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. 2019 Jan 25;43(2):147–157. doi: 10.1002/cbin.11072

Human neonatal stem cell‐derived skin substitute improves healing of severe burn wounds in a rat model

Ruhma Mahmood 1,2,✉, Azra Mehmood 1, Mahmood S Choudhery 1, Sana Javaid Awan 1, Shaheen N Khan 1, Sheikh Riazuddin 1,2
PMCID: PMC13397396  PMID: 30443955

Abstract

Conventional approaches can repair minor skin injuries; however, severe burn injuries require innovative approaches for efficient and better wound repair. Recent studies indicate that stem cell‐based regenerative therapies can restore severe damaged skin both structurally and functionally. The current study aims to evaluate the wound healing potential of skin substitute derived from human neonatal stem cells (hNSCs) using a severe burn injury rat model. Amniotic epithelial cells (AECs) and mesenchymal stem cells (MSCs) were isolated from placenta (a source of neonatal stem cells) by explant culture method. After characterization, AECs and umbilical cord‐MSCs were differentiated into keratinocyte and fibroblasts, respectively. Morphological changes, and expression of corresponding keratinocyte and fibroblast specific markers were used to verify differentiation into respective lineage. A skin substitute was developed by mixing hNSCs‐derived skin cells (hNSCs‐SCs) in plasma for transplantation in a rat model of severe burn injury. Results indicated that placenta‐derived AECs and MSCs were efficiently differentiated into skin cells, that is, keratinocytes and fibroblasts, respectively, as indicated by morphological changes, immunostaining, and polymerase chain reaction analysis. Further, transplantation of hNSCs‐SCs seeded in plasma significantly improved basic skin architecture, re‐epithelization rate, and wound healing concurrent with reduced apoptosis. In conclusion, neonatal stem cell‐derived skin substitute efficiently improved severe burn wounds in a rat model of burn injury. Unique properties of placenta‐derived stem cells make them superlative candidates for the development of “off‐the‐shelf” artificial skin substitutes for future use.

Keywords: burn injury model, epithelial cells, mesenchymal stem cells, neonatal stem cells

Abbreviations

AECs

amniotic epithelial cells

hNSCs

human neonatal stem cells

hNSCs‐SCs

human neonatal stem cells‐derived skin cells

pDFs

pre‐differentiated fibroblasts

pDKs

pre‐differentiated keratinocytes

pDSLCs

pre‐differentiated skin like cells

UC‐MSCs

umbilical cord mesenchymal stem cell

Introduction

Severe burn injuries are difficult to manage due to multiple problems ranging from pain to infection. The loss of skin integrity results in compromised skin functions and therefore needs immediate treatment especially for severe burn injuries. In severe burn injuries, healing process does not start suddenly due to damage to both skin layers (Wooldridge and Surveyer, 1980). Moreover, in severe burns, healing starts from wound margins, which is a time‐taking process with greater risk of infections. Moreover, due to large burn surface area, there is less availability of auto‐graft donor sites in such patients. Therefore, novel remedies that are available immediately for such severe burn injuries are required. Recent research in this regard is promising and has created a hope for patients with incurable diseases.

Embryonic, adult, as well as neonatal tissues are the main sources of stem cells. Neonatal stem cell sources such as placenta appear more appropriate for severe burn injuries as they are readily available in large numbers. In addition, as compared to adipose tissue and bone marrow, acquisition of placenta poses no risk to donor and it contains large number of regenerative cells. Furthermore, stem cells isolated from placenta possess immunomodulatory and immunosuppressive properties (Insausti et al., 2014), which make these cells ideal even for allogenic use and for making “off‐the‐shelf” skin substitutes (Cargnoni et al., 2009).

