As the incidence of diabetes has increased, microvascular dysfunction, delayed wound healing and hard-to-heal ulcer formation caused by the microenvironment of diabetic skin have become common in clinical situations.1,2 However, the treatment of hard-to-heal wounds in patients with diabetes is extremely difficult, and there are few effective treatment methods. Adipose-derived stem cells (ASCs), which are extracted from human adipose tissue and are multipotent, have been used in a number of clinical studies.3,4 Several studies have indicated that stem cells can promote the proliferation of endothelial cells,5 promote angiogenesis6 and restrict fibroblast proliferation and scar formation.7 ASCs have been used in clinical studies of tissue regeneration with some success,8,9,10 but because of the complexity of the diabetic wound environment, the effect of ASC transplantation in diabetic wounds remains unclear. Preliminary experiments have shown that the proliferation of ASCs is inhibited, that the number of apoptotic cells increases, and that differentiation activity decreases in the in vitro diabetic model environment.11 Therefore, we designed experiments to determine whether diabetic ASCs can be used for treating normal or diabetic wounds.
Methods
Ethical approval
In this study, 40 male inbred Wistar rats (16 weeks old, specific-pathogen-free) were used. Feeding and experimental methods met ethical requirements (approval by the Laboratory Animal Ethics Committee of Rui-Jin Hospital, No 148).
Rat grouping and modelling
Rats were randomly divided into a diabetic model group (group D) and a non-diabetic (normal) group (group N). Before the experiment, the rats were kept in abrosia for 12 hours, but were permitted water. After weighing and blood glucose measurement, group D was intraperitoneally injected with streptozocin (STZ) (Sigma, US) buffer solution (45mg/kg), and group N was injected with the same amount of citric acid buffer. After seven days, the blood glucose level of group D was >18mmol/l, indicating successful establishment of the diabetes model.12 We measured blood glucose and body weight for all rats every week and statistically analysed the differences between group D and group N. We also analysed the correlation between body weight and blood glucose of diabetic rats during the 12 weeks using SPSS 19.0 (IBM, US).
Extraction and amplification of ASC-rich cells
At 12 weeks after the establishment of the rat model, three rats from each group (N and D) were randomly selected. Inguinal fat pads were completely removed under abdominal anaesthesia. The large blood vessels and fascia were removed, and the fat pads were weighed and stored at 4°C.13
The fat pads were homogenised in a sterile environment and digested with 0.5% type I collagenase (Gibco, US) for 30 minutes. The resulting suspension was centrifuged at 180g for five minutes, and the pellets were cultured in ASC amplification medium (Mesenchymal Stem Cell Growth Medium Ready-to-use; PromoCell, Germany).
After 24 hours of incubation, the suspended cells were removed. On day three, adherent cells were counted. Adherent cells were cultured to the third generation (passage 3, P3).
P3 cells from the D and N groups were microscopically observed (×400 magnification). In each sample, 10 fields were randomly selected, and the aged cells (identified as having an enlarged nucleus, impurities in the cytoplasm, increased volume and >3 pseudopodia) were counted. In each group, three samples were counted and the data were statistically analysed.
The P3 cells had a nearly fusiform shape and grew quickly, so we named the p3 cells which contained the most adipose stem cells ‘rich adipose stem cells’ (ASC-rich cells). They were then used for wound transplantation. ASC-rich cells were labelled by CD31, CD34, CD44 and CD105 fluorescent antibody (Abcam, US) and detected by flow cytometry. P3 ASC-rich cells and rich diabetic adipose stem cells (d-ASC-rich cells) were stained by AnnexinV/PI (BD Falcon, US) to detect the percentage of early, late and total apoptotic ASC-rich cells in each group, respectively, by using flow cytometry. From each group, four samples were collected.
Extraction and enrichment of nuclear cells from adipose tissue
From group N, three rats were randomly selected and their bilateral inguinal fat pads were homogenised. After type I collagenase digestion, the suspension was filtered through a nylon filter (200 mesh, 70μm; BD Falcon, US) to enrich nuclear cells.14 The filtered suspension underwent density centrifugation (at 180g for five minutes). The nuclear cell layer in the deposit was collected and resuspended in phosphate-buffered saline (PBS). The suspension of ‘nucleus-enriched cells’ contained 2×107 nuclear cells/ml and was stored at 4°C for transplantation. Nucleus-enriched cell transplantation is the most common way in clinic to heal wounds. The nuclear cell forms the majority of the enriched cells and plays a critical role in wound healing. We named the nucleus-enriched cells by density centrifugation and filtration ‘adipose nuclear cells’ (ANCs).
