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Differentiation within autologous fibrin scaffolds of porcine dermal cells with the mesenchymal stem cell phenotype
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319 (2013) 144–152
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Research Article
Differentiation within autologous fibrin scaffolds of porcine dermal cells with the mesenchymal stem cell phenotype ˜ab, Marta Lo´peza, Jennifer Ramosa, Javier Iglesiasa Pilar de la Puentea,n, Dolores Luden a
´n, Spain Tissue Bank, San Francisco Clinic Foundation, Av./Facultad 51, 51, 24004 Leo Pathology Service, University Hospital of Salamanca, P/San Vicente 58-182, 37007 Salamanca, Spain
b
article information
abstract
Article Chronology:
Porcine mesenchymal stem cells (pMSCs) are an attractive source of cells for tissue engineering
Received 13 July 2012
because their properties are similar to those of human stem cells. pMSCs can be found in
Received in revised form
different tissues but their dermal origin has not been studied in depth. Additionally, MSCs
17 October 2012
differentiation in monolayer cultures requires subcultured cells, and these cells are at risk of
Accepted 20 October 2012
dedifferentiation when implanting them into living tissue. Following this, we attempted to
Available online 1 November 2012
characterize the MSCs phenotype of porcine dermal cells and to evaluate their cellular
Keywords:
proliferation and differentiation in autologous fibrin scaffolds (AFSs). Dermal biopsies and
Porcine mesenchymal stem cells
blood samples were obtained from 12 pigs. Dermal cells were characterized by flow cytometry.
Dermal cells
Frozen autologous plasma was used to prepare AFSs. pMSC differentiation was studied in
Fibrin scaffolds
standard structures (monolayers and pellets) and in AFSs. The pMSCs expressed the CD90 and
Differentiation
CD29 markers of the mesenchymal lineage. AFSs afforded adipogenic, osteogenic and chondrogenic differentiation. The porcine dermis can be proposed to be a good source of MSCs with adequate proliferative capacity and a suitable expression of markers. The pMSCs also showed optimal proliferation and differentiation in AFSs, such that these might serve as a promising autologous and implantable material for use in tissue engineering. & 2012 Elsevier Inc. All rights reserved.
Introduction Mesenchymal stem cells (MSCs) are a heterogeneous subset of stromal stem cells that have the capacity for self-renewal, and are multipotent and display immunological features, antiinflammation properties, and trophic effects [1,2]. In view of these properties, MSCs can be considered as an ideal candidate cell type for tissue engineering and regenerative medicine. The International Society for Cellular Therapy (ISCT) has defined the minimum criteria for defining human MSCs [3]. They must be plastic-adherent when maintained in standard culture conditions; Z95% of the MSC population must express CD105, CD73, and CD90, and they must lack the expression (r2% n
positive) of CD45, CD34, CD14 or CD11b, Cd79a or CD19, and HLA class II. Furthermore, the cells must be able to differentiate into adipocytes, osteoblasts and chondrocytes under standard in vitro differentiating conditions. Most stem cells have been described in studies carried out on humans and small animals. Owing to the similarity in size and anatomical features of pigs and humans, porcine stem cells may offer a more attractive possibility for evaluating the efficacy and applicability of MSCs. Porcine MSCs (pMSCs) have also been found in bone marrow [4–6], amniotic fluid [7], umbilical cord blood [7], muscle [8], and fetal skin [9]. However, not much is known about the stem cells of the porcine dermis. Multipotent cell populations in skin have been identified from different locations, such as epidermal stem cells
Corresponding author. Postal address: Paseo Salamanca 59, 21B, 24010, Leo´n, Spain. Fax: þ34 987218801. E-mail address: [email protected] (P. de la Puente).
0014-4827/$ - see front matter & 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.yexcr.2012.10.009
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from the epidermis and hair follicle stem cells from appendages [10]. Despite this, the dermis may represent a larger reservoir for adult stem cells than the epidermis and the hair follicles together [11]. In 2001, mesenchymal stem cells in the dermis were first isolated in mice and later they were also found in the human dermis [12]. We believe that the dermis might provide an abundant alternative source of MSCs for autologous and allogenic stem cell therapy, because the dermis is a large organ and is readily accessible. Differentiation in monolayer cultures is the technique most frequently referred to in the literature, but for these cells to be suitable for implantation, prior subculture is required. Subculture involves the dissociation of the cells from each other and the substrate in order to generate a suspension of single cells, which are therefore at risk of losing their differentiation properties. These pitfalls are usually overcome by the use of tissueengineered implantable scaffolds, capable of stimulating cellular growth in different biomaterials by providing a 3D structure so that the cells can adhere, proliferate and differentiate. We have recently developed easy-to-process, readily available and totally autologous fibrin scaffolds (AFSs) [13], in which cells are able to integrate and adhere themselves, and proliferate within a biocompatible and porous structure. Although some types of fibrin scaffold have long been used in tissue engineering for many applications [14], scaffolds of autologous fibrin with platelet-poor plasma or platelet-rich plasma represent an excellent biological support for stromal cells, whereas those made from commercial fibrins show poor stromal cell survival [15]. Some studies have shown that human embryonic stem cells [16] and human stromal stem cells [15] may be successfully differentiated in fibrin scaffolds, but very little is known about their properties as regards the differentiation of porcine MSCs in such structures. Accordingly, here we were prompted to assess the ability of porcine dermal cells to differentiate into mesenchymal lineages when cultured inside AFSs. A further aim was to check the mesenchymal stem cell phenotype and multilineage potential of these cells.
