membrane-associated molecules on LSCs which in turn impair the stem cell survival and adhesion.

In summary, human ASCs have the capacity to support the expansion of LSCs in vitro. A fine-tuned ASC-supported culture system including seeding LEC clusters and using the 3D culture method can achieve a significantly high cell expansion meanwhile maintaining the percentage of putative limbal stem/progenitor cell population with minimal cell-cell contacts from feeder cells, which enables the generation of sufficient amount of LSCs for transplantation from fewer cells derived from small biopsies while minimizing cell contamination from feeder cells. Therefore, 3D CC-ASC appears to be a good substitute for the standard 3T3 culture to expand LSCs in vitro for clinical application.

Supporting information

S1 Fig.tif

Representative images showing the cell clusters of LECs.
The cell clusters of LECs were obtained by Dispase digestion of corneoscleral rim followed by mechanical scraping and pippeting. The cell clusters of LECs were composed of mainly single cells and some small cell clusters (usually around 2 to 20 cells/cluster). The cell clusters in dashed rectangles are enlarged at the sides of the images. Scale bar = 100 μm.
(TIF)

S1 Fig. Representative images showing the cell clusters of LECs.

The cell clusters of LECs were obtained by Dispase digestion of corneoscleral rim followed by mechanical scraping and pippeting. The cell clusters of LECs were composed of mainly single cells and some small cell clusters (usually around 2 to 20 cells/cluster). The cell clusters in dashed rectangles are enlarged at the sides of the images. Scale bar = 100 μm.
https://doi.org/10.1371/journal.pone.0186238.s001

(TIF)

S1 Table. Primers used in qRT-PCR.

https://doi.org/10.1371/journal.pone.0186238.s002

(DOCX)

S2 Table. Primary antibodies used in immunocytochemistry.

https://doi.org/10.1371/journal.pone.0186238.s003

(DOCX)

Acknowledgments

Human adipose-derived stem cells were a generous gift from Prof. Bruno Peault (Professor of Orthopedic Surgery, University of California, Los Angeles).