Some of the ANCs were fluorescent labelled with CD31, CD34, CD44 and CD105 antibodies, and the expression of surface antigens was evaluated by flow cytometry. The other ANCs were ready for transplantation.
Production and enrichment of ASC and enriched nuclear cell transplantation in rat wound models
After removing the back hair of diabetic and normal rats (9×9cm), we drew a straight line along the spine with a perpendicular line through its midpoint, thereby dividing the back skin into four regions. In the centre of each region, a 12mm-diameter borer was used to make a full-thickness skin defect.
Each wound edge was divided into eight equal parts. The cell suspension was injected into the dermal and subdermal layer of each part (0.05ml for each part, 2×107 cells/ml), and another 0.1ml of cell suspension was injected into the centre of the defect wound. In total, approximately 107 cells were transplanted into each wound.
The four wounds in the back of the diabetic rats and normal rats were injected with ASC-rich cells, d-ASC-rich cells, enriched ANCs or PBS. We named the ASC-rich cell transplanted wounds ‘ASC wounds’, the d-ASC-rich cell transplanted wounds ‘d-ASC wounds’, the ANC transplanted wounds ‘ANC wounds’, and the PBS-injected wounds ‘control wounds’.
After cell transplantation, water gel wound dressing (Comfeel Plus Ulcer Dressing, Coloplast, Denmark) was used to cover all wounds. Bandages were tied and fixed externally.
Observation of the wound-healing process and statistical analysis
The sizes of wounds were observed 24 hours, three days, seven days and 12 days after injury. From each group at each time point, three rats were randomly selected and whole-skin tissue samples from the back wound edges were collected. The samples were fixed in formalin for 48 hours before being stained with haematoxylin and eosin (H&E; Sigma Chemical, US) and microscopically observed.
The thicknesses of skin, subcutaneous fat layer, dartos fascia and connective tissue layer were measured by AxioVision 4.0 software (Carl Zeiss Vision, UK).
Adobe Photoshop CS5 (Adobe Systems Inc., US) was used to measure the number of pixels in a 2×2cm area; the area of the wound was measured by counting the number of pixels. The formula is as follows:
Wound area (cm2) = 4P1/P2
Where:
- P1=number of pixels in the whole wound area
- P2=number of pixels in the 2×2cm area
Each wound area was measured three times and the average taken. From each wound group, three wound area samples were collected.
All data were analysed by the normal distribution test (W-test) and homogeneity test of variance (F-test) with SPSS 19.0 and the data compared by t-test.
From both group D and from group N, four wound areas were compared at the same time point by a t-test (SPSS 19.0). The thicknesses of new granulation tissue and skin tissue were also measured and compared by t-test (SPSS 19.0).
Results
Body weight and blood glucose changes during modelling
There was a positive correlation between body weight and blood glucose of group D rats after culturing for six weeks. However, a negative linear correlation was found between blood sugar and body weight of group D on week eight (|r|<0.4, low) and week 12 (|r|>0.4 week 12, significant). (R is negative in negative linear correlation, and so |r| is used to express its absolute value.)
The normal rats of group N all showed a positive correlation between body weight and blood glucose at all time points (0.4≤|r|<0.7).
Cell density and growth rate of nuclear cells in diabetic adipose tissue were lower than those in normal tissue
Diabetic rats in group D had smaller fat pads (9.23±1.62g) than normal rats (23.75±1.12g) (p<0.01) (Fig 1a, 1b). Cell counts indicated that the number of primary ANCs on day three in diabetic rats ((0.78±0.11)×106) was statistically significantly lower than that in normal rats ((2.54±0.16)×106) (p<0.01). The cell density of diabetic rats ((0.43±0.07)×105/g) was also decreased compared with that of normal rats ((1.07±0.05)×105/g) (Fig 1c, 1d, 1g; p<0.01). The number of aged cells in group D (9±3 per field) was increased compared with group N (3±2 per field), and the difference was statistically significant (Fig 1e, 1f, 1h; p<0.01).