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All animals received humane care in compliance with the 86/609/CEE European Economic Community Directive and the 1201/2005 Spanish Royal Decree at the Veterinary School of Leo´n. The experimental protocol was reviewed and approved by the Research Committee of the University of Leo´n. The animals were premedicated intramuscularly with 0.35 mg/kg Midazolam (Normon, Madrid, Spain) and 5 mg/kg ketamine (Imalgene, Marquez, Mexico). Anesthesia was maintained with inhaled 2% isofluorane (Isoflo, Abbot, Chicago, USA). The animals were not euthanized after the study, because they were used for other experiments not directly linked to the present study. Dermal biopsies and blood were obtained as previously described by us [13]: small pieces of dermis (1–3 g) were incubated with type I collagenase (Sigma Aldrich, St. Louis, USA) at 2 mg/ml for 22–24 h. 104 dermal cells/cm2 were seeded in six-well plates and cultured in complete culture medium (Table 1), which was refreshed every 2–3 days. Cells were subcultured every 5 days using trypsin/EDTA (T/E, PAA, Pasching, Austria).The proliferative capacity of porcine MSCs was evaluated by cell doublings, using the following formula: log (N)/log (2), where N¼cells harvested/cells seeded. The results are expressed as cumulative cell doublings. Blood samples of 25 ml from each animal were centrifuged at 1620g for 10 min, after which the plasma was frozen until it was used to prepare the fibrin scaffolds. AFSs were formed by mixing 400 ml/ml plasma (autologous from each animal), 104 cells/ml (autologous from each animal), 4 mg/ml tranexamic acid (Rottapharm, Valencia, Spain) and 4 mg/ml calcium chloride (Braun, Barcelona, Spain) diluted in complete medium to elaborate 2 ml of fibrin gel, as described previously [13]. P1 cells were counted every 3 days using the Trypan blue (Sigma Aldrich) exclusion method in order to obtain a growth curve of the cells in the autologous fibrin scaffolds (AFSs), treated enzymatically with type I collagenase (Sigma Aldrich) at 2 mg/ml for 24 h at 37 1C, and in tissue culture polystyrene (TCPS) treated enzymatically with trypsin/EDTA at 0.05/0.02% for 3 min at 37 1C.
Materials and methods Flow cytometry Isolation and culture of pMSCs Dermis and blood from Large-White crossbred pigs of 3 months of age (n¼ 12, 15–35 kg) were extracted under sterile conditions.
Cells at the 2–3 passage were trypsinized, centrifuged, and 500,000 cells were resuspended in PBS/BSA buffer (Phosphate Buffer Saline (Sigma Aldrich), pH 7.4, and 1% Bovine Serum
Table 1 – Composition of culture and differentiation media, and methods to assess lineage differentiation (DMEM, Dulbecco’s Modified Eagle Medium; FBS, Fetal Bovine Serum; IBMX, 3-isobutyl-1-methylxanthine; TGF-b1, Transforming growth factor b1). Culture medium
Complete culture medium
Supplements
Histochemical and immunohistochemical analysis
DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM
Controls with the same techniques as induction cells. H&E, and Oil Red O staining
L-Glutamine
Adipogenic differentiation medium Osteogenic differentiation medium Chondrogenic differentiation medium
DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ100 nM Dexamethasomeþ0,5 mM IBMXþ10 mg/ml Insulinþ200 mM Indomethacinþ500 mM ascorbate 2-phosphate DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ10 nM Dezamethasomeþ500 mM ascorbate 2phosphateþ10 mM b-glycerophosphate DMEMþ1% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ10 ng/ml TGF b1þ6,25 mg/ml Insulinþ50 mg/ml ascorbate 2-phosphateþ100 nM Dexamethasome
H&E, Alizarin Red Staining and immunostaining of osteopontin and osteocalcin markers H&E, Alcian Blue staining and immunostaining of collagen type II
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Albumin (PAA, Pasching, Austria)). An isotype control (Mouse IgG2a and IgG1, BD Pharmingen, Bedford, USA) or primary antibody was added and the mixture was incubated at room temperature for 30 min. We used the mouse primary anti-human CD90 antibody, which reacted with porcine antigens (1:500, Clone 5E10, BD Pharmingen), anti-human CD14 (1:10, Clone M5E2, BD Pharmingen), anti-pig CD29 (1:10, Clone NaM160-1A3, BD Pharmingen), and anti-pig CD45 (1:20, Clone K252.1E4, abdSerotec). FITC Goat anti-mouse IgG/IgM (BD Pharmingen) was used to detect primary antibodies. Flow cytometry analysis was performed using CyAnADP (Dako) and Summit 4.3 software.
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expression of type II collagen (MonosanXtra) in fibrin chondrogenic differentiation. All micrographs of pellets and fibrin scaffolds were taken with an Axiophot Zeiss Microscope and a Nikon Digital Sight D5-U1 camera. In addition, two scaffolds (control and differentiation) from each experiment were cut and fixed in 2.5% glutaraldehyde (Sigma Aldrich), dehydrated in a graded ethanol series, and sputter-coated with gold for visualization by scanning electron microscopy (SEM, JEOL 6100).
Multilineage differentiation The P1 porcine dermal cells were cultured in vitro with specific differentiation media (Table 1) to determine multipotency; cells cultured in complete culture medium were used as a control. Differentiation media were added on the third day in culture after which the media were changed every 2–3 days. Each experiment was repeated 12 times.
Traditional differentiation methods: monolayers and pellets Adipogenic and osteogenic induction was applied to cells growing in monolayers at a density of 104 cells/cm2 over 21 days of culture. All micrographs of monolayer cultures were taken with an inverted microscope (Leica DMIL) and a Leica DFC 300FX camera on days 7, 14, and 21 of culture. In adipogenic differentiation, we used 0.35% (w/v) Oil Red O dye (Sigma Aldrich) and in osteogenic differentiation the cells were stained with 1% (w/v) Alizarin Red dye (Sigma Aldrich). Chondrogenic differentiation was induced in pellets over 21 days at a density of 106 cells/pellet. Pellets were collected on day 21, fixed in 4% phosphate-buffered formalin, embedded in paraffin, and cut into 4 mm sections. The sections were stained with Hematoxylin & Eosin (H&E, Merck, Darmstadt, Germany) and Alcian Blue (Sigma Aldrich). Finally, to detect the expression of type II collagen we used type II anti-human collagen antibody (1:50; MonosanXtra, Uden, Netherlands).