References

  1. 1.Davanger M, Evensen A. Role of the pericorneal papillary structure in renewal of corneal epithelium. Nature. 1971;229(5286):560–1. pmid:4925352.
  2. 2.Cotsarelis G, Cheng SZ, Dong G, Sun TT, Lavker RM. Existence of slow-cycling limbal epithelial basal cells that can be preferentially stimulated to proliferate: implications on epithelial stem cells. Cell. 1989;57(2):201–9. pmid:2702690.
  3. 3.Tsai RJ, Sun TT, Tseng SC. Comparison of limbal and conjunctival autograft transplantation in corneal surface reconstruction in rabbits. Ophthalmology. 1990;97(4):446–55. pmid:1691476.
  4. 4.Huang AJ, Tseng SC. Corneal epithelial wound healing in the absence of limbal epithelium. Invest Ophthalmol Vis Sci. 1991;32(1):96–105. pmid:1702774.
  5. 5.Ebato B, Friend J, Thoft RA. Comparison of central and peripheral human corneal epithelium in tissue culture. Invest Ophthalmol Vis Sci. 1987;28(9):1450–6. pmid:3623831.
  6. 6.Chen SY, Hayashida Y, Chen MY, Xie HT, Tseng SC. A new isolation method of human limbal progenitor cells by maintaining close association with their niche cells. Tissue Eng Part C Methods. 2011;17(5):537–48. pmid:21175372; PubMed Central PMCID: PMC3129703.
  7. 7.Funderburgh JL, Funderburgh ML, Du Y. Stem Cells in the Limbal Stroma. The ocular surface. 2016;14(2):113–20. pmid:26804252; PubMed Central PMCID: PMC4842326.
  8. 8.Xie HT, Chen SY, Li GG, Tseng SC. Isolation and expansion of human limbal stromal niche cells. Invest Ophthalmol Vis Sci. 2012;53(1):279–86. pmid:22167096; PubMed Central PMCID: PMC3292364.
  9. 9.Li W, Hayashida Y, Chen YT, Tseng SC. Niche regulation of corneal epithelial stem cells at the limbus. Cell Res. 2007;17(1):26–36. pmid:17211449; PubMed Central PMCID: PMC3190132.
  10. 10.Hayashi R, Yamato M, Sugiyama H, Sumide T, Yang J, Okano T, et al. N-Cadherin is expressed by putative stem/progenitor cells and melanocytes in the human limbal epithelial stem cell niche. Stem Cells. 2007;25(2):289–96. pmid:17008425.
  11. 11.Tsai RJ, Li L, Chen J. Reconstruction of damaged corneas by transplantation of autologous limbal epithelial cells(1). Am J Ophthalmol. 2000;130(4):543. pmid:11024443.
  12. 12.Rama P, Bonini S, Lambiase A, Golisano O, Paterna P, De Luca M, et al. Autologous fibrin-cultured limbal stem cells permanently restore the corneal surface of patients with total limbal stem cell deficiency. Transplantation. 2001;72(9):1478–85. pmid:11707733.
  13. 13.Baylis O, Figueiredo F, Henein C, Lako M, Ahmad S. 13 years of cultured limbal epithelial cell therapy: a review of the outcomes. J Cell Biochem. 2011;112(4):993–1002. pmid:21308743.
  14. 14.Rama P, Matuska S, Paganoni G, Spinelli A, De Luca M, Pellegrini G. Limbal stem-cell therapy and long-term corneal regeneration. N Engl J Med. 2010;363(2):147–55. pmid:20573916.
  15. 15.Martin MJ, Muotri A, Gage F, Varki A. Human embryonic stem cells express an immunogenic nonhuman sialic acid. Nat Med. 2005;11(2):228–32. pmid:15685172.
  16. 16.Johnen S, Wickert L, Meier M, Salz AK, Walter P, Thumann G. Presence of xenogenic mouse RNA in RPE and IPE cells cultured on mitotically inhibited 3T3 fibroblasts. Invest Ophthalmol Vis Sci. 2011;52(5):2817–24. pmid:21220557.
  17. 17.Mendoza R, Vaughan AE, Miller AD. The left half of the XMRV retrovirus is present in an endogenous retrovirus of NIH/3T3 Swiss mouse cells. J Virol. 2011;85(17):9247–8. pmid:21697491; PubMed Central PMCID: PMC3165791.
  18. 18.Nakamura T, Inatomi T, Sotozono C, Ang LP, Koizumi N, Yokoi N, et al. Transplantation of autologous serum-derived cultivated corneal epithelial equivalents for the treatment of severe ocular surface disease. Ophthalmology. 2006;113(10):1765–72. pmid:16905193.
  19. 19.Kolli S, Ahmad S, Lako M, Figueiredo F. Successful clinical implementation of corneal epithelial stem cell therapy for treatment of unilateral limbal stem cell deficiency. Stem Cells. 2010;28(3):597–610. pmid:20014040.
  20. 20.Meller D, Fuchsluger T, Pauklin M, Steuhl KP. Ocular surface reconstruction in graft-versus-host disease with HLA-identical living-related allogeneic cultivated limbal epithelium after hematopoietic stem cell transplantation from the same donor. Cornea. 2009;28(2):233–6. pmid:19158575.
  21. 21.Kawashima M, Kawakita T, Satake Y, Higa K, Shimazaki J. Phenotypic study after cultivated limbal epithelial transplantation for limbal stem cell deficiency. Arch Ophthalmol. 2007;125(10):1337–44. pmid:17923540.
  22. 22.Grueterich M, Espana EM, Touhami A, Ti SE, Tseng SC. Phenotypic study of a case with successful transplantation of ex vivo expanded human limbal epithelium for unilateral total limbal stem cell deficiency. Ophthalmology. 2002;109(8):1547–52. pmid:12153809.
  23. 23.Baradaran-Rafii A, Ebrahimi M, Kanavi MR, Taghi-Abadi E, Aghdami N, Eslani M, et al. Midterm outcomes of autologous cultivated limbal stem cell transplantation with or without penetrating keratoplasty. Cornea. 2010;29(5):502–9. pmid:20299977.
  24. 24.Ang LP, Sotozono C, Koizumi N, Suzuki T, Inatomi T, Kinoshita S. A comparison between cultivated and conventional limbal stem cell transplantation for Stevens-Johnson syndrome. Am J Ophthalmol. 2007;143(1):178–80. pmid:17188066.
  25. 25.Liu J, Sheha H, Fu Y, Liang L, Tseng SC. Update on amniotic membrane transplantation. Expert Rev Ophthalmol. 2010;5(5):645–61. pmid:21436959; PubMed Central PMCID: PMC3061461.
  26. 26.Hopkinson A, McIntosh RS, Tighe PJ, James DK, Dua HS. Amniotic membrane for ocular surface reconstruction: donor variations and the effect of handling on TGF-beta content. Invest Ophthalmol Vis Sci. 2006;47(10):4316–22. pmid:17003421.
  27. 27.Chen YT, Li W, Hayashida Y, He H, Chen SY, Tseng DY, et al. Human amniotic epithelial cells as novel feeder layers for promoting ex vivo expansion of limbal epithelial progenitor cells. Stem Cells. 2007;25(8):1995–2005. pmid:17495107; PubMed Central PMCID: PMC3197019.
  28. 28.Zhang X, Sun H, Li X, Yuan X, Zhang L, Zhao S. Utilization of human limbal mesenchymal cells as feeder layers for human limbal stem cells cultured on amniotic membrane. J Tissue Eng Regen Med. 2010;4(1):38–44. pmid:19813216.
  29. 29.Dziasko MA, Tuft SJ, Daniels JT. Limbal melanocytes support limbal epithelial stem cells in 2D and 3D microenvironments. Exp Eye Res. 2015;138:70–9. pmid:26142953.
  30. 30.Murphy MB, Moncivais K, Caplan AI. Mesenchymal stem cells: environmentally responsive therapeutics for regenerative medicine. Exp Mol Med. 2013;45:e54. pmid:24232253; PubMed Central PMCID: PMC3849579.
  31. 31.Hwang ST, Kang SW, Lee SJ, Lee TH, Suh W, Shim SH, et al. The expansion of human ES and iPS cells on porous membranes and proliferating human adipose-derived feeder cells. Biomaterials. 2010;31(31):8012–21. pmid:20674000.
  32. 32.Kim JS, Kwon D, Hwang ST, Lee DR, Shim SH, Kim HC, et al. hESC expansion and stemness are independent of connexin forty-three-mediated intercellular communication between hESCs and hASC feeder cells. PLoS One. 2013;8(7):e69175. pmid:23922689; PubMed Central PMCID: PMC3724839.
  33. 33.Sugiyama H, Maeda K, Yamato M, Hayashi R, Soma T, Hayashida Y, et al. Human adipose tissue-derived mesenchymal stem cells as a novel feeder layer for epithelial cells. J Tissue Eng Regen Med. 2008;2(7):445–9. pmid:18792424.
  34. 34.Mei H, Gonzalez S, Nakatsu MN, Baclagon ER, Lopes VS, Williams DS, et al. A three-dimensional culture method to expand limbal stem/progenitor cells. Tissue Eng Part C Methods. 2014;20(5):393–400. pmid:24047104; PubMed Central PMCID: PMC4005490.
  35. 35.Feng Y, Borrelli M, Reichl S, Schrader S, Geerling G. Review of alternative carrier materials for ocular surface reconstruction. Curr Eye Res. 2014;39(6):541–52. pmid:24405104.
  36. 36.de Paiva CS, Chen Z, Corrales RM, Pflugfelder SC, Li DQ. ABCG2 transporter identifies a population of clonogenic human limbal epithelial cells. Stem Cells. 2005;23(1):63–73. pmid:15625123; PubMed Central PMCID: PMC2906389.
  37. 37.Di Iorio E, Barbaro V, Ruzza A, Ponzin D, Pellegrini G, De Luca M. Isoforms of DeltaNp63 and the migration of ocular limbal cells in human corneal regeneration. Proc Natl Acad Sci U S A. 2005;102(27):9523–8. pmid:15983386; PubMed Central PMCID: PMCPMC1172259.
  38. 38.Schermer A, Galvin S, Sun TT. Differentiation-related expression of a major 64K corneal keratin in vivo and in culture suggests limbal location of corneal epithelial stem cells. J Cell Biol. 1986;103(1):49–62. pmid:2424919; PubMed Central PMCID: PMC2113783.
  39. 39.Kasper M, Moll R, Stosiek P, Karsten U. Patterns of cytokeratin and vimentin expression in the human eye. Histochemistry. 1988;89(4):369–77. pmid:2457569.
  40. 40.Joyce NC, Meklir B, Joyce SJ, Zieske JD. Cell cycle protein expression and proliferative status in human corneal cells. Invest Ophthalmol Vis Sci. 1996;37(4):645–55. pmid:8595965.
  41. 41.Corselli M, Chin CJ, Parekh C, Sahaghian A, Wang W, Ge S, et al. Perivascular support of human hematopoietic stem/progenitor cells. Blood. 2013;121(15):2891–901. pmid:23412095; PubMed Central PMCID: PMC3707421.
  42. 42.Sousa BR, Parreira RC, Fonseca EA, Amaya MJ, Tonelli FM, Lacerda SM, et al. Human adult stem cells from diverse origins: an overview from multiparametric immunophenotyping to clinical applications. Cytometry A. 2014;85(1):43–77. pmid:24700575.
  43. 43.Maleki M, Ghanbarvand F, Reza Behvarz M, Ejtemaei M, Ghadirkhomi E. Comparison of mesenchymal stem cell markers in multiple human adult stem cells. Int J Stem Cells. 2014;7(2):118–26. pmid:25473449; PubMed Central PMCID: PMC4249894.
  44. 44.Dominici M, Le Blanc K, Mueller I, Slaper-Cortenbach I, Marini F, Krause D, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 2006;8(4):315–7. pmid:16923606.
  45. 45.Zimmerlin L, Donnenberg VS, Rubin JP, Donnenberg AD. Mesenchymal markers on human adipose stem/progenitor cells. Cytometry A. 2013;83(1):134–40. pmid:23184564; PubMed Central PMCID: PMC4157311.
  46. 46.Zimmerlin L, Donnenberg VS, Pfeifer ME, Meyer EM, Peault B, Rubin JP, et al. Stromal vascular progenitors in adult human adipose tissue. Cytometry A. 2010;77(1):22–30. pmid:19852056; PubMed Central PMCID: PMC4148047.
  47. 47.Satish L, Krill-Burger JM, Gallo PH, Etages SD, Liu F, Philips BJ, et al. Expression analysis of human adipose-derived stem cells during in vitro differentiation to an adipocyte lineage. BMC Med Genomics. 2015;8:41. pmid:26205789; PubMed Central PMCID: PMC4513754.
  48. 48.Hauschka PV. Osteocalcin: the vitamin K-dependent Ca2+-binding protein of bone matrix. Haemostasis. 1986;16(3–4):258–72. pmid:3530901.
  49. 49.Nakatsu MN, Ding Z, Ng MY, Truong TT, Yu F, Deng SX. Wnt/beta-catenin signaling regulates proliferation of human cornea epithelial stem/progenitor cells. Invest Ophthalmol Vis Sci. 2011;52(7):4734–41. pmid:21357396; PubMed Central PMCID: PMC3175950.
  50. 50.Shiffman MA, Mirrafati S. Fat transfer techniques: the effect of harvest and transfer methods on adipocyte viability and review of the literature. Dermatol Surg. 2001;27(9):819–26. pmid:11553171.
  51. 51.Locke M, Windsor J, Dunbar PR. Human adipose-derived stem cells: isolation, characterization and applications in surgery. ANZ J Surg. 2009;79(4):235–44. pmid:19432707.
  52. 52.Bourin P, Bunnell BA, Casteilla L, Dominici M, Katz AJ, March KL, et al. Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT). Cytotherapy. 2013;15(6):641–8. pmid:23570660; PubMed Central PMCID: PMC3979435.
  53. 53.Wosnitza M, Hemmrich K, Groger A, Graber S, Pallua N. Plasticity of human adipose stem cells to perform adipogenic and endothelial differentiation. Differentiation. 2007;75(1):12–23. pmid:17244018.
  54. 54.Aksu AE, Rubin JP, Dudas JR, Marra KG. Role of gender and anatomical region on induction of osteogenic differentiation of human adipose-derived stem cells. Ann Plast Surg. 2008;60(3):306–22. pmid:18443514.
  55. 55.Kang SK, Jun ES, Bae YC, Jung JS. Interactions between human adipose stromal cells and mouse neural stem cells in vitro. Brain Res Dev Brain Res. 2003;145(1):141–9. pmid:14519500.
  56. 56.Kim JY, Park CD, Lee JH, Lee CH, Do BR, Lee AY. Co-culture of melanocytes with adipose-derived stem cells as a potential substitute for co-culture with keratinocytes. Acta Derm Venereol. 2012;92(1):16–23. pmid:21879248.
  57. 57.Pellegrini G, Rama P, Matuska S, Lambiase A, Bonini S, Pocobelli A, et al. Biological parameters determining the clinical outcome of autologous cultures of limbal stem cells. Regen Med. 2013;8(5):553–67. pmid:23725042.
  58. 58.Vogel C, Marcotte EM. Insights into the regulation of protein abundance from proteomic and transcriptomic analyses. Nat Rev Genet. 2012;13(4):227–32. pmid:22411467; PubMed Central PMCID: PMC3654667.
  59. 59.Gonzalez S, Mei H, Nakatsu MN, Baclagon ER, Deng SX. A 3D culture system enhances the ability of human bone marrow stromal cells to support the growth of limbal stem/progenitor cells. Stem Cell Res. 2016;16(2):358–64. pmid:26896856; PubMed Central PMCID: PMC4828302.
  60. 60.Nakatsu MN, Gonzalez S, Mei H, Deng SX. Human limbal mesenchymal cells support the growth of human corneal epithelial stem/progenitor cells. Invest Ophthalmol Vis Sci. 2014;55(10):6953–9. pmid:25277234; PubMed Central PMCID: PMC4215742.
  61. 61.Gonzalez S, Deng SX. Presence of native limbal stromal cells increases the expansion efficiency of limbal stem/progenitor cells in culture. Exp Eye Res. 2013;116:169–76. pmid:24016868; PubMed Central PMCID: PMC3900305.
  62. 62.Miyashita H, Shimmura S, Higa K, Yoshida S, Kawakita T, Shimazaki J, et al. A novel NIH/3T3 duplex feeder system to engineer corneal epithelial sheets with enhanced cytokeratin 15-positive progenitor populations. Tissue Eng Part A. 2008;14(7):1275–82. pmid:18433313.
  63. 63.Kawakita T, Shimmura S, Hornia A, Higa K, Tseng SC. Stratified epithelial sheets engineered from a single adult murine corneal/limbal progenitor cell. J Cell Mol Med. 2008;12(4):1303–16. pmid:18318692; PubMed Central PMCID: PMCPMC3225011.