d-ASC-rich cells showed a higher proportion of apoptosis cells
To further test the underlying mechanism that may explain the lower density of ANCs in diabetic tissue, AnnexinV/PI (BD Falcon, Finland) staining was used to measure the proportion of apoptosis cells in P3 d-ASC-rich and P3 ASC-rich cells. The early apoptosis (AnnexinV+/PI–, 5.68±0.6%), late apoptosis (AnnexinV+/PI+, 0.65±0.13%) and total apoptosis (AnnexinV+/PI– and AnnexinV+/PI+, 6.33±0.53%) of d-ASC-rich cells was higher than that of ASC-rich cells (2.05±2.35%, 0.38±0.13% and 2.43±2.44%, respectively) on passage 3 (Fig 1i; p<0.05).
Expression of characteristic ASC surface antigens on enriched nuclear cells
Next, we compared the ASC positive markers CD105 and CD44, and negative markers CD31 and CD34, on the enriched nuclear cells to those on primary cells. In normal ASC-rich cells (P3 ANCs), the average expression of CD105 was 46.37±2.81%, whereas the average expression of CD105 in primary cells (P1 ANCs) was 15.22±3.85%. This difference was statistically significant (p<0.05) (Fig 2e). The average expression of CD44 on P3 cells was 98.43±0.55%, representing an increase over the average expression of primary cells, which was 74.57±5.58% (p<0.05) (Fig 2f). Normal P3 ANCs did not contain CD31- or CD34-positive cells (<1% positive rate).
Diabetic P3 ANCs contained 38.73±2.89% CD105-positive cells, which is less than the proportion among normal P3 cells (p<0.05) (Fig 2a). P3 ANCs do not express CD31 or CD34. The average CD44 expression was 97.37±0.96%, and there was no significant difference between the D and N groups (p=0.17) (Fig 2b). Compared with the numbers in normal P3 cells, the numbers of CD31- (2.47±0.55%) and CD34-positive (4.63±1.76%) cells were higher among diabetic P3 cells (p<0.05; Fig 2c, 2d). The data above indicate that the number and viability of ASC-rich cells were all decreased in the diabetic environment.
Wound healing after cell transplantation
We tested the effect of cell transplantation to investigate whether a diabetic environment or diabetic ASC-rich cells would change the result of ASC therapy for a skin wound. The water gel wound dressings were removed 24 hours after cell transplantation. Fresh blood scabs could be seen in the wounds, and the wound sizes were not obviously different. On day three, the scabs became darker and harder. By day seven, hard, dry scabs had formed. By day 12, many of the wounds had healed, and the surfaces of the wounds were covered by soft, pink epithelial tissue.
For group D, on day three and day 12, the ASC wound areas (0.63±0.13cm2 and 0.01±0.03cm2, respectively) were smaller than in the control group (0.78±0.08cm2 and 0.04±0.02cm2, respectively) (p<0.05). There was no significant difference in the other wound areas or the other time points.
In group N, on day three, ASC wounds, d-ASC wounds and ANC wounds (0.46±0.32cm2, 0.57±0.29cm2 and 0.52±0.08cm2, respectively) were smaller than the control wounds (0.85±0.29cm2) (p<0.05).
On day seven, the ASC wound areas (0.33±0.07cm2) were smaller than the areas of the d-ASC wounds and ANC wounds (0.5±0.19cm2 and 0.4±0.07cm2, respectively) (p<0.05). There was no difference in the sizes of d-ASC wounds (0.5±0.19cm2) and ANC wounds (0.4±0.07cm2), but both were smaller than the control wounds (0.67±0.16cm2) (p<0.05).
On day 12, the ASC and d-ASC wound areas (0.03±0.02cm2 and 0.06±0.03cm2, respectively) were smaller than control and ANC wounds (0.19±0.07cm2 and 0.14±0.04cm2, respectively) (p<0.01), and the ASC wounds (0.03±0.02cm2) were the smallest (p<0.05). There was no significant difference in size between ANC wounds (0.14±0.04cm2) and the control wounds (0.19±0.07cm2) on day 12 (p=0.063) (Fig 3).