Adipogenic, osteogenic and chondrogenic differentiation in fibrin scaffolds Fibrin scaffolds were collected at 14, 21, 28, and 35 days of culture, fixed in 4% phosphate-buffered formalin, embedded in paraffin, and cut into 4 mm sections. In the three groups of differentiation, sections were stained with H&E (Merck) to reveal the histological structure. The cells of fibrin scaffolds used in the adipogenic differentiation studies were stained with Oil Red O (Sigma Aldrich) to determine the production of lipid droplets and the scaffolds were frozen with Tissue-Tek OCT (Sakura, Kampenhout Belgium) and cut with cryostat at 6 mm. In osteogenic differentiation, sections were stained with Alizarin Red (Sigma Aldrich) to determine mineralization as sign of matrix ossification in osteogenic differentiation. To confirm osteogenic differentiation we detected the expression of osteopontin (1:50, Thermo Scientific, Cheshire, UK) and osteocalcin (1:50, Thermo Scientific, Cheshire, UK). In chondrogenic differentiation, sections were stained with Alcian Blue (Sigma Aldrich) to determine mucopolysacharide production in chondrogenic differentiation. Finally, we detected the
Fig. 1 – Characterization of porcine dermal cells: (A) SEM image reveals cell attachment of pMSCs throughout the autologous fibrin scaffolds (AFSs), together with the internal porous structure. Scale bar¼ 5 lm. (B) Proliferative capacity of pMSCs expressed as cumulative cell-doubling. (C) Numbers of dermal cells cultured in TCPS and inside the fibrin scaffold along 15 days in culture. All values are means7SD. n ¼12 (po0.05 from 12 to 15 days in culture).
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Statistical analysis Data are expressed as means7standard deviation. The SPSS Statistics 17.0 program was used to determine statistical significance between groups; po0.05 was considered significant. The Kolmogorov–Smirnov and Levene tests were used in the parametric normality and homoscedasticity assumptions respectively, after checking the criteria for parametric analysis, which was performed using Student’s t-test for independent samples.
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2.0870.47; P6, 2.2270.47; P7, 2.1570.55; P8, 2.2170.50; P9, 2.0570.49; P10, 1.9170.50), and hence cells could be continuously passaged every 5 days for 10 passages (Fig. 1B). The cell numbers of pMSCs on Tissue Culture Polystyrene (TCPS) and in AFSs were similar along the first 9 days of culture. The proliferative capacity in AFSs was more evident than in TCPS as from 12 days (po0.05), and the number of cells on TCPS decreased at 15 days, while in the AFSs they increased further (po0.05) (Fig. 1C).
Expression of surface markers
Results Isolation and culture The cells had a spindle-shaped morphology, and these cells exhibited a homogeneous distribution in the autologous fibrin scaffolds (AFSs). The porous structure of the fibrin allowed cell adhesion (Fig. 1A). The proliferative capacity of porcine MSCs (pMSCs), expressed as cell-doubling showed that the cells divided approximately twice during each passage (The cell doubling P1 was 1.9970.58; P2, 2.0970.64; P3, 2.1570.60; P4, 2.0770.60; P5,
Dermal cells were characterized by flow cytometry for the expression of the following markers. The cells expressed the CD90 (99.867 0.11%) and CD29 (99.9470.04%) markers of the mesenchymal lineage, but did not express the CD45 (1.3870.43%) and CD14 (1.58 70.52%) hematopoietic cell lineage-specific surface markers (Fig. 2).
Adipogenic differentiation Oil Red O dye confirmed the presence of intracellular red lipid droplets in adipogenic differentiated cells in monolayers
Fig. 2 – Flow cytometry for the surface markers CD14, CD29, CD45, and CD90. Porcine MSCs were positive for CD90 and CD29, whereas they were negative for CD14, and CD45. Purple curves represent the distributions of cells incubated with primary antibodies. The isotype control is represented by a black-line. All values are means7SD. n ¼ 5. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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(Fig. 3A), with an evident change in morphology towards adipocytes; these were not seen in cells maintained in complete culture medium (Fig. 3B). We observed the morphology of adipocytes in fibrin adipogenic differentiation scaffolds with H&E staining (Fig. 3C), and Oil-Red-O staining (Fig. 3E) confirmed the intracellular accumulation of red lipid droplets, which was not seen in the controls (H&E (Fig. 3D) and Oil-Red-O (Fig. 3F)).
Osteogenic differentiation Osteogenic differentiation as evidenced by the presence of calcium-rich red deposits following Alizarin Red staining revealed an evident mineralization of osteogenic differentiated cells in monolayers (Fig. 4A), which was not observed in the controls (Fig. 4B). In the fibrin osteogenic differentiation scaffolds a change in morphology occurred (Fig. 4C) and calcium deposits, as a sign of matrix mineralization, were shown up by Alizarin Red staining (Fig. 4E), which was not the case of cells cultured in complete medium (H&E (Fig. 4D) and Alizarin Red (Fig. 4F)). The immunohistochemistry of osteopontin revealed that the cellular production of osteopontin was more evident in the fibrin
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scaffolds with differentiation to the osteogenic lineage (Fig. 4G) than in controls (Fig. 4H), and the focal expression of osteocalcin was detected in differentiation scaffolds (Fig. 4I); this was not seen in the controls (Fig. 4J).
Chondrogenic differentiation Pellets with chondrogenic differentiation showed foci of cell necrosis (Fig. 5A) and the large majority had disintegrated, while in the fibrin scaffolds a good chondrogenic differentiation was obtained (Fig. 5B). H&E staining revealed that cell morphology was compatible with chondroblasts, flat cells with small and displaced nuclei, which were located in lacunae in the case of the cells of fibrin chondrogenic differentiation scaffolds (Fig. 5B). In contrast, the controls had a spindle-shaped morphology (Fig. 5C). Alcian Blue staining revealed mucopolysaccharides synthesized by the chondrogenic cell type in the fibrin scaffolds with differentiation to the chondrogenic lineage (Fig. 5D), which was not seen in the controls (Fig. 5E). The immunohistochemistry of type II collagen revealed that the cellular production of collagen fibers was more evident in the differentiation scaffolds (Fig. 5F) than in the cells maintained in complete culture medium (Fig. 5G).
Fig. 3 – Oil-Red-O staining (A and B) of the adipogenic differentiation of porcine MSCs at 14 days in culture for monolayers (A) and controls (B). Note the morphology of the adipocyte (box A). H&E staining (C and D) and Oil-Red-O staining (E and F) of porcine MSCs at 28 days in culture for fibrin adipogenic differentiation scaffolds (C–E) and fibrin controls (D–F). Scale bar: 100 lm (A and B) and 50 lm (C–F).