Stem cells characteristics such as release of specific growth factors and cytokines, and division and differentiation into functional tissues support the healing process efficiently. In the current study, the use of skin substitute developed using pre‐differentiated skin cells [i.e., pre‐differentiated fibroblasts (pDFs) and pre‐differentiated keratinocytes (pDKs)] was proposed for spontaneous healing. Stem cells already committed to skin lineage will start the healing process soon after transplantation. This study explores wound healing effect of skin substitute derived from pDKs and pDFs using a rat model of severe burn injury. Amniotic epithelial cells (AECs) and umbilical cord mesenchymal stem cells (UC‐MSCs) isolated by explant culture method were characterized and induced into keratinocytes and fibroblasts, respectively. Differentiation of AECs and UC‐MSCs into respective cell types was confirmed using parameters such as morphological changes, immunostaining, and polymerase chain reaction (PCR). A rat model of severe burn injury was developed using a hot brass rod. The pDKs and pDFs were mixed with rat blood plasma to be used as a matrix for easy transport and application of cells on surface of rat wounds. Differentiation of cells (AECs and UC‐MSCs) into respective lineage was confirmed by morphological changes, reverse transcription‐PCR (RT‐PCR), and immunostaining. Further, transplantation of neonatal stem cell‐derived skin substitutes in a rat burn injury model significantly improved skin architecture, re‐epithelization rate, and wound healing. The study design will provide new opportunities for the development of skin substitutes that may be available off‐the‐shelf for repair of severe burn wounds.

Materials and methods

Collection of human placenta

Human placenta were collected after informed consent and processed according to the protocols approved by institutional review board (IRB) at CEMB, University of The Punjab, Lahore, Pakistan and were processed within 2 h to isolate AECs and UC‐MSCs.

Isolation and culturing of amniotic epithelial cells (AECs)

For AECs isolation, amniotic membranes were separated from the placenta and washed with PBS. The amniotic membrane minced into small pieces (∼1–2 cm2) was taken in culture flask and incubated in a humidified CO2 incubator at 37°C (Raeder et al., 2007; Mahmood et al., 2015). Following culture medium was used for AECs expansion: DMEM, 10% FCS, and 1% penicillin/streptomycin (Sigma–Aldrich, USA). Without disturbing the tissue pieces, the medium was replaced within 48 h.

Mesenchymal stem cells (MSCs) isolation from cord tissue pieces

For MSCs isolation, a non‐enzymatic digestion technique was used (Choudhery et al., 2013a; Mahmood et al., 2015). Briefly, minced cord tissue pieces were cultured in Dulbecco's modified Eagle's medium (DMEM) containing 1% antibiotics (penicillin/streptomycin) solution (Sigma–Aldrich) and 10% FBS (fetal bovine serum). The culture conditions were 37°C and 5% CO2 under humidity. Minced cord pieces were discarded when cell colonies were observed (after 8–10 days).

Differentiation of AECs and UC‐MSCs

After isolation, both types of cells, that is, AECs and UC‐MSCs were differentiated into keratinocytes and fibroblasts, respectively. Differentiation into skin cells (fibroblasts and keratinocytes) was assessed by morphological changes, PCR, and immunocytochemistry as described below.

Induction of amniotic epithelial cells (AECs) into keratinocytes

A total of 10,000 AECs at passage 2 were cultured in culture plates (six‐well) in AECs expansion medium as mentioned in section Isolation and culturing of amniotic epithelial cells (AECs). After 24 h, AECs expansion medium was replaced with keratinocyte induction medium that consisted of DMEM+HAM F12 (3:1), 10% FCS, 1% solution of penicillin/streptomycin, 0.5 mg/mL hydrocortisone, 1% insulin transferrin (Sigma–Aldrich), and 15 ng/mL KGF (keratinocytes growth factor, Invitrogen, USA) as described previously by us (Mahmood et al., 2015). This induction medium was changed after every 72 h for 15 days.

Induction of cord tissue‐derived mesenchymal stem cells (UC‐MSCs) into fibroblasts

At passage 2, UC‐MSCs (1 × 104) were cultured using six‐well plate in MSCs expansion medium as mentioned in section Mesenchymal stem cells (MSCs) isolation from cord tissue pieces. For induction of UC‐MSCs, following medium (fibroblast induction medium) was used: DMEM containing 10% FCS, 1% streptomycin/penicillin solution, 5.0 μg/mL insulin, and 1 ng/mL bFGF (Sigma–Aldrich) as described previously by us (Mahmood et al., 2015). Cells were cultured in this induction medium for 15 days, with medium changed twice a week.

RT‐PCR analysis of induced keratinocytes and fibroblasts

RT‐PCR was carried out to identify the expression of lineage‐specific genes in induced fibroblast‐like and keratinocyte‐like cells. Briefly, 15 days after initiation of induction, total RNA was extracted using TRIzol reagent (Invitrogen) and cDNA was synthesized using 1.5 μg of RNA using RT enzyme (Invitrogen). The expression of CK1, CK10, CK14, and E‐Cad was assessed to confirm differentiation of cells into keratinocytes‐like cells, while Coll‐3, Desmin, FGF7, and vimentin were used to confirm differentiation of UC‐MSCs into fibroblast‐like cells. β‐actin was used as an internal control. Primer sequences and their lengths in base pairs have been given in Table 1. After electrophoresis ImageJ software (http://rsbweb.nih.gov/ij/) was used to quantify gel bands.