2. https://stemcells.nih.gov/info/basics/7.htm

Can Stem Cells Mend a Broken Heart?: Stem Cells for the Future Treatment of Heart Disease

Cardiovascular disease (CVD), which includes hypertension, coronary heart disease, stroke, and congestive heart failure, has ranked as the number one cause of death in the United States every year since 1900 except 1918, when the nation struggled with an influenza epidemic. Nearly 2,600 Americans die of CVD each day, roughly one person every 34 seconds. Given the aging of the population and the relatively dramatic recent increases in the prevalence of cardiovascular risk factors such as obesity and type 2 diabetes, CVD will be a significant health concern well into the 21st century.
Cardiovascular disease can deprive heart tissue of oxygen, thereby killing cardiac muscle cells (cardiomyocytes). This loss triggers a cascade of detrimental events, including formation of scar tissue, an overload of blood flow and pressure capacity, the overstretching of viable cardiac cells attempting to sustain cardiac output, leading to heart failure, and eventual death. Restoring damaged heart muscle tissue, through repair or regeneration, is therefore a potentially new strategy to treat heart failure.
The use of embryonic and adult-derived stem cells for cardiac repair is an active area of research. A number of stem cell types, including embryonic stem (ES) cells, cardiac stem cells that naturally reside within the heart, myoblasts (muscle stem cells), adult bone marrow-derived cells including mesenchymal cells (bone marrow-derived cells that give rise to tissues such as muscle, bone, tendons, ligaments, and adipose tissue), endothelial progenitor cells (cells that give rise to the endothelium, the interior lining of blood vessels), and umbilical cord blood cells, have been investigated as possible sources for regenerating damaged heart tissue. All have been explored in mouse or rat models, and some have been tested in larger animal models, such as pigs.

A few small studies have also been carried out in humans, usually in patients who are undergoing open-heart surgery. Several of these have demonstrated that stem cells that are injected into the circulation or directly into the injured heart tissue appear to improve cardiac function and/or induce the formation of new capillaries. The mechanism for this repair remains controversial, and the stem cells likely regenerate heart tissue through several pathways. However, the stem cell populations that have been tested in these experiments vary widely, as do the conditions of their purification and application. Although much more research is needed to assess the safety and improve the efficacy of this approach, these preliminary clinical experiments show how stem cells may one day be used to repair damaged heart tissue, thereby reducing the burden of cardiovascular disease.

 2011;702:289-98. doi: 10.1007/978-1-61737-960-4_21.

Epithelial differentiation of human adipose-derived stem cells.

Baer PC1, Brzoska M, Geiger H.

Abstract

The versatile differentiation potential of adipose-derived stem cells (ASC) into cells of mesodermal, entodermal, and ectodermal origin places these cells at the forefront of cell-based therapies and cell transplantation. Epithelial differentiation of ASC may either be initiated by direct cell-cell or cell-matrix contacts, by chemical factors like retinoic acid, or via secreted cellular factors like cytokines, interleukins, or growth factors included in conditioned media.This protocol describes methods to induce the in vitro differentiation of ASC from human adipose tissue into the epithelial lineage, and describes the methods used to verify this induced differentiation. We present two differentiation protocols based on either retinoic acid or conditioned medium of cultured epithelial cells.