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Fig. 4 – Alizarin Red staining (A and B) of the adipogenic differentiation of porcine MSCs at 14 days in culture for monolayers (A) and controls (B). H&E staining (C and D), Alizarin Red staining (E and F), and immunohistochemistry of osteopontin (G and H) and osteocalcin (I and J) of porcine MSCs at 28 days in culture for fibrin osteogenic differentiation scaffolds (C, E, G, and I) and fibrin controls (D, F, H, and J). Scale bar: 100 lm (A and B) and 50 lm (C–J).
4.
Discussion
The search for new cell sources for use as substitutes of native cells is one of the main objectives of tissue engineering and regenerative medicine. Owing to their low immunogenicity MSCs are immunoprivileged cells for both autologous and allogenic transplantation. Dermal porcine MSCs are an attractive cell source for tissue engineering and regenerative medicine owing to their easy isolation and abundance, their suitable proliferative capacity, and their ability to differentiate in vitro into osteoblasts,
chondrocytes and adipocytes. In addition, the anatomy and physiology of the pig are very similar to those of the human. For dermis-derived MSCs, using different culture systems several multipotent cell populations have been identified in rodents as well as in humans. Low-temperature preserved human foreskin biopsies have afforded dermal mesenchymal stem cells [17]. Skin-derived precursors (SKPs) other than mesenchymal stem cells have been isolated from mammalian and human dermis [12], neonatal rat dermis enzymatically dissociated isolated dermal multipotent cells (DMCs) [10,11]. In human foreskin
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Fig. 5 – H&E staining (A) of the chondrogenic differentiation of porcine MSCs at 21 days in the pellet culture system (A). H&E staining (B and C), Alcian Blue staining (D and E), and immunohistochemistry of type II collagen (F and G) of porcine MSCs at 35 days in culture for fibrin chondrogenic differentiation scaffolds (B, D, and F) and fibrin controls (C, E, and G). Scale bar: 50 lm.
biopsies, nestin vimentinþ fibroblasts have been observed, representing a new concept of multipotent stem cells [18]. Since 2009 another new concept has been introduced: cutaneous mesenchymal stem cells. In these, the connective tissue sheath and the papilla of the hair follicle probably represent their anatomical niche [19]. Here we attempted to confirm the results reported for humans in adult porcine dermis. We observed that dermal porcine MSCs had a typical spindle shape, a good proliferative capacity, and that pMSCs expressed the cell markers of the mesenchymal lineage, such as CD90 (Thy-1) and CD29 (Integrin b1). Determination of specific surface antigen expression for pMSCs is complicated because the minimum criteria for defining MSCs have been applied in humans, together with some markers such as CD105 or CD73 (positive 495%), the clone not being present in pigs. To identify pMSCs and not confuse them with
other cells, we assessed the lack of expression of hematopoietic markers such as CD45 (a pan-leukocyte marker) and CD14. Unfortunately, the cellular implantation of cells differentiated by classical methods as monolayers involves the generation of a single cell suspension, which involves a risk of cell necrosis and loss of the cells’ differentiation properties. The pellet system showed foci of cell necrosis and proved to be a non-reproducible method. These pitfalls are usually overcome by the use of tissueengineered implantable scaffolds capable of favoring cellular growth on different biomaterials by providing a 3D structure for the cells to adhere, to proliferate and differentiate. Our AFSs, which are highly biocompatible, readily available and easy to process, have proved to be an optimal substrate for these purposes [13,20,21]. The proliferative capacity of dermal pMSCs in AFSs was good; after 15 days, the cell population had
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increased 100-fold. Thus, we obtained large cell numbers in a short period of time. Additionally, our analyses confirmed the cellular plasticity of porcine dermal cells when cultured in adipogenic, osteogenic, and chondrogenic induction media in AFSs. We observed intracellular lipid droplets in adipogenic differentiation; an evident matrix mineralization with calcium deposits and osteogenic markers, such as osteopontin and osteocalcin, were seen in osteogenic differentiation. Finally, chondrocytes located in lacunae and the production of mucopolysacchrides and type II collagen were observed in chondrogenic differentiation. Adipogenic differentiation is easily obtainable in monolayer cultures, but only some authors have attempted differentiation using 3D structures [22]. The use of scaffolds in osteogenic differentiation has risen in the last decade because a three-dimensional structure is required to support bone formation [22–24]. Pellet culture systems and scaffolds are necessary for chondrogenic differentiation [25–27]. Previously Ohgushi et al. used a ceramic engineered tissue for the treatment of osteoarthritis [28]. In this procedure, the artificial scaffold was incubated in differentiation medium to stimulate MSC differentiation, after which the scaffold was implanted. We speculated that it would be interesting to demonstrate cell multidifferentiation to the adipogenic, osteogenic and chondrogenic lineages within an autologous and implantable 3D structure, where we could implant the same previously differentiated cells. Accordingly, we believe that the present study provides sufficient evidence for the effectiveness of 3D culture and the differentiation of dermal pMSCs in autologous fibrin scaffolds (AFSs).
5.
Conclusions
The porcine dermis is a good source of MSCs, with a suitable proliferative capacity and expression of mesenchymal lineage markers. pMSCs also showed optimum adhesion, proliferation and differentiation in our AFSs, and hence they might serve as a promising autologous and implantable material for use in tissue engineering.
Acknowledgments The authors thank the staff from the Laboratory of Microscopy of the University of Leo´n, Angustias Pe rez, a technician at the Pathology Services of University Hospital of Salamanca, and Marta Regueiro at the School of Veterinary Sciences in Leo´n for her excellent care of the animals used in the study.
Appendix Figures with essential color discrimination. Figs. 2–5 in this article are difficult to interpret in black and white. The full color images can be found in the on-line version.
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[25] D. Bosnakovski, M. Mizuno, G. Kim, T. Ishiguro, M. Okumura, T. Iwanaga, T. Kadosawa, T. Fujinaga, Chondrogenic differentiation of bovine bone marrow mesenchymal stem cells in pellet cultural system, Exp. Hematol. 32 (2004) 502–509. [26] G.I. Im, Chondrogenesis from mesenchymal stem cells derived from adipose tissue on the fibrin scaffold, Curr. Appl. Phys. 5 (2005) 438–443. [27] H.A. Awad, M.Q. Wickham, H.A. Leddy, J.M. Gimble, F. Guilak, Chondrogenic differentiation of adipose-derived adult stem cells in agarose, alginate and gelatin scaffolds, Biomaterials 25 (2004) 3211–3222. [28] H. Ohgushi, N. Kotobuki, H. Funaoka, H. Machida, M. Hirose, Y. Tanaka, Y. Takakura, Tissue engineered ceramic artificial joint—ex vivo osteogenic differentiation of patient mesenchymal cells on total ankle joints for treatment of osteoarthritis, Biomaterials 26 (2005) 4654–4661.