Table 1.

Primer sequences for genes

Genes 5′–3′ sequence Size (bp)
BAX AGAGGTCTTTTTCCGAGTGG (F) 95
CAGTTTGCTGGCAAAGTAGAAA (R)
Beta actin CGCATGGGTCAGAAGGATTC (F) 137
TAGAAGGTGTGGTGCCAGATTT (R)
CK1 GGAGGAGGAGGTGGTAGATTTT (F) 388
GAGGTTGCTGATGTATGACTCG (R)
CK 10 GAGCAAGGAACTGACTACAG (F) 249
CTCGGTTTCAGCTGCAATCT (R)
CK 14 TGCTATTGGTGTCAGGGAAG (F) 277
GTGGCAAGGTTCTTTTCTCC (R)
Collagen‐3 GTTGACCCTAACCAAGGATGCA (F) 203
GGAAGTTCAGGATTGCCGTAG (R)
E‐Cadherin CACCTGGAGAGAGGCCATGT (F) 187
TGGGAAACATGAGCAGCTCT (R)
FADD CTGCGGGAGTAGTTGGAAAGT (F) 241
GGAAATGGGACAAAACATCCT (R)
FGF‐7 TGGTGAAGTTCATGGATGTCTATC (F) 212
CACAGGATGGCTTGAAGATGTA (R)
Desmin CATCCTCAAGAAGGTGTTGGAG (F) 112
CAAAGAGACGTGGGACGAGT (R)
Oct‐4 GGCGTTCTCTTTGGAAAGGTGTTC (F) 145
CTCGAACCACATCCTTCTCT (R)
Nanog GGACGCGTGGGGGCTGGAGAC (F) 174
GGCTCGAGGGGGACCAGGAAG (R)
PCNA CATCCTCAAGAAGGTGTTGGAG (F) 112
CAAAGAGACGTGGGACGAGT (R)
SSEA4 CCGCGTCAAGAGGCCCATGAA (F) 148
CCCGCTTCTCGGTCTCGGACAA (R)
VEGF TGGTGAAGTTCATGGATGTCTATC(F) 114
CACAGGATGGCTTGAAGATGTA (R)
Vimentin CTGCGGGAGTAGTTGGAAAGT (F) 241
GGAAATGGGACAAAACATCCT (R)
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Immunocytochemistry

We further performed immunostaining using fibroblast‐specific markers (collagen 3; Santa Cruz Biotechnology, USA), desmin (Santa Cruz Biotechnology), FAPα (Abcam, UK), procollagen 1 (Santa Cruz Biotechnology), and keratinocyte‐specific markers (CK5; Abcam), CK10 (Abcam), Involucrin (Abcam), Loricrin (Abcam) at 1:200. Briefly, induced cells were fixed with 4% paraformaldehyde (PFA) and incubated overnight with primary antibodies at 4°C. Next day primary antibodies were washed with PBS and cells were incubated for 60 min with respective secondary antibody in an oven at 37°C. Afterwards the cell nuclei were stained with DAPI (4′,6‐diamidino‐2‐phenylindole; Sigma–Aldrich) and observed under Olympus BX‐61 equipped with DP‐70 digital camera (Olympus, Japan) to take images.

Rat model of full thickness burn injury

Burn injury model was developed to evaluate the effect of pre‐differentiated skin‐like cells (pDSLCs) for treatment and repair of burn injury. Briefly, SD rats (n = 12) aged 9–12 months were anesthetized with Ketasol and xylazene solution, and hair were removed from dorsal side. A brass rod (2 × 2 cm) was placed in water at 100°C for 15 min and applied immediately on shaved area for 20 s as described (Singer et al., 2009). Two injuries, one on right side and other on left side, were given to each rat. Wounds were excised 2 days after injury under anesthesia using scalpel and blade. Immediately after excision, plasma‐based bioengineered skin substitute containing differentiated keratinocyte‐like and fibroblast‐like cells was applied on excised wounds, one side and the side served as control in each rat. All the protocols for heat injury, excision, and handling of rats were approved by the IRB at the CEMB, Lahore, Pakistan.