Collectively, ASC-rich cell transplantation increased wound healing in group N (all time points) and group D (day three and day 12). d-ASC-rich cells increased wound healing in group N (all time points), but could not increase the diabetic wound healing in group D. ANC transplantation only increased the wound healing of group N on days three and seven. We conclude that not only the diabetic wound environment, but also d-ASC-rich cells decreased the treatment effect of ASC transplantation in diabetic wound healing.
Histological changes and collagen synthesis in the wound-healing process
We further tested histological changes caused by the diabetic environment. We compared the skin tissue section (H&E) between group N and group D. Diabetic rats in group D had thinner skin (1.98±0.12mm) than group N rats (2.84±0.21mm) (p<0.01). The subcutaneous fat layer was also thinner (diabetic rats 0.08±0.06mm, normal rats 0.36±0.12mm) (p<0.01). The muscle layer and connective tissue layer for the group D rats (0.5±0.16mm) below the fascia were both thinner than those for normal rats (0.77±0.3mm) (p<0.05).
Blood clots composed of fibrinogen and inflammatory cells were found in all wounds at 24 hours (Fig 4a, 4e, 4i, 4m). The scabs in ASC wounds (1.63±0.41mm) were thicker than those in d-ASC wounds (0.53±0.15mm), ANC wounds (0.25±0.14mm) and control wounds (0.1±0.03mm) in group N (p<0.01), which indicates that there were more repair cells involved in the formation of the initial blood scab of ASC wounds. The d-ASC wounds have the second thickest scab (p<0.05), which indicates that there are also many repair cells involved in scab formation, but that the cell number or cell proliferation may be affected by the diabetic environment in which the d-ASC-rich cells were formerly sited.
On day three, more granulation tissue was found in ASC wounds (2.48±0.63mm) than in the control wounds (1.51±0.23mm) in group N (p<0.01). A slice of adipose tissue could be found in this tissue (Fig 4b, arrow). The thickness of the granulation tissue in the four types of wounds did not obviously vary in group D (ASC wound p=0.11; d-ASC wound p=0.062; ANC wound p=0.94) (Fig 5b1, f1, j1, n1). A small amount of dermal tissue grew into the granulation tissue, and epithelial cells migrated on the granulation tissue near the wound margin in ASC wounds (Fig 5b1, arrow). This phenomenon was not observed in the other wound types.
On day seven, the granulation tissue in ASC wounds (1.61±0.37mm) was thicker than in the ANC (1.27±0.23mm) and control wounds (0.69±0.11mm) of group N (p<0.05). There was no difference in thickness between the ASC and d-ASC wounds (p=0.78), but the d-ASC wounds had a lower density of granulation tissue (Fig 4g, arrow), and the connection with the wound edge was looser relative to the ASC wounds (Fig 4c, arrow). The granulation tissue thicknesses of the four wounds in group D did not differ (ASC wound p=0.079; d-ASC wound p=0.058; ANC wound p=0.097) (Fig 5c1, g1, k1, o1). Re-epithelialisation was obvious in the ASC wounds (1.69±0.22mm; control: 1.1±0.61mm) (p<0.05) of group D (Fig 5c1, arrow).
On day 12, the ASC and d-ASC wounds of group N had all healed, with dense, mature granulation tissue that was nearly covered by stratified epithelium (Fig 4d, 4h, arrow). In ANC and control wounds, the granulation tissue was loose, and there were some fat vacuoles in it (Fig 4l, arrow). The ANC and control wounds were not completely covered by epithelium (Fig 4l, p, arrow). In group D, the ASC and d-ASC wounds were all completely repaired by mature granulation tissue and covered with epithelial tissue. There were more hair follicles and newborn dermal tissue in the granulation tissue of ASC wounds, and a layer of continuous, loose connective tissue was formed at its bottom (Fig 5d1, arrow). The ANC and control wounds were incompletely covered by epithelium (Fig 5l1, p1, arrow). This strongly suggested that ASC-rich cell and d-ASC-rich cell transplantation promoted wound re-epithelialisation and epithelial tissue reconstruction.
Discussion
Changes in body weight and blood glucose in diabetic model Wistar rats
Body weight was significantly negatively correlated with blood glucose in group D rats at week 12 (R= –0.510, |r|≥0.4), and the symptoms of type I diabetes were more