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Available online at www.sciencedirect.com
journal homepage: www.elsevier.com/locate/yexcr
Research Article
Differentiation within autologous fibrin scaffolds of porcine dermal cells with the mesenchymal stem cell phenotype ˜ab, Marta Lo´peza, Jennifer Ramosa, Javier Iglesiasa Pilar de la Puentea,n, Dolores Luden a
´n, Spain Tissue Bank, San Francisco Clinic Foundation, Av./Facultad 51, 51, 24004 Leo Pathology Service, University Hospital of Salamanca, P/San Vicente 58-182, 37007 Salamanca, Spain
b
article information
abstract
Article Chronology:
Porcine mesenchymal stem cells (pMSCs) are an attractive source of cells for tissue engineering
Received 13 July 2012
because their properties are similar to those of human stem cells. pMSCs can be found in
Received in revised form
different tissues but their dermal origin has not been studied in depth. Additionally, MSCs
17 October 2012
differentiation in monolayer cultures requires subcultured cells, and these cells are at risk of
Accepted 20 October 2012
dedifferentiation when implanting them into living tissue. Following this, we attempted to
Available online 1 November 2012
characterize the MSCs phenotype of porcine dermal cells and to evaluate their cellular
Keywords:
proliferation and differentiation in autologous fibrin scaffolds (AFSs). Dermal biopsies and
Porcine mesenchymal stem cells
blood samples were obtained from 12 pigs. Dermal cells were characterized by flow cytometry.
Dermal cells
Frozen autologous plasma was used to prepare AFSs. pMSC differentiation was studied in
Fibrin scaffolds
standard structures (monolayers and pellets) and in AFSs. The pMSCs expressed the CD90 and
Differentiation
CD29 markers of the mesenchymal lineage. AFSs afforded adipogenic, osteogenic and chondrogenic differentiation. The porcine dermis can be proposed to be a good source of MSCs with adequate proliferative capacity and a suitable expression of markers. The pMSCs also showed optimal proliferation and differentiation in AFSs, such that these might serve as a promising autologous and implantable material for use in tissue engineering. & 2012 Elsevier Inc. All rights reserved.
Introduction Mesenchymal stem cells (MSCs) are a heterogeneous subset of stromal stem cells that have the capacity for self-renewal, and are multipotent and display immunological features, antiinflammation properties, and trophic effects [1,2]. In view of these properties, MSCs can be considered as an ideal candidate cell type for tissue engineering and regenerative medicine. The International Society for Cellular Therapy (ISCT) has defined the minimum criteria for defining human MSCs [3]. They must be plastic-adherent when maintained in standard culture conditions; Z95% of the MSC population must express CD105, CD73, and CD90, and they must lack the expression (r2% n
positive) of CD45, CD34, CD14 or CD11b, Cd79a or CD19, and HLA class II. Furthermore, the cells must be able to differentiate into adipocytes, osteoblasts and chondrocytes under standard in vitro differentiating conditions. Most stem cells have been described in studies carried out on humans and small animals. Owing to the similarity in size and anatomical features of pigs and humans, porcine stem cells may offer a more attractive possibility for evaluating the efficacy and applicability of MSCs. Porcine MSCs (pMSCs) have also been found in bone marrow [4–6], amniotic fluid [7], umbilical cord blood [7], muscle [8], and fetal skin [9]. However, not much is known about the stem cells of the porcine dermis. Multipotent cell populations in skin have been identified from different locations, such as epidermal stem cells
Corresponding author. Postal address: Paseo Salamanca 59, 21B, 24010, Leo´n, Spain. Fax: þ34 987218801. E-mail address: [email protected] (P. de la Puente).
0014-4827/$ - see front matter & 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.yexcr.2012.10.009
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from the epidermis and hair follicle stem cells from appendages [10]. Despite this, the dermis may represent a larger reservoir for adult stem cells than the epidermis and the hair follicles together [11]. In 2001, mesenchymal stem cells in the dermis were first isolated in mice and later they were also found in the human dermis [12]. We believe that the dermis might provide an abundant alternative source of MSCs for autologous and allogenic stem cell therapy, because the dermis is a large organ and is readily accessible. Differentiation in monolayer cultures is the technique most frequently referred to in the literature, but for these cells to be suitable for implantation, prior subculture is required. Subculture involves the dissociation of the cells from each other and the substrate in order to generate a suspension of single cells, which are therefore at risk of losing their differentiation properties. These pitfalls are usually overcome by the use of tissueengineered implantable scaffolds, capable of stimulating cellular growth in different biomaterials by providing a 3D structure so that the cells can adhere, proliferate and differentiate. We have recently developed easy-to-process, readily available and totally autologous fibrin scaffolds (AFSs) [13], in which cells are able to integrate and adhere themselves, and proliferate within a biocompatible and porous structure. Although some types of fibrin scaffold have long been used in tissue engineering for many applications [14], scaffolds of autologous fibrin with platelet-poor plasma or platelet-rich plasma represent an excellent biological support for stromal cells, whereas those made from commercial fibrins show poor stromal cell survival [15]. Some studies have shown that human embryonic stem cells [16] and human stromal stem cells [15] may be successfully differentiated in fibrin scaffolds, but very little is known about their properties as regards the differentiation of porcine MSCs in such structures. Accordingly, here we were prompted to assess the ability of porcine dermal cells to differentiate into mesenchymal lineages when cultured inside AFSs. A further aim was to check the mesenchymal stem cell phenotype and multilineage potential of these cells.