Preparation of plasma‐based bioengineered skin substitute

Blood (6–8 mL) was collected from rats and centrifuged to collect plasma. A total of 3.33 mL plasma was taken in a new tube and 10 mg/mL tranexamic acid (40 μL) was added to make a plasma gel. 2.5 × 104 fibroblast‐like cells and 700 μL of 1% CaCl2 solution were taken in a tube and mixed. This mixture was taken in a petri plate and was then placed for 60 min at 37°C in 5%CO2 to allow it to semi‐solidify. A total of 5 mL fibroblast expansion medium was added and plates were placed in CO2 incubator at standard condition for 24 h.

Keratinocyte‐like cells (derived from AECs) were trypsinized (passage 2) and 3 × 104 cells were overlaid on plasma (containing fibroblasts) and again incubated for 24 h before it is was finally applied on excised wounds of rats.

Application of plasma‐based skin substitute to excised burn wound

Bioengineered skin substitutes (2 × 2 cm) were applied on excised wounds on one side. To avoid infections, paraffin guaze was used to cover wounds. Finally the wounds were covered with Tegaderm bandage (Health Care, USA).

Assessment of wound healing

Re‐epithelization rate

Re‐epithelization rate after application of pSCs‐derived‐pDSLCs was determined. Rats were observed visually every day, and days that control wounds and pSCs‐derived‐pDSLCs‐treated wounds take to re‐epithelize completely were counted.

Measurement of wound healing

Wound healing assay was performed at days 5, 10, and 15 to measure reduction in wounded area (Gebrehiwot et al., 2015). Briefly, a transparent sheet of plastic was placed on wounds of anesthetized rats that was positioned with ventral side down. The wounded area was marked through on the sheet using pointer. Percentage reduction in wound size was calculated by using the initial and current values of wound.

Histological analysis of skin sections from wounds

Three weeks after transplantation, 0.5 cm2 skin samples was taken from the wounded area and fixed in formalin (Sigma, USA). Skin samples were dehydrated, embedded in wax, and cut using a microtome (5 μm). H&E was performed to stain the skin section and to observe skin architecture.

Further the skin sections were incubated with Annexin V, CK10, and Coll1 primary antibodies overnight at 4°C followed by 1‐h incubation with relevant secondary antibodies at 37°C. Vectashield mounting media containing DAPI was used to mount the stained skin section. The sections were observed under Olympus BX‐61 microscope in at least 10 randomly chosen fields at a magnification of 200×.

RNA isolation from skin and quantitative real‐time

Skin samples taken from the control and treated wounds were minced and treated with TRIzol Reagent (Invitrogen) to isolate total RNA. cDNA synthesized using reverse transcriptase system and 1.5 μg of RNA was used to perform PCR. We used SYBR Green PCR Super Mix (Fermentas, USA) on BioRad System iQ5. Normalized fold expression of PCNA, VEGF, FADD, and Bax was measured.

Statistical analysis

Graphpad Prism version 5 was used for statistical analysis of data. The data are shown as mean ± standard deviation. To compare two groups, unpaired t‐test was carried out. P values <0.05 were considered statistically significant.

Results

Characterization of AECs and UC‐MSCs

Both types of cells (AECs and UC‐MSCs) were isolated using explant culture method. Cell outgrowth from both types of tissues (amniotic membrane and umbilical cords) was observed 3–5 days of initial culture. Both AECs and UC‐MSCs grew into colonies and exhibited plastic adherent growth. Morphologically, AECs were round in shape (Supplementary Figure 1A), while UC‐MSCs displayed typical spindle‐shaped morphology (Supplementary Figure 1B). In addition, AECs displayed a positive expression of cytokeratins such as CK8, CK18, and CK19 (Supplementary Figure 1C), while UC‐MSCs were strongly positive for CD49, CD73, and CD90 (Supplementary Figure 1D). Furthermore as shown in Supplementary Figure 1E, both types of cells were positive for NANOG, SSEA4, and OCT4, indicating their embryonic origin.