. 2014; 2014: 395781.
Published online 2014 Jan 20. doi:  10.1155/2014/395781
PMCID: PMC3915932
PMID: 24563864

Direct Cell-Cell Contact between Mesenchymal Stem Cells and Endothelial Progenitor Cells Induces a Pericyte-Like Phenotype In Vitro

Markus Loibl, 1 , 2 Andreas Binder, 1 , 2 Marietta Herrmann, 1 Fabian Duttenhoefer, 1 , 3 R. Geoff Richards, 1Michael Nerlich, 2 Mauro Alini, 1 and Sophie Verrier 1 ,*
Academic Editor: Sanga Gehmert
Author information ▼ Article notes ► Copyright and License information ▼ Disclaimer
Copyright © 2014 Markus Loibl et al.
This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

1. Introduction

The remarkable regeneration potential of bone tissue is based on the presence of a highly branched vessel network, providing oxygen, nutrients, growth factors, and precursor cells to the injured tissue []. However, the capacity of regeneration is limited and fails in large bone defects. Thus, the gold-standard strategy remains autologous bone implants, associated with an additional surgical procedure at the harvesting site causing an increase of operation time, pain, and risk for infection at the donor site.
Cell-based tissue engineering strategies have been developed and are accepted since a few years to address the challenge of bone repair. The approach of engineering bone tissue not only depends on the presence of osteogenic cells at the healing site but also requires an adequate vascularisation of the applied biomaterial []. Therefore, bone-forming osteoblasts and endothelial cells (or their progenitors) are playing a crucial role for successful engraftment of cell seeded biomaterials [].
Bone marrow is a natural reservoir of mesenchymal stem cells (MSCs) able to develop an osteogenic, chondrogenic, and adipogenic phenotype upon stimulation. The recruitment, growth, and differentiation of MSCs into mature osteoblasts are regulated by many cytokines and growth factors. These factors are secreted not only by the osteoblasts themselves, but also from endothelial cells from the tightly connected vascular network [, , ]. This crosstalk between different cell types is enabled by secreted paracrine factors, as well as by direct cell-to-cell interactions.
In 1997, Asahara et al. identified the presence of bone marrow derived endothelial progenitor cells (EPCs) able to develop an endothelial phenotype in peripheral blood of adults. Those progenitor cells are known to express CD34 [], CD133 [], and CD309 [] and as mature endothelial cells, they specifically bind UEA-1 lectins [, ]. As they are present in the blood compartment as well as in the bone marrow, they can easily be isolated in an autologous way, which is a fundamental advantage for further tissue engineering applications.
Recently, we developed an in vitro prevascularised 3D polyurethane (PU) bone implant seeded with EPCs (CD34+/CD133+) and MSCs, which showed the formation of luminal tubular structures after 7 days of coculture []. These capillary-like structures expressed not only mature endothelial cell markers (i.e., PECAM-1, vWF), but also pericyte (PC) markers (i.e., CD146, NG2, and αSMA). PCs are still poorly characterised [] and their reported markers are also expressed on other cells associated with blood vessels [, ]. Coexpression of several markers (CD146 [, ], NG2 [], αSMA [], and PDGFR-β[]) and absence of endothelial markers (PECAM-1) [] may be used to identify PCs in vitro. However, coexpression of these markers by PCs is variable and depends not only on the tissue of origin [], but also on culture conditions []. Hellström et al. [] and more recently Blocki et al. suggested that PCs represent a subpopulation of MSCs in bone marrow, contributing to microvessel maturation, stability, structure, and function [, ]. PCs establish important direct cell-cell contact with endothelial cells of immature blood vessels [] and some studies suggested that PCs may serve as guiding structures aiding outgrowth of endothelial cells to form early capillary sprouts [].
Based on our previous findings concerning the participation of PC-like cells in the overall structure of in vitro preformed capillary network [], the aim of the present study was to determine the origin of PC-like cells in cocultures of MSCs and EPCs in vitro. We analysed the influence of cell-cell interactions in two different 2D coculture systems. In addition, two cell culture media were tested; on one hand, cells were cultured in a classical cell culture medium without any growth factor supplements; on the other hand, the medium was supplemented with autologous platelet lysate growth factors (PL) to provide the optimal condition for EPCs [].

2. Materials and Methods

2.1. Preparation of Platelet Lysate Growth Factors (PL)

PL was prepared from platelet concentrates, as described earlier []. Platelet bags were obtained from the blood bank of Kantonsspital Graubünden in Chur in accordance with the current ethical laws of Switzerland. The platelet bags contained a standardised platelet density (5 times above physiological concentration), obtained by blood apheresis. The platelet density was further increased by a centrifugation at 2000 g for 20 minutes. After two washing steps in phosphate buffer saline (PBS) and subsequent centrifugation, the platelet pellets were resuspended in half of the original volume of PBS to obtain a final density 10 times higher than that in normal blood (2.5 million (±10%) platelets/μL). PL samples were pooled from platelet concentrates from three different donors and randomly matched.

2.2. Bone Marrow Aspirates

Human bone marrow aspirates (20 mL) were obtained upon informed consent and ethical approval (EK Regensburg 12-101-0127) from 6 donors undergoing routine orthopaedic surgery (20 to 76 years old, average age 43 years; 4 females and 2 males). The samples were processed within 24 hours after harvesting.

2.3. Mesenchymal Stem Cells (MSC)

Bone marrow mononucleated cells (BMCs) were isolated from bone marrow aspirates by Histopaque-1077 (Sigma-Aldrich) density gradient centrifugation, as previously described []. Briefly, bone marrow was homogenised and diluted 1 : 4 with PBS. After slowly pipetting on Histopaque, the samples were centrifuged at 800 g for 20 minutes. The interphase containing BMCs was collected and washed twice in Iscove Modified Dulbecco Medium (IMDM, Gibco) containing 10% human MSC qualified fetal calf serum (FCS, Gibco), followed by 15 min centrifugation at 400 g. BMCs were seeded at the density of 16 × 106 mononucleated cells per 300 cm2 cell culture flask (BD Biosciences, Falcon Cell Culture Dishes) and cultured in Minimum Essential Medium Alpha (MEMα, Gibco) containing PenStrep (PS) (100 U/mL, Gibco), 10% FCS, and 5 ng/mL basic fibroblast growth factor (bFGF) (R&D Systems). After the first passage, adhesion selected MSCs were further subcultured at a density of 0.9 × 106 per 300 cm2 cell culture flask.

2.4. Endothelial Progenitor Cells (EPC) and CD34-CD133-CD146-Depleted MSCs (Depleted-MSCs)

CD34+, CD133+, and CD146+ cell populations were further selected from the previously isolated BMCs using magnetic microbeads linked to antibodies specific to CD133, CD34, or CD146 (MACS system, Miltenyi Biotec) according to the manufacturer instructions. After isolation, the different cell populations were referred to as CD133+, CD34+, or CD146+, respectively. The remaining cell population (depleted from CD133+, CD34+, and CD146+ cells) was termed depleted-MSCs. CD34+ and CD133+ cells were pooled and from then on referred to as EPCs. Cells from the depleted-MSCs population were propagated in MEMα containing PS (100 U/mL), 10% FCS, and 5 ng/mL bFGF. EPCs were cultured in IMDM containing PS (100 U/mL), supplemented with 5% PL, 5% FCS, and 1% nonessential amino acids (NEAA, Gibco). All cell types were cultured at 37°C 5% CO2 humidified atmosphere incubator and media were changed twice a week.

2.5. Cell Culture

For all subsequent experiments, cells between passages 2 and 3 were used. MSCs, depleted-MSCs, and EPCs (CD34+/CD133+) were enzymatically detached (Trypsin-EDTA), counted, seeded at a density of 5,000 cells/cm2 in 3 different culture setups (Figure 1), and incubated for 3, 7, or 10 days in presence of IMDM-FCS (IMDM supplemented with 10% FCS and 1% NEAA) or IMDM-PL (IMDM supplemented with 5% FCS, 5% PL, and 1% NEAA).