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All animals received humane care in compliance with the 86/609/CEE European Economic Community Directive and the 1201/2005 Spanish Royal Decree at the Veterinary School of Leo´n. The experimental protocol was reviewed and approved by the Research Committee of the University of Leo´n. The animals were premedicated intramuscularly with 0.35 mg/kg Midazolam (Normon, Madrid, Spain) and 5 mg/kg ketamine (Imalgene, Marquez, Mexico). Anesthesia was maintained with inhaled 2% isofluorane (Isoflo, Abbot, Chicago, USA). The animals were not euthanized after the study, because they were used for other experiments not directly linked to the present study. Dermal biopsies and blood were obtained as previously described by us [13]: small pieces of dermis (1–3 g) were incubated with type I collagenase (Sigma Aldrich, St. Louis, USA) at 2 mg/ml for 22–24 h. 104 dermal cells/cm2 were seeded in six-well plates and cultured in complete culture medium (Table 1), which was refreshed every 2–3 days. Cells were subcultured every 5 days using trypsin/EDTA (T/E, PAA, Pasching, Austria).The proliferative capacity of porcine MSCs was evaluated by cell doublings, using the following formula: log (N)/log (2), where N¼cells harvested/cells seeded. The results are expressed as cumulative cell doublings. Blood samples of 25 ml from each animal were centrifuged at 1620g for 10 min, after which the plasma was frozen until it was used to prepare the fibrin scaffolds. AFSs were formed by mixing 400 ml/ml plasma (autologous from each animal), 104 cells/ml (autologous from each animal), 4 mg/ml tranexamic acid (Rottapharm, Valencia, Spain) and 4 mg/ml calcium chloride (Braun, Barcelona, Spain) diluted in complete medium to elaborate 2 ml of fibrin gel, as described previously [13]. P1 cells were counted every 3 days using the Trypan blue (Sigma Aldrich) exclusion method in order to obtain a growth curve of the cells in the autologous fibrin scaffolds (AFSs), treated enzymatically with type I collagenase (Sigma Aldrich) at 2 mg/ml for 24 h at 37 1C, and in tissue culture polystyrene (TCPS) treated enzymatically with trypsin/EDTA at 0.05/0.02% for 3 min at 37 1C.
Materials and methods Flow cytometry Isolation and culture of pMSCs Dermis and blood from Large-White crossbred pigs of 3 months of age (n¼ 12, 15–35 kg) were extracted under sterile conditions.
Cells at the 2–3 passage were trypsinized, centrifuged, and 500,000 cells were resuspended in PBS/BSA buffer (Phosphate Buffer Saline (Sigma Aldrich), pH 7.4, and 1% Bovine Serum
Table 1 – Composition of culture and differentiation media, and methods to assess lineage differentiation (DMEM, Dulbecco’s Modified Eagle Medium; FBS, Fetal Bovine Serum; IBMX, 3-isobutyl-1-methylxanthine; TGF-b1, Transforming growth factor b1). Culture medium
Complete culture medium
Supplements
Histochemical and immunohistochemical analysis
DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM
Controls with the same techniques as induction cells. H&E, and Oil Red O staining
L-Glutamine
Adipogenic differentiation medium Osteogenic differentiation medium Chondrogenic differentiation medium
DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ100 nM Dexamethasomeþ0,5 mM IBMXþ10 mg/ml Insulinþ200 mM Indomethacinþ500 mM ascorbate 2-phosphate DMEMþ10% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ10 nM Dezamethasomeþ500 mM ascorbate 2phosphateþ10 mM b-glycerophosphate DMEMþ1% FBSþ100 U/ml Penicillimþ100 mg/ml Streptomicinþ2 mM L-Glutamineþ10 ng/ml TGF b1þ6,25 mg/ml Insulinþ50 mg/ml ascorbate 2-phosphateþ100 nM Dexamethasome
H&E, Alizarin Red Staining and immunostaining of osteopontin and osteocalcin markers H&E, Alcian Blue staining and immunostaining of collagen type II
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Albumin (PAA, Pasching, Austria)). An isotype control (Mouse IgG2a and IgG1, BD Pharmingen, Bedford, USA) or primary antibody was added and the mixture was incubated at room temperature for 30 min. We used the mouse primary anti-human CD90 antibody, which reacted with porcine antigens (1:500, Clone 5E10, BD Pharmingen), anti-human CD14 (1:10, Clone M5E2, BD Pharmingen), anti-pig CD29 (1:10, Clone NaM160-1A3, BD Pharmingen), and anti-pig CD45 (1:20, Clone K252.1E4, abdSerotec). FITC Goat anti-mouse IgG/IgM (BD Pharmingen) was used to detect primary antibodies. Flow cytometry analysis was performed using CyAnADP (Dako) and Summit 4.3 software.
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expression of type II collagen (MonosanXtra) in fibrin chondrogenic differentiation. All micrographs of pellets and fibrin scaffolds were taken with an Axiophot Zeiss Microscope and a Nikon Digital Sight D5-U1 camera. In addition, two scaffolds (control and differentiation) from each experiment were cut and fixed in 2.5% glutaraldehyde (Sigma Aldrich), dehydrated in a graded ethanol series, and sputter-coated with gold for visualization by scanning electron microscopy (SEM, JEOL 6100).
Multilineage differentiation The P1 porcine dermal cells were cultured in vitro with specific differentiation media (Table 1) to determine multipotency; cells cultured in complete culture medium were used as a control. Differentiation media were added on the third day in culture after which the media were changed every 2–3 days. Each experiment was repeated 12 times.
Traditional differentiation methods: monolayers and pellets Adipogenic and osteogenic induction was applied to cells growing in monolayers at a density of 104 cells/cm2 over 21 days of culture. All micrographs of monolayer cultures were taken with an inverted microscope (Leica DMIL) and a Leica DFC 300FX camera on days 7, 14, and 21 of culture. In adipogenic differentiation, we used 0.35% (w/v) Oil Red O dye (Sigma Aldrich) and in osteogenic differentiation the cells were stained with 1% (w/v) Alizarin Red dye (Sigma Aldrich). Chondrogenic differentiation was induced in pellets over 21 days at a density of 106 cells/pellet. Pellets were collected on day 21, fixed in 4% phosphate-buffered formalin, embedded in paraffin, and cut into 4 mm sections. The sections were stained with Hematoxylin & Eosin (H&E, Merck, Darmstadt, Germany) and Alcian Blue (Sigma Aldrich). Finally, to detect the expression of type II collagen we used type II anti-human collagen antibody (1:50; MonosanXtra, Uden, Netherlands).