AECs and UC‐MSCs exhibited differentiation into skin cells

After 15 days in induction, medium AECs were differentiated into keratinocytes. Differentiation of AECs into keratinocytes was indicated by the positive expression of E‐Cadherin, CK1, CK10, and CK14 (Supplementary Figures 2A and 2B). Additionally, the differentiated AECs showed positive expression for Loricrin, CK5, CK10, and Involucrin as indicated by immunostaining of induced cultures (Supplementary Figure 2C).

Figure 2.

Figure 2

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Re‐epithelization after transplantation of pre‐differentiated cells in rat model of burn injury. Re‐epithelization was improved in rat wounds in which pre‐differentiated cells were transplanted (right) as compared to control wounds (left). Re‐epithelization in rats at day 0 (A), day 10 (B), and day 20 (C) has been shown. Graphical representation of wound contraction in control and transplanted wounds at days 10 and 20 (D). The rate of re‐epithelization in various groups has been shown in panel E. Values were expressed as mean ± SD. P < 0.05 was considered significant.

RT‐PCR analysis and immunostaining revealed successful differentiation of UC‐MSCs into fibroblasts. PCR results demonstrated a significant higher expression of desmin, coll‐3, vimentin, and FGF‐7 (Supplementary Figures 2D and 2E) in induced cultures of UC‐MSCs as compared to respective control. As shown in Supplementary Figure 2F, the immunostaining results indicated positive expression of collagen 3, desmin, FAPα, and procollagen 1.

Development of plasma‐based skin substitute

Plasma‐based skin substitutes were prepared by using pDKs and pDFs. Figure 1A shows the morphology of pDKs in the plasma gel. A plasma‐based skin substitute containing both types of pre‐differentiated cells, that is, pDKs and pDFs, have been shown in Figure 1B. When solidified, these skin substitutes appeared suitable in terms of ease to transfer to wounds and for “off‐the‐shelf” use.

Figure 1.

Figure 1

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Development of skin substitute. A skin substitute was developed using pre‐differentiated fibroblasts and pre‐differentiated keratinocytes in a plasma gel matrix. (A) Fibroblasts within the plasma gel. (B) Skin substitute with keratinocytes cultured on top of the gel. Rat burn injury model (C). A rat burn injury model was developed using a hot brass rod (100°C). On back of each rat, two wounds were created. One such wound served as control (left) with no transplantation, while other (right) was transplanted with plasma‐based skin substitute made of pre‐differentiated skin‐like cells (keratinocyte‐like and fibroblast‐like cells).

On back of each rat two wounds were created (Figure 1C) side by side. In each rat, left‐sided wound served as control and only bandage was applied, while on right‐sided wound, plasma‐based skin substitute (made of pDKs and pDFs) was applied. In both groups, all animals were survived after application of plasma‐based skin substitute.

Wound healing and rate of re‐epithelization after transplantation of pre‐differentiated cells

The burn injured rats showed better healing ability when treated with plasma‐based skin substitutes. Re‐epithelization of injured skin was evaluated after different time periods (after 10 and 20 days). The re‐epithelization was high in wounds that were transplanted with a skin substitute made of hNSCs‐derived skin cells (Figures 2A–2C, right) as compared to control (Figures 2A–2C, left). Rate of wound healing at day 10 was 78.0 ± 1.00% in transplanted group as compared to 31.80 ± 1.76% in control wounds. After 20 days, control group showed 87.57 ± 1.30% in control as compared to 97.6 ± 0.61% contraction in wounded area of transplanted wound (Figure 2C). Additionally, complete healing of wounds in control group occurred after 27.67 ± 2.51 days, while in pre‐differentiated group, it occurred after 20.00 ± 2.00 days (Figure 2D).

Skin architect and expression of CK10 and Coll1 after transplantation of skin substitute

Figures 3A–3D show H&E staining of skin sections. The results indicate a more well‐preserved architect in wounds transplanted with hNSCs‐derived skin cells (Figure 3A) as compared to control wounds (Figure 3B). For comparison, skin section from normal skin was used a control (Figure 3C).

Figure 3.

Figure 3

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H&E staining and expression of CK10 and Coll1 after transplantation of pre‐differentiated skin‐like cells. Results indicated more well‐preserved architect in wounds transplanted with pre‐differentiated skin‐like cells (A) as compared to control wounds (B). Panel C shows skin section from normal skin. Expression of CK10 and Coll1 in control and treatment groups after transplantation was determined by immunostaining. Panel D shows expression of CK10 in normal skin epidermis. As compared to control (E), transplanted (F) skin tissue of injured rats showed more intact epidermis. The expression of collagen 1 in normal, control, and transplanted wounds is been shown in panels G–I, respectively. The results indicated that the collagen 1 expression was significantly reduced in control (H) as compared to transplanted samples (I).