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Cell culture setups. MSCs or depleted-MSCs were seeded with EPCs in transwell culture (a) or direct coculture (b). MSCs, depleted-MSCs, and EPCs were seeded in single cultures (c) as controls. All experiments were performed in the presence of IMDM-FCS (IMDM supplemented with 10% FCS) or IMDM-PL (IMDM supplemented with 5% FCS, 5% PL).
For indirect cocultures (transwell), MSCs (or depleted-MSCs) were seeded in the bottom part of 6-well plates at a density of 5,000 cells/cm2 for transwell culture setup (Figure 1(a)), whereas EPCs were seeded at the same density in the corresponding transwell cell culture inserts (0.4 μm pore size, Sigma-Aldrich).
For direct coculture experiments (direct coculture), MSCs and EPCs, as well as depleted-MSCs and EPCs, were coseeded in a 1 : 1 ratio in 75 cm2 cell culture plate (BD Biosciences) at an initial density of 5,000 cells/cm2 (Figure 1(b)). In addition, each cell type was seeded individually in 6-well plates (5,000 cells/cm2) for single culture as controls (Figure 1(c)).

2.6. Cell Sorting

Prior to gene expression analysis, cells in direct cocultures were separated by fluorescence activated cell sorting (FACS) using FITC labelled lectin (UEA-1, Sigma-Aldrich) which specifically labels EPCs. Adherent cells were incubated for 60 minutes at 37°C in the dark with 10 μg/mL FITC-lectin in IMDM-FCS as previously described []. Afterwards, cells were washed three times with PBS and trypsinised, followed by centrifugation at 2000 g for 10 min. The resulting cell pellet was resuspended in 500 μL PBS 1% FCS, filtered through a 40 μm cell strainer, and subjected to FACS analysis (FACSAria III, BD Biosciences). FITC signal was identified with a 488 nm excitation laser and 502LP filter. Cell analysis was performed on at least 10,000 events for each sample and analysed using FACS DIVA software version 6.1.3 (BD Biosciences). A primary gate based on physical parameters (forward and side scatter) was set to exclude dead cells or small debris. For the separation, gating was implemented based on the negative-control staining profile of MSCs in direct coculture with EPCs. Sorting gates were determined by fluorescence intensity of FITC maintaining a gap between the lectin negative and the lectin positive gate. Cells were fractionated with an efficiency of >90% in lectin positive and lectin negative cell populations and collected in PBS 1% FCS. The lectin positive cell population represents EPCs, and the lectin negative population represents MSCs or depleted-MSCs. At least 50,000 events per sorted cell type were collected. After separation, cells were spin down and mRNA was extracted for further analysis. In parallel, the sorted cell populations were reseeded (1 : 1 ratio) onto a 2-well glass slide (LabTek chamber slides, Thermo Fisher scientific) at 5,000 cells per cm2 for immunofluorescence staining.

2.7. Real-Time PCR

At days 3, 7, and 10, total RNA was extracted from monolayers or sorted cells using Tri Reagent (Molecular Research Centre, Inc.) according to the manufacturer’s protocol and stored at −80°C until further use. cDNA was synthesised from 1 μg RNA using TaqMan reverse transcription reagents (Applied Biosystems, Invitrogen) with MultiScribe reverse transcriptase (50 U/μL) and random hexamer primers. Real-time Polymerase chain reaction (PCR) was performed on the StepOne Plus machine (Applied Biosystems). Genes of interest were detected using CD146/MCAM (HS00174838_m1), NG2/CSPG4 (HS00426981_m1), αSMA/ACTA2 (HS00909449_m1), PECAM-I (HS01065282_m1), and PDGFR-β(HS00182163_m1) all purchased from Applied Biosystems. Human GAPDH (Cat no. 4326317E, Applied Biosystems) was used as a housekeeping gene. PCR conditions were 95°C for 10 min, followed by 45 cycles of amplification at 95°C for 15 sec and 60°C for 1 min using the Stepone software v2.1 (Applied Biosystems). Relative quantification of mRNA targets was performed according to the comparative ΔΔCt method.

2.8. Immunocytochemistry

After 7 days of cell culture in IMDM-FCS or IMDM-PL, cells were separated and seeded onto LabTek chamber slides. Cells were allowed to adhere for at least 6 hours prior to fixation with 70% methanol for 10 minutes and 100% methanol for 2 minutes and stored at −20°C until further use. Cells were rehydrated in PBS prior to staining procedure. Nonspecific bindings were blocked by incubation with 5% normal goat serum (Vector Labs) in PBS for 60 minutes. Thereafter, slides were incubated with a mouse anti-human antibody (anti-hCD146 (clone P1H12, ab24577) or anti-hNG2 (clone LHM2, ab20156)) and a rabbit anti-human antibody (anti-hNG2 (ab104535) or anti-hαSMA (ab5694)) simultaneously. Mouse anti-human and rabbit anti-human antibodies were used in 3 different combinations: CD146/NG2, CD146/αSMA, and NG2/αSMA. After 1 h of incubation at room temperature, the samples were washed and stained with corresponding secondary fluorescent antibodies: Alexa Fluor 488 conjugated anti-mouse antibody (Life Technologies, A11029) and TRITC conjugated anti-rabbit antibody (abcam, ab50598).
Primary and secondary antibodies were both incubated for 1 hour at room temperature. Stained slides were cover-slipped with Prolong Gold Antifade reagent with DAPI (Molecular Probes, Life Technologies). A digital image was obtained by using an AxioCam HRc camera and AxioVision software V3.1 (Carl Zeiss).

2.9. Statistical Analysis

Results are presented as mean ± standard error of the mean. Statistical analysis was performed using Prism 4 software (Graphpad). One-way ANOVA with Tukey’s multiple comparison test was used to calculate the overall differences in nonsize matched experimental groups. Level of significance was P < 0.05.

3. Results

3.1. Cell Characterisation

The baseline expression of the pericyte and endothelial marker genes in each of the 3 tested cell populations (MSCs, depleted-MSCs, and EPCs) was measured prior to exposure to the different culture conditions (day 0) using GAPDH as a housekeeping gene. Similar expression profile was observed on day 0 for the MSCs and depleted-MSCs populations when propagated in MEMα supplemented with 10% FCS and 5 ng/mL bFGF (Figures 2(a) and 2(b)). Moreover, αSMA and PDGFR-β were expressed at a higher level compared to CD146 and NG2 in MSCs and depleted-MSCs, without a significant difference (P > 0.05). However, as expected PECAM-1 was expressed at a lower level than the PC marker genes.


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Baseline gene expression at day 0 of MSCs, depleted-MSCs, and EPCs (ΔCt values relative to GAPDH in logarithmic scale). Pericyte marker genes (CD146, NG2, αSMA, and PDGFR-β) and an endothelial marker gene (PECAM-1) were detected in MSCs (a), depleted-MSCs (b), and EPCs (c) after single culture in culture medium at day 0 in 5 independent experiments.
A similar trend of baseline gene expression was observed for EPCs population after culturing in IMDM supplemented with 5% FCS and 5% PL, without a significant difference for all tested genes (P > 0.05). The endothelial marker PECAM-1 was expressed at a lower level than the PC marker genes similar to MSCs and depleted-MSCs.

3.2. Influence of Cell Culture Setups on Gene Expression

Gene expression analysis of MSCs cultured in IMDM-FCS revealed an upregulation of CD146 (Figure 3(a)) for all time points and all cell media when compared to day 0. Representative data are shown for day 3 and are summarised in Supplementary Table  1 (see Supplementary Material available online at http://dx.doi.org/10.1155/2014/395781). CD146 expression demonstrated a 15.1 ± 6.99-fold increase compared to day 0 in coculture and a 3.25 ± 0.89- and 1.97 ± 0.58-fold increase in single and transwell culture, respectively (Figure 3(a)). A similar trend was detected for NG2 but to a lesser extent. NG2 expression demonstrated a 6.4 ± 2.6-fold increase compared to day 0 in coculture and a 3.21 ± 0.36- and 2.06 ± 0.21-fold increases in single and transwell culture, respectively (Figure 3(b)). An overall decrease of αSMA and PDGFR-β expression was observed for MSCs, with a lower extent for the direct coculture condition (Figures 3(c) and 3(d)). The expression of PECAM-1 was decreased in MSC in all culture conditions and at all time points (Figure 3(e)). A similar trend of gene expression was shown for depleted-MSCs (Supplementary Figure  1).