Adipogenic, osteogenic and chondrogenic differentiation in fibrin scaffolds Fibrin scaffolds were collected at 14, 21, 28, and 35 days of culture, fixed in 4% phosphate-buffered formalin, embedded in paraffin, and cut into 4 mm sections. In the three groups of differentiation, sections were stained with H&E (Merck) to reveal the histological structure. The cells of fibrin scaffolds used in the adipogenic differentiation studies were stained with Oil Red O (Sigma Aldrich) to determine the production of lipid droplets and the scaffolds were frozen with Tissue-Tek OCT (Sakura, Kampenhout Belgium) and cut with cryostat at 6 mm. In osteogenic differentiation, sections were stained with Alizarin Red (Sigma Aldrich) to determine mineralization as sign of matrix ossification in osteogenic differentiation. To confirm osteogenic differentiation we detected the expression of osteopontin (1:50, Thermo Scientific, Cheshire, UK) and osteocalcin (1:50, Thermo Scientific, Cheshire, UK). In chondrogenic differentiation, sections were stained with Alcian Blue (Sigma Aldrich) to determine mucopolysacharide production in chondrogenic differentiation. Finally, we detected the
Fig. 1 – Characterization of porcine dermal cells: (A) SEM image reveals cell attachment of pMSCs throughout the autologous fibrin scaffolds (AFSs), together with the internal porous structure. Scale bar¼ 5 lm. (B) Proliferative capacity of pMSCs expressed as cumulative cell-doubling. (C) Numbers of dermal cells cultured in TCPS and inside the fibrin scaffold along 15 days in culture. All values are means7SD. n ¼12 (po0.05 from 12 to 15 days in culture).
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Statistical analysis Data are expressed as means7standard deviation. The SPSS Statistics 17.0 program was used to determine statistical significance between groups; po0.05 was considered significant. The Kolmogorov–Smirnov and Levene tests were used in the parametric normality and homoscedasticity assumptions respectively, after checking the criteria for parametric analysis, which was performed using Student’s t-test for independent samples.
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2.0870.47; P6, 2.2270.47; P7, 2.1570.55; P8, 2.2170.50; P9, 2.0570.49; P10, 1.9170.50), and hence cells could be continuously passaged every 5 days for 10 passages (Fig. 1B). The cell numbers of pMSCs on Tissue Culture Polystyrene (TCPS) and in AFSs were similar along the first 9 days of culture. The proliferative capacity in AFSs was more evident than in TCPS as from 12 days (po0.05), and the number of cells on TCPS decreased at 15 days, while in the AFSs they increased further (po0.05) (Fig. 1C).
Expression of surface markers
Results Isolation and culture The cells had a spindle-shaped morphology, and these cells exhibited a homogeneous distribution in the autologous fibrin scaffolds (AFSs). The porous structure of the fibrin allowed cell adhesion (Fig. 1A). The proliferative capacity of porcine MSCs (pMSCs), expressed as cell-doubling showed that the cells divided approximately twice during each passage (The cell doubling P1 was 1.9970.58; P2, 2.0970.64; P3, 2.1570.60; P4, 2.0770.60; P5,
Dermal cells were characterized by flow cytometry for the expression of the following markers. The cells expressed the CD90 (99.867 0.11%) and CD29 (99.9470.04%) markers of the mesenchymal lineage, but did not express the CD45 (1.3870.43%) and CD14 (1.58 70.52%) hematopoietic cell lineage-specific surface markers (Fig. 2).
Adipogenic differentiation Oil Red O dye confirmed the presence of intracellular red lipid droplets in adipogenic differentiated cells in monolayers
Fig. 2 – Flow cytometry for the surface markers CD14, CD29, CD45, and CD90. Porcine MSCs were positive for CD90 and CD29, whereas they were negative for CD14, and CD45. Purple curves represent the distributions of cells incubated with primary antibodies. The isotype control is represented by a black-line. All values are means7SD. n ¼ 5. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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(Fig. 3A), with an evident change in morphology towards adipocytes; these were not seen in cells maintained in complete culture medium (Fig. 3B). We observed the morphology of adipocytes in fibrin adipogenic differentiation scaffolds with H&E staining (Fig. 3C), and Oil-Red-O staining (Fig. 3E) confirmed the intracellular accumulation of red lipid droplets, which was not seen in the controls (H&E (Fig. 3D) and Oil-Red-O (Fig. 3F)).
Osteogenic differentiation Osteogenic differentiation as evidenced by the presence of calcium-rich red deposits following Alizarin Red staining revealed an evident mineralization of osteogenic differentiated cells in monolayers (Fig. 4A), which was not observed in the controls (Fig. 4B). In the fibrin osteogenic differentiation scaffolds a change in morphology occurred (Fig. 4C) and calcium deposits, as a sign of matrix mineralization, were shown up by Alizarin Red staining (Fig. 4E), which was not the case of cells cultured in complete medium (H&E (Fig. 4D) and Alizarin Red (Fig. 4F)). The immunohistochemistry of osteopontin revealed that the cellular production of osteopontin was more evident in the fibrin
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scaffolds with differentiation to the osteogenic lineage (Fig. 4G) than in controls (Fig. 4H), and the focal expression of osteocalcin was detected in differentiation scaffolds (Fig. 4I); this was not seen in the controls (Fig. 4J).
Chondrogenic differentiation Pellets with chondrogenic differentiation showed foci of cell necrosis (Fig. 5A) and the large majority had disintegrated, while in the fibrin scaffolds a good chondrogenic differentiation was obtained (Fig. 5B). H&E staining revealed that cell morphology was compatible with chondroblasts, flat cells with small and displaced nuclei, which were located in lacunae in the case of the cells of fibrin chondrogenic differentiation scaffolds (Fig. 5B). In contrast, the controls had a spindle-shaped morphology (Fig. 5C). Alcian Blue staining revealed mucopolysaccharides synthesized by the chondrogenic cell type in the fibrin scaffolds with differentiation to the chondrogenic lineage (Fig. 5D), which was not seen in the controls (Fig. 5E). The immunohistochemistry of type II collagen revealed that the cellular production of collagen fibers was more evident in the differentiation scaffolds (Fig. 5F) than in the cells maintained in complete culture medium (Fig. 5G).