Reconstruction of new epidermis in normal, control, and transplanted wounds was evaluated by CK10 staining. Figure 3D shows expression of CK10 in normal skin epidermis. As compared to control (Figure 3E), transplanted (Figure 3F) skin tissue of injured rats showed more intact epidermis. The expression of collagen 1 in normal, control, and transplanted wounds has been shown in Figures 3G–3I, respectively. The results indicated that the collagen 1 expression was significantly reduced in control (Figure 3H) as compared to transplanted samples (Figure 3I).

Transplantation of skin substitute decreases apoptosis and increases expression of PCNA and VEGF

There were more apoptotic cells (Annexin‐V positive) in sections of skin taken from wounds in which no substitute was applied (Figure 4A) as compared to wounds transplanted with skin substitute derived from pDKs and pDFs (Figure 4B). Figure 4C shows apoptosis in normal rat skin. A significant reduction in apoptotic cells was observed in wounds transplanted with skin‐substitute made of neonatal pre‐differentiated skin cells as compared to control.

Figure 4.

Figure 4

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Assessment of apoptosis in different treatment groups after transplantation of pre‐differentiated skin‐like cells. Apoptosis, as measured by expression of annexin V, was significantly higher in control (A) as compared to wounds transplanted with pre‐differentiated skin‐like cells (B). The expression of annexin V in skin samples of normal rat skin has been shown in panel C. Similarly, real‐time polymerase chain reaction analysis of skin samples after transplantation of pre‐differentiated skin‐like cells was performed (D). The expression of BAX, FADD, that are pro‐apoptotic genes, decreased in skin samples that were transplanted with a skin substitute. Contrary to the expression of pro‐apoptotic genes, the expression of PCNA and VEGF was significantly up‐regulated after treatment with skin‐substitutes. P < 0.05 was considered significant.

Similarly, the expression of BAX and FADD that are pro‐apoptotic genes decreased in skin samples that were transplanted with a skin substitute made of pre‐differentiated skin cells. RT‐PCR analysis indicated that BAX expression was significantly reduced in transplanted wounds (1.547 ± 0.39) as compared to control wounds (2.850 ± 0.18). Similarly, the expression of FADD was 2.23 ± 0.47 in transplanted wounds and 4.00 ± 0.23 in controls. Contrary to the expression of Bax and FADD, the expression of PCNA and VEGF was significantly up‐regulated after treatment with skin‐substitutes (hNSCs‐SG). The expression of PCNA was 2.30 ± 0.35 in transplanted wounds, while in controls, it was 0.72 ± 0.07. VEGF expression was increased from 0.723 ± 0.24 in control to 2.39 ± 0.57 in wounds transplanted with skin substitute.

Discussion

Thermal injuries are common worldwide especially in developing countries like Pakistan. Severe burn injuries not only damage the skin and may expose internal organs but also may result in life‐threatening infections. Traditional treatment options take long time to heal severe wounds and thus long‐term exposure to external environment may poses risk of infections, which is a serious concern for health‐care providers and patients. Therefore, treatment of such severe burn patients in shortest possible time can save infections and in turn patient's life. The aim of the current study is, therefore, to develop a skin substitute using readily available source of cells, that is, placenta, which is neonatal source of stem cells.

We propose that severe burn injuries require an immediate mechanism to start repair. In the current study, already committed cells to skin lineages (keratinocytes and fibroblasts) were used that can meet the above‐mentioned requirement. The major conclusions of this study are: (i) AECs and UC‐MSCs derived from placenta efficiently differentiate into skin cells (keratinocytes and fibroblasts, respectively); and (ii) burn wounds in a rat model were more efficiently healed when plasma‐based skin substitute (prepared using in vitro differentiated keratinocytes and fibroblasts) was applied.