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Gene regulation in MSCs and EPCs after 3 days of culture in IMDM-FCS. Pericyte marker genes CD146 ((a), (f)), NG2 ((b), (g)), αSMA ((c), (h)), and PDGFR-β ((d), (i)) and endothelial marker gene PECAM-1 ((e), (j)) were detected in MSCs and EPCs after 3 days in transwell culture, direct coculture, and single culture in IMDM-FCS (IMDM supplemented with 10% FCS). Results are presented for each gene as relative change in gene expression (2−ΔΔCt) over time between day 0 and day 3 in 5 independent experiments (donor 1–5) and as average ± error of the mean of all experiments.
Analysis of EPCs revealed an increased CD146 expression in all culture setups. In contrast to MSC, direct coculture induced the lowest level of upregulation (2−ΔΔCt2.42 ± 0.78, Figure 3(f)). A similar trend was apparent for NG2 but to a lesser extent (Figure 3(g)).
The αSMA expression showed a constant decrease in single culture or transwell culture (2−ΔΔCt0.63 ± 0.1 and 2−ΔΔCt0.58 ± 0.07, resp.). However, EPCs in direct coculture with MSC demonstrated no change in αSMA expression (2−ΔΔCt0.97 ± 0.2) (Figure 3(h)). The PDGFR-β expression showed a marginal increase in direct coculture with MSCs (2−ΔΔCt1.46 ± 0.27) (except donor 1), a minor increase in transwell culture (2−ΔΔCt1.35 ± 0.65) (except donor 4), and no changes in gene expression in single culture (Figure 3(i)).
Of interest, a significant upregulation of PECAM-1 was observed in EPCs (2−ΔΔCt8.16 ± 3.49) when cocultured in direct cell-cell contact with MSCs in comparison to single or transwell culture (2−ΔΔCt0.26 ± 0.08 and 2−ΔΔCt0.21 ± 0.09) (Figure 3(j)).
EPC gene expression in all culture setups revealed similar trends when EPCs were cocultured with depleted-MSCs (Supplementary Figure  1).

3.3. Influence of Culture Medium on Gene Expression

Gene expression analysis of MSCs, depleted-MSCs, and EPCs cultured in IMDM-PL (Figure 4 and Supplementary Figure  2) revealed the same trend of changes as for IMDM-FCS at all time points. Representative data are presented for day 3 (Figure 4 and Supplementary Figure  2). However, the differences between the cell culture conditions were less prominent as detected for IMDM-FCS medium. Similar to IMDM-FCS, highest upregulation of CD146 and NG2 expression was detected in MSCs in direct coculture with EPCs (Figures 4(a) and 4(b)). CD146 expression demonstrated a 8.44 ± 3.5-fold increase in direct coculture and a 4.3 ± 1.62- and 3.47 ± 0.98-fold increases in single and transwell culture, respectively (Figure 4(a)). In addition, NG2 expression revealed a 3.11 ± 0.64-fold increase for cells in direct coculture, a 2.32 ± 0.55-fold increase for single culture, and a 1.51 ± 0.34-fold increase for transwell culture (Figure 4(b)).


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Gene regulation in MSCs and EPCs after 3 days of culture in IMDM-PL. Pericyte marker genes CD146 ((a), (f)), NG2 ((b), (g)), αSMA ((c), (h)), and PDGFR-β ((d), (i)) and endothelial marker gene PECAM-1 ((e), (j)) were detected in MSCs and EPCs after 3 days in transwell culture, direct coculture, and single culture in IMDM-PL (IMDM supplemented with 5% FCS and 5% PL). Results are presented for each gene as relative change in gene expression (2−ΔΔCt) over time between day 0 and day 3 in 5 independent experiments (donor 1–5) and as average ± error of the mean of all experiments.
An overall decrease in gene expression of αSMA and PDGFR-β was observed in MSCs after 3 days when cultured in IMDM FCS (Figures 4(c) and 4(d)). The 2−ΔΔCt values (day 3/day 0) of PECAM-1 decreased in MSCs for all conditions (Figure 4(e)).
EPCs in IMDM-PL showed an upregulation of CD146 and NG2 at day 3 in all culture setups. In contrast to MSCs, EPCs showed the lowest upregulation in direct coculture with MSCs in comparison to single or transwell cultures. In detail, CD146 expression in EPCs direct coculture with MSCs propagated in IMDM-PL showed a 1.7 ± 0.51-fold increase compared to a 2.13 ± 0.82- and 3.0 ± 0.94-fold increases in single or transwell culture, respectively (Figure 4(f)). A 1.36 ± 0.48-fold increase was observed for NG2 expression in EPCs in coculture with MSCs in IMDM-PL, and a 2.54 ± 1.03- and 3.22 ± 1.48-fold upregulation in single and transwell culture were observed, respectively (Figure 4(g)).
Gene expression of αSMA and PDGFR-β in EPCs was likewise not affected by the different cell-culture setups. Both genes were downregulated at day 3 without significant differences between culture conditions (Figures 4(h) and 4(i)).
Noteworthy, a 10.2 ± 5.94-fold increase of PECAM-1 was observed in EPCs in direct coculture with MSCs in IMDM-PL. PECAM-1 expression of EPCs was of lesser extent in single and transwell culture, with a 1.12 ± 0.40- and 2.53 ± 0.89-fold increase, respectively (both > 0.05) (Figure 4(j)).

3.4. MSCs in Direct Coculture Coexpress CD146, αSMA, and NG2

Expression of the PC markers CD146, αSMA, and NG2, were detected by immunofluorescence staining on MSCs after 7 days when cocultured with EPCs and consecutive separation. Single positive and double positive cells were detectable, in both IMDM-FCS and IMDM-PL (Figure 5).

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Coexpression of pericyte markers on MSCs after direct coculture with EPCs for 7 days in IMDM-FCS (top row) or IMDM-PL (bottom row). Immunofluorescence staining for CD146, NG2, and αSMA demonstrated double positive cells for each combination (CD146 and NG2, CD146 and αSMA, and NG2 and αSMA). Negative controls were stained only with the fluorescent labelled secondary antibody. Cell nuclei are stained with DAPI (blue). Scale bars = 50 μm.