Fig. 3 – Oil-Red-O staining (A and B) of the adipogenic differentiation of porcine MSCs at 14 days in culture for monolayers (A) and controls (B). Note the morphology of the adipocyte (box A). H&E staining (C and D) and Oil-Red-O staining (E and F) of porcine MSCs at 28 days in culture for fibrin adipogenic differentiation scaffolds (C–E) and fibrin controls (D–F). Scale bar: 100 lm (A and B) and 50 lm (C–F).
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Fig. 4 – Alizarin Red staining (A and B) of the adipogenic differentiation of porcine MSCs at 14 days in culture for monolayers (A) and controls (B). H&E staining (C and D), Alizarin Red staining (E and F), and immunohistochemistry of osteopontin (G and H) and osteocalcin (I and J) of porcine MSCs at 28 days in culture for fibrin osteogenic differentiation scaffolds (C, E, G, and I) and fibrin controls (D, F, H, and J). Scale bar: 100 lm (A and B) and 50 lm (C–J).
4.
Discussion
The search for new cell sources for use as substitutes of native cells is one of the main objectives of tissue engineering and regenerative medicine. Owing to their low immunogenicity MSCs are immunoprivileged cells for both autologous and allogenic transplantation. Dermal porcine MSCs are an attractive cell source for tissue engineering and regenerative medicine owing to their easy isolation and abundance, their suitable proliferative capacity, and their ability to differentiate in vitro into osteoblasts,
chondrocytes and adipocytes. In addition, the anatomy and physiology of the pig are very similar to those of the human. For dermis-derived MSCs, using different culture systems several multipotent cell populations have been identified in rodents as well as in humans. Low-temperature preserved human foreskin biopsies have afforded dermal mesenchymal stem cells [17]. Skin-derived precursors (SKPs) other than mesenchymal stem cells have been isolated from mammalian and human dermis [12], neonatal rat dermis enzymatically dissociated isolated dermal multipotent cells (DMCs) [10,11]. In human foreskin
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Fig. 5 – H&E staining (A) of the chondrogenic differentiation of porcine MSCs at 21 days in the pellet culture system (A). H&E staining (B and C), Alcian Blue staining (D and E), and immunohistochemistry of type II collagen (F and G) of porcine MSCs at 35 days in culture for fibrin chondrogenic differentiation scaffolds (B, D, and F) and fibrin controls (C, E, and G). Scale bar: 50 lm.
biopsies, nestin vimentinþ fibroblasts have been observed, representing a new concept of multipotent stem cells [18]. Since 2009 another new concept has been introduced: cutaneous mesenchymal stem cells. In these, the connective tissue sheath and the papilla of the hair follicle probably represent their anatomical niche [19]. Here we attempted to confirm the results reported for humans in adult porcine dermis. We observed that dermal porcine MSCs had a typical spindle shape, a good proliferative capacity, and that pMSCs expressed the cell markers of the mesenchymal lineage, such as CD90 (Thy-1) and CD29 (Integrin b1). Determination of specific surface antigen expression for pMSCs is complicated because the minimum criteria for defining MSCs have been applied in humans, together with some markers such as CD105 or CD73 (positive 495%), the clone not being present in pigs. To identify pMSCs and not confuse them with
other cells, we assessed the lack of expression of hematopoietic markers such as CD45 (a pan-leukocyte marker) and CD14. Unfortunately, the cellular implantation of cells differentiated by classical methods as monolayers involves the generation of a single cell suspension, which involves a risk of cell necrosis and loss of the cells’ differentiation properties. The pellet system showed foci of cell necrosis and proved to be a non-reproducible method. These pitfalls are usually overcome by the use of tissueengineered implantable scaffolds capable of favoring cellular growth on different biomaterials by providing a 3D structure for the cells to adhere, to proliferate and differentiate. Our AFSs, which are highly biocompatible, readily available and easy to process, have proved to be an optimal substrate for these purposes [13,20,21]. The proliferative capacity of dermal pMSCs in AFSs was good; after 15 days, the cell population had
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increased 100-fold. Thus, we obtained large cell numbers in a short period of time. Additionally, our analyses confirmed the cellular plasticity of porcine dermal cells when cultured in adipogenic, osteogenic, and chondrogenic induction media in AFSs. We observed intracellular lipid droplets in adipogenic differentiation; an evident matrix mineralization with calcium deposits and osteogenic markers, such as osteopontin and osteocalcin, were seen in osteogenic differentiation. Finally, chondrocytes located in lacunae and the production of mucopolysacchrides and type II collagen were observed in chondrogenic differentiation. Adipogenic differentiation is easily obtainable in monolayer cultures, but only some authors have attempted differentiation using 3D structures [22]. The use of scaffolds in osteogenic differentiation has risen in the last decade because a three-dimensional structure is required to support bone formation [22–24]. Pellet culture systems and scaffolds are necessary for chondrogenic differentiation [25–27]. Previously Ohgushi et al. used a ceramic engineered tissue for the treatment of osteoarthritis [28]. In this procedure, the artificial scaffold was incubated in differentiation medium to stimulate MSC differentiation, after which the scaffold was implanted. We speculated that it would be interesting to demonstrate cell multidifferentiation to the adipogenic, osteogenic and chondrogenic lineages within an autologous and implantable 3D structure, where we could implant the same previously differentiated cells. Accordingly, we believe that the present study provides sufficient evidence for the effectiveness of 3D culture and the differentiation of dermal pMSCs in autologous fibrin scaffolds (AFSs).
5.
Conclusions
The porcine dermis is a good source of MSCs, with a suitable proliferative capacity and expression of mesenchymal lineage markers. pMSCs also showed optimum adhesion, proliferation and differentiation in our AFSs, and hence they might serve as a promising autologous and implantable material for use in tissue engineering.
Acknowledgments The authors thank the staff from the Laboratory of Microscopy of the University of Leo´n, Angustias Pe rez, a technician at the Pathology Services of University Hospital of Salamanca, and Marta Regueiro at the School of Veterinary Sciences in Leo´n for her excellent care of the animals used in the study.
Appendix Figures with essential color discrimination. Figs. 2–5 in this article are difficult to interpret in black and white. The full color images can be found in the on-line version.
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