Neonatal stem cells such as those derived from human placental tissue are readily available and can be isolated in large numbers for use in tissue engineering and regenerative medicine. As placentas are readily available, they can be collected without any risk to donor. Various parts of placenta have already been investigated by researcher for use in tissue engineering purposes (Kesting et al., 2008a,b). The properties of cells isolated from placenta such as high proliferation, multi‐lineage differentiation, and immunomodulation and immunosuppression make them superlative candidates for regenerative medicine applications even for allogenic use. Considering the objective of the study and above‐mentioned properties, full‐term placentas were selected to isolate the cells (AEC and MSCs) for differentiation into skin cells and to make a skin substitute for burn‐injured skin. In the current study, two types of cells, that is, AECs and MSCs, were isolated from amniotic membrane and cord tissue, respectively. These two types of cells were selected to differentiate into respective skin‐like cells considering similarities in basic origin and features.

We used explants culture method to isolate these cells from tissue pieces as it is inexpensive and gives pure population of cells. Morphologically, AECs were oval shaped (Barbati et al., 2012) similar to other cells of epithelial origin (Noruddin et al., 2007). Similarly, immunostaining of AECs indicated a positive expression of several cytokeratins (CK8, CK18, CK19). Similar findings have been observed by others (Fatimah et al., 2012). UC‐MSCs exhibited spindle‐shaped morphology that resembles with morphology of MSCs derived from other sources such as bone marrow (Shetty et al., 2010). In addition, UC‐MSCs were positive for CD49, CD73, and CD90 as previously reported (Dominici et al., 2006; Pratama et al., 2011; Choudhery et al., 2013b). Both AECs and UC‐MSCs were negative for CD45, which is a marker of hematopoietic cells (Dominici et al., 2006). The positive expression of stemness markers (OCT4, NANOG, SSEA4) by AECs and UC‐MSCs indicates that these cells have qualities of stem cells with embryonic origin (Miki et al., 2007; Simat et al., 2008; Chen et al., 2011).

After isolation of AECs and UC‐MSCs, both types of cells were differentiated into skin‐like cells, that is, AECs into keratinocytes and UC‐MSCs into fibroblasts. Differentiation of both types of cells was triggered using respective medium for 15 days and differentiation into functional cells was assessed using parameters such as morphological changes, immunostaining, and RT‐PCR. AECs when differentiated into keratinocytes exhibited a polygonal morphology along with high expression of CK1, CK10, CK14, and E‐Cadherin as compared to control cultures. These results are similar to those already published in literature for cells undergoing differentiation into keratinocytes (Choi et al., 1990; Alkhalaf et al., 1999). UC‐MSCs have immense ability to differentiate into multiple lineages (Choudhery et al., 2013a) and therefore successfully differentiated into fibroblasts in the current study. The differentiated UC‐MSCs were more elongated with up‐regulated expression of desmin, FGF7, and vimentin as measured with RT‐PCR. Similar results were shown by Raffa et al. (2012) and Lederle et al. (2006). The successful differentiation of AECs into keratinocytes and UC‐MSCs into fibroblasts indicates that these differentiated cells could be used as seed cells for constructing off‐the‐shelf skin substitutes.

For the transplantation of differentiated cells, skin substitute was prepared using fibroblast‐like and keratinocyte‐like cells. We used rat plasma as a matrix to easily transport and apply the skin substitute on rat wounds. In addition, it has been shown that plasma enhances the viability of skin cells for longer periods (Cetin et al., 2000). The application of skin substitute in a rat model of wound injury healed the wound in less time as compared to control wound. The efficient improvement in wound healing was obvious in terms of quicker re‐epithelization and wound contraction. H&E staining of skin sections from regenerated skin showed better skin architect in treated groups as compared to control. Similarly, positive expression of CK10 and Coll1 indicated enhanced healing of both layers of skin, that is, epidermis and dermis after transplantation of skin substitute. Epidermal keratinocytes strongly express Ck10 (Yang et al., 2016), while newly formed healthy fibroblasts indicate formation of ECM as expressed by Coll1 deposition (Midwood et al., 2004; Noorafshan et al., 2014). Staining of skin sections with annexin V, which is a marker of apoptosis, demonstrated reduced more apoptotic cells in control wounds as compared to wounds that were transplanted with plasma‐based skin substitute made from pre‐differentiated skin‐like cells. Expression of other apoptosis‐related genes, such as BAX and FADD, was reduced after transplantation of skin substitute, while expression of PCNA and VEGF was up‐regulated. Overall results of current study indicate that neonatal cells (AECs and UC‐MSCs) differentiated into skin cells (pDKs and pDFs) improve wound healing in a rat model of burn injury. There are certain limitations of the study such as we have not determined the exact mechanism of wound healing when a skin substitute is applied.