4. Discussion

The aim of the present study was to investigate the interactions between EPCs and MSCs in cocultures to determine the origin of PC-like cells in vitro. In addition, we evaluated the influence of platelet derived growth factors on EPCs and MSCs in this experimental setup.
In our previous work, we successfully established a 3D prevascularised scaffold in vitro []. We demonstrated a beneficial effect of MSCs coseeded with EPCs and observed the formation of luminal tubular structures already after 7 days in vitro. Notably, tubular structures were positive for CD146, NG2, and αSMA, suggesting the presence of PC-like cells.
In the present study, MSCs and EPCs were cultured in direct contact or in a transwell system to elucidate the influence of soluble, paracrine factors and to identify the population of origin of in vitro differentiated PCs.
It is well accepted that CD34 and CD133 are EPC markers []. In contrast to EPC, PCs do not express CD34 and CD133; however, they do express CD146 [, , ]. Therefore, we assume that the depleted-MSC cell population, characterised by the absence of CD34, CD133, or CD146, is free of EPCs and CD146+ PCs.
This was confirmed by gene expression analysis of depleted-MSCs at day 0, which was similar to the profile obtained for MSCs.
To date, there is no single surface marker that is specific or exclusively expressed on PCs []. However, PCs can be identified by coexpressing CD146 [, ], NG2 [], αSMA [], and PDGFR-β [] while being negative for PECAM-1 []. We observed a remarkable increase of CD146 and NG2 expression on MSCs and depleted-MSCs when cocultured with EPCs. PDGFR-β expression decreased but was still the highest expressed gene of all examined PC markers. Analysing the gene expression after different days of coculture, we found the most prominent differential regulation of genes on day 3. The increased CD146 and NG2 expression, the maintained high expression of PDGFR-β, and the complete reduction of PECAM-1 in MSCs and depleted-MSCs support the assumption that MSCs and depleted-MSCs differentiate towards a PC-like phenotype.
In addition, the gene expression of depleted-MSCs and MSCs revealed similar results, indicating that CD146 positive cells present in MSCs did not proliferate. This underlines the hypothesis that MSCs and depleted-MSCs can differentiate towards a PC phenotype. In line, immunocytochemistry revealed positive signals of CD146, NG2, and αSMA on MSCs which indicates the presence of triple positive cells after 7 days of direct coculture. Interestingly, the immunofluorescence analysis demonstrated a positive signal for αSMA at day 7, whereas the mRNA level was already decreased at this time point (data not shown).
We reported the highest upregulation of CD146 and NG2 for MSCs and depleted-MSCs in direct coculture with EPC. Therefore, direct cell-cell contact with EPCs might be an important factor for the differentiation of MSCs and depleted-MSCs towards a PC-like phenotype. The underlying mechanism remains unknown. However, integrins may be involved in this promotion of the angiogenic response of endothelial cells since the α6 integrin subunit in MSCs has been reported to play a role in vessel formation and PC differentiation [].
PCs were originally described to be indistinguishable in culture from MSCs []. Covas et al. demonstrated a similar gene expression profile of retinal PCs compared to MSCs from 12 different fetal and adult tissues [], highlighting the close relationship of both cell populations. Therefore, Covas et al. suggested that MSCs and PCs might be related cells, present at the vascular wall, and constitute a MSC compartment extending throughout the entire organism []. The large overall similarity between PCs and MSCs, with regard to their immunophenotype and differentiation potential in vitro, was already described by Shi and Gronthos and Armulik et al. [, ]. This supports our hypothesis that MSCs and depleted-MSCs differentiate towards a PC-like phenotype.
Armulik et al. noted that the expression of the combination of markers which were used to confirm the PC phenotype in our study is variable and depends on culture conditions []. We used two media conditions based on IMDM. MEMα was used for MSCs/depleted-MSCs expansion prior to the experiment. Armulik et al. demonstrated a strong NG2 expression of human placenta derived PCs when cultured in PC growth media (PGM, Promocell); however, staining intensity decreased after transfer to Dulbecco’s Modified Eagle Medium (DMEM). In contrast to this finding, αSMA staining was enhanced for both human placenta derived PCs and MSCs when cultured in DMEM. This emphasises a crucial, but still unknown, impact of culture conditions.
CD133+ and CD34+ separated cells were highly purified in our study, but these antigens are not expressed on mature endothelial cells [, ]. On mRNA level, the investigated EPCs lack the marker of mature endothelial cells PECAM-1 at the beginning. However, lectin positive cells showed a significant increase of PECAM-1 expression. Aguirre et al. [] reported an upregulation of the endothelial phenotype in EPCs and MSCs after 3 days of direct coculture without any supplementary growth factors. Our data support that EPCs can differentiate into mature ECs by direct cell-cell contact with MSCs. However, in the present study, cocultured EPCs were separated by lectin binding, which favours an additional purification of endothelial cells from a more heterologous EPCs population.
Previous studies showed a positive effect on proliferation and differentiation of endothelial cells treated with growth factors released by PL [, ]. Our data support these findings, since PECAM-1 expression of EPCs is higher in IMDM-PL than in IMDM-FCS. Furthermore, we observed that PL diminished the difference in gene expression for all PC genes, in both MSCs/depleted-MSCs and EPCs. PL consists of a composition of growth factors and cytokines, which might be responsible for the reported changes in the gene expression.
Interestingly, PECAM-1 expression seems to be particularly influenced by PL, since PECAM-1 expression is increased in most cell culture conditions in IMDM-PL whereas only in direct cocultures in IMDM-FCS. Therefore, PL might be a trigger for the differentiation of EPCs towards ECs. More recently, a study demonstrated the importance of PDGF and EGF signalling in controlling PC recruitment to luminal tubular structures []. Although we have not explicitly examined these signalling molecules in the present study, future work can determine if PL influences these pathways in the same way.

5. Conclusion

In summary, we report preliminary data on the effect of cell-cell contact of bone marrow derived MSCs and EPCs with regard to the induction of PC-like cells. The upregulation of several PC marker genes (CD146, NG2, and αSMA) indicates the differentiation of MSCs/depleted-MSCs towards PCs in vitro. The change of phenotype is detectable as early as day 3 of coculture.
Moreover, our data confirm that several growth factors, released by PL, contribute to the differentiation of EPCs towards endothelial cells. The effect of PL on PC induction may be further evaluated in future studies. These findings emphasise the concept of coculture strategies to promote angiogenesis for cell-based tissue engineered bone grafts.

Supplementary Material

Gene regulation in Depleted-MSCs and EPCs after 3 days of culture in IMDM-FCS. Pericyte marker genes CD146 (A, F), NG2 (B, G), αSMA (C, H), and PDGFR-β (D, I) and endothelial marker gene PECAM-1 (E, J) were detected in depleted-MSCs and EPCs after 3 days in transwell culture, direct coculture and single culture in IMDM-FCS (IMDM supplemented with 10% FCS). Results are presented for each gene as relative change in gene expression (2-ΔΔCt) over time between day 0 and day 3 for transwell culture, direct co-culture, and single culture in 5 independent experiments (donor 1-5), and as mean ± error of the mean of all experiments.
Gene regulation in Depleted-MSCs and EPCs after 3 days of culture in IMDM-PL. Pericyte marker genes CD146 (A, F), NG2 (B, G), αSMA (C, H), and PDGFR-β (D, I) and endothelial marker gene PECAM-1 (E, J) were detected in depleted-MSCs and EPCs after 3 days in transwell culture, direct coculture and single culture in IMDM-PL (IMDM supplemented with 5% FCS, 5% PL). Results are presented for each gene as relative change in gene expression (2-ΔΔCt) over time between day 0 and day 3 for transwell culture, direct coculture, and single culture in 5 independent experiments (donor 1-5) and as mean ± standard error of the mean of all experiments.
Gene regulation in MSCs, Depleted-MSCs and corresponding EPCs after 3 days of culture in IMDM-FCS and IMDM-PL. Pericyte marker genes CD146, NG2, αSMA, and PDGFR-β, and endothelial marker gene PECAM-1 were detected in MSCs, depleted-MSCs and corresponding EPCs after 3 days in transwell culture, direct coculture and single culture in IMDM-FCS (IMDM supplemented with 10% FCS) and IMDM-PL (IMDM supplemented with 5% FCS, 5% PL). Results are presented for each gene as relative change in gene expression (2-ΔΔCt) over time between day 0 and day 3 for transwell culture, coculture, and single culture as mean ± standard error of the mean of 5 independent experiments.
Click here for additional data file.(251K, pdf)

Acknowledgments

The authors would like to thank Dr. Ursula Menzel for excellent technical assistance with the flow cytometer. This work was supported by the AO Foundation. The publication cost of this work was supported by the German Research Foundation (DFG) within the funding programme Open Access Publishing. The FACS AriaIII was kindly donated by the Innovationsstiftung Graubünden.

Conflict of Interests

The authors declare that they have no conflict of interests regarding the publication of this paper.

Authors’ Contribution

Markus Loibl and Andreas Binder contributed equally to this work.