In conclusion, neonatal sources such as placenta are the important and promising source of cells for applications in tissue engineering and regenerative medicine. Placental tissue‐derived cells, that is, AECs and MSCs, can be differentiated into keratinocytes and fibroblasts, respectively. In addition, these pre‐differentiated skin cells efficiently improve severe burn wounds after transplantation in a rate model of burn injury. Furthermore, unique properties of placenta‐derived stem cells make them superlative candidates for “off‐the‐shelf” artificial skin substitutes.

Conflict of interest

None.

Supporting information

Additional supporting information may be found in the online version of this article at the publisher's web‐site.

Figure S1. Morphology of AECs and UC‐MSCs. AECs showed rounded morphology (A), while UC‐MSCs were spindle shaped (B) in morphology. AECs displayed a positive expression of CK8, CK18, and CK 19, while UC‐MSCs were positive for CD49, CD73, and CD90 (C). Both types of cells were negative for hematopoietic marker CD45. Both AECs and UC‐MSCs were positive for stemness markers SSEA4, OCT4, and NANOG (D).

CBIN-43-147-s001.tif (12.5MB, tif)

Figure S2. Differentiation of AECs and UC‐MSCs into keratinocyte‐like and fibroblasts‐like cells in vitro. RT‐PCR analysis showed positive expression of keratinocyte‐specific markers, that is, CK1, CK10, CK14, and E‐Cadherin (A) after differentiation. Quantification of gel bands using ImageJ software showing relative gene expression in pre‐differentiated AECs (B). The induced AECs were CK5, CK10, Involucrin, and loricrin positive as determined by immunostaining (C). UC‐MSCs were differentiated into fibroblasts‐like cells (D–F). RT‐PCR analysis showed positive expression of dermal fibroblasts‐specific markers, that is, Coll 3, Desmin, FGF7, and Vimentin (D) in differentiation medium. Quantification of gel bands showed up‐regulation of these markers in induced cultures (E). Results of immunostaining indicated negative expression of dermal fibroblasts‐specific markers Coll 3, Desmin, FAPα, and Procollagen 1 in untreated UC‐MSCs, while positive expression on differentiated UC‐MSCs (F).

CBIN-43-147-s002.tif (6.5MB, tif)

Acknowledgments and funding

The authors thank their colleagues for the review of this manuscript. This work was supported by research grant from Higher Education Commission (HEC) of Pakistan.

Present address of Mahmood S. Choudhery is Tissue Engineering and Regenerative Medicine Laboratory, Department of Biomedical Sciences, King Edward Medical University, Lahore, Pakistan.

Present address of Sana Javaid Awan is IMBB, University of the Lahore, Lahore, Pakistan.

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Supplementary Materials

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Figure S1. Morphology of AECs and UC‐MSCs. AECs showed rounded morphology (A), while UC‐MSCs were spindle shaped (B) in morphology. AECs displayed a positive expression of CK8, CK18, and CK 19, while UC‐MSCs were positive for CD49, CD73, and CD90 (C). Both types of cells were negative for hematopoietic marker CD45. Both AECs and UC‐MSCs were positive for stemness markers SSEA4, OCT4, and NANOG (D).

CBIN-43-147-s001.tif (12.5MB, tif)

Figure S2. Differentiation of AECs and UC‐MSCs into keratinocyte‐like and fibroblasts‐like cells in vitro. RT‐PCR analysis showed positive expression of keratinocyte‐specific markers, that is, CK1, CK10, CK14, and E‐Cadherin (A) after differentiation. Quantification of gel bands using ImageJ software showing relative gene expression in pre‐differentiated AECs (B). The induced AECs were CK5, CK10, Involucrin, and loricrin positive as determined by immunostaining (C). UC‐MSCs were differentiated into fibroblasts‐like cells (D–F). RT‐PCR analysis showed positive expression of dermal fibroblasts‐specific markers, that is, Coll 3, Desmin, FGF7, and Vimentin (D) in differentiation medium. Quantification of gel bands showed up‐regulation of these markers in induced cultures (E). Results of immunostaining indicated negative expression of dermal fibroblasts‐specific markers Coll 3, Desmin, FAPα, and Procollagen 1 in untreated UC‐MSCs, while positive expression on differentiated UC‐MSCs (F).

CBIN-43-147-s002.tif (6.5MB, tif)

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