References

1. Johnson EO, Troupis T, Soucacos PN. Tissue-engineered vascularized bone grafts: basic science and clinical relevance to trauma and reconstructive microsurgery. Microsurgery. 2011;31(3):176–182.[PubMed]
2. Grellier M, Bordenave L, Amédée J. Cell-to-cell communication between osteogenic and endothelial lineages: implications for tissue engineering. Trends in Biotechnology. 2009;27(10):562–571. [PubMed]
3. Krenning G, van Luyn MJA, Harmsen MC. Endothelial progenitor cell-based neovascularization: implications for therapy. Trends in Molecular Medicine. 2009;15(4):180–189. [PubMed]
4. Aguirre A, Planell JA, Engel E. Dynamics of bone marrow-derived endothelial progenitor cell/mesenchymal stem cell interaction in co-culture and its implications in angiogenesis. Biochemical and Biophysical Research Communications. 2010;400(2):284–291. [PubMed]
5. Bianchi G, Banfi A, Mastrogiacomo M, et al. Ex vivo enrichment of mesenchymal cell progenitors by fibroblast growth factor 2. Experimental Cell Research. 2003;287(1):98–105. [PubMed]
6. Asahara T, Murohara T, Sullivan A, et al. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997;275(5302):964–967. [PubMed]
7. Quirici N, Soligo D, Caneva L, Servida F, Bossolasco P, Deliliers GL. Differentiation and expansion of endothelial cells from human bone marrow CD133+ cells. British Journal of Haematology. 2001;115(1):186–194. [PubMed]
8. Peichev M, Naiyer AJ, Pereira D, et al. Expression of VEGFR-2 and AC133 by circulating human CD34+ cells identifies a population of functional endothelial precursors. Blood. 2000;95(3):952–958.[PubMed]
9. Holthofer H, Virtanen I, Kariniemi AL. Ulex europaeus I lectin as a marker for vascular endothelium in human tissues. Laboratory Investigation. 1982;47(1):60–66. [PubMed]
10. Duttenhoefer F, Lara de Freitas R, Meury T, et al. 3D scaffolds co-seeded with human endothelial progenitor and mesenchymal stem cells: evidence of prevascularisation within 7 days. European Cells and Materials. 2013;26:49–65. [PubMed]
11. Bondjers C, He L, Takemoto M, et al. Microarray analysis of blood microvessels from PDGF-B and PDGF-Rbeta mutant mice identifies novel markers for brain pericytes. The FASEB Journal. 2006;20(10):1703–1705. [PubMed]
12. Ozerdem U, Alitalo K, Salven P, Li A. Contribution of bone marrow-derived pericyte precursor cells to corneal vasculogenesis. Investigative Ophthalmology and Visual Science. 2005;46(10):3502–3506.[PMC free article] [PubMed]
13. Covas DT, Panepucci RA, Fontes AM, et al. Multipotent mesenchymal stromal cells obtained from diverse human tissues share functional properties and gene-expression profile with CD146+ perivascular cells and fibroblasts. Experimental Hematology. 2008;36(5):642–654. [PubMed]
14. Li Q, Yu Y, Bischoff J, Mulliken JB, Olsen BR. Differential expression of CD146 in tissues and endothelial cells derived from infantile haemangioma and normal human skin. Journal of Pathology. 2003;201(2):296–302. [PubMed]
15. Huang F-J, You W-K, Bonaldo P, Seyfried TN, Pasquale EB, Stallcup WB. Pericyte deficiencies lead to aberrant tumor vascularizaton in the brain of the NG2 null mouse. Developmental Biology. 2010;344(2):1035–1046. [PMC free article] [PubMed]
16. Nehls V, Drenckhahn D. The versatility of microvascular pericytes: from mesenchyme to smooth muscle? Histochemistry. 1993;99(1):1–12. [PubMed]
17. Winkler EA, Bell RD, Zlokovic BV. Pericyte-specific expression of PDGF beta receptor in mouse models with normal and deficient PDGF beta receptor signaling. Molecular Neurodegeneration. 2010;5(1, article 32) [PMC free article] [PubMed]
18. Song S, Ewald AJ, Stallcup W, Werb Z, Bergers G. PDGFRβ+ perivascular progenitor cells in tumours regulate pericyte differentiation and vascular survival. Nature Cell Biology. 2005;7(9):870–879.[PMC free article] [PubMed]
19. Blocki A, Wang Y, Koch M, et al. Not all MSCs can act as pericytes: functional in vitro assays to distinguish pericytes from other mesenchymal stem cells in angiogenesis. Stem Cells and Development. 2013;22(17):2347–2355. [PMC free article] [PubMed]
20. Hellström M, Gerhardt H, Kalén M, et al. Lack of pericytes leads to endothelial hyperplasia and abnormal vascular morphogenesis. Journal of Cell Biology. 2001;152(3):543–553. [PMC free article][PubMed]
21. Adams RH, Alitalo K. Molecular regulation of angiogenesis and lymphangiogenesis. Nature Reviews Molecular Cell Biology. 2007;8(6):464–478. [PubMed]
22. Amselgruber WM, Schäfer M, Sinowatz F. Angiogenesis in the bovine corpus luteum: an immunocytochemical and ultrastructural study. Anatomia, Histologia, Embryologia. 1999;28(3):157–166.[PubMed]
23. Lippross S, Loibl M, Hoppe S, et al. Platelet released growth factors boost expansion of bone marrow derived CD34+ and CD133+ endothelial progenitor cells for autologous grafting. Platelets. 2011;22(6):422–432. [PubMed]
24. Meury T, Verrier S, Alini M. Human endothelial cells inhibit BMSC differentiation into mature osteoblasts in vitro by interfering with osterix expression. Journal of Cellular Biochemistry. 2006;98(4):992–1006. [PubMed]
25. Zimmerlin L, Donnenberg VS, Pfeifer ME, et al. Stromal vascular progenitors in adult human adipose tissue. Cytometry Part A. 2010;77(1):22–30. [PMC free article] [PubMed]
26. Armulik A, Genové G, Betsholtz C. Pericytes: developmental, physiological, and pathological perspectives, problems, and promises. Developmental Cell. 2011;21(2):193–215. [PubMed]
27. Carrion B, Kong YP, Kaigler D, Putnam AJ. Bone marrow-derived mesenchymal stem cells enhance angiogenesis via their alpha6beta1 integrin receptor. Experimental Cell Research. 2013;319(19):2964–2976. [PMC free article] [PubMed]
28. Péault B. Are mural cells guardians of stemness?: from pluri-to multipotency via vascular pericytes. Circulation. 2012;125(1):12–13. [PubMed]
29. Shi S, Gronthos S. Perivascular niche of postnatal mesenchymal stem cells in human bone marrow and dental pulp. Journal of Bone and Mineral Research. 2003;18(4):696–704. [PubMed]
30. Armulik A, Abramsson A, Betsholtz C. Endothelial/pericyte interactions. Circulation Research. 2005;97(6):512–523. [PubMed]
31. Gehling UM, Ergün S, Schumacher U, et al. In vitro differentiation of endothelial cells from AC133-positive progenitor cells. Blood. 2000;95(10):3106–3112. [PubMed]
32. Verrier S, Meury TR, Kupcsik L, Heini P, Stoll T, Alini M. Platelet-released supernatant induces osteoblastic differentiation of human mesenchymal stem cells: potential role of BMP-2. European Cells & Materials. 2010;20:403–414. [PubMed]
33. Stratman AN, Schwindt AE, Malotte KM, Davis GE. Endothelial-derived PDGF-BB and HB-EGF coordinately regulate pericyte recruitment during vasculogenic tube assembly and stabilization. Blood. 2010;116(22):4720–4730. [PMC free article] [PubMed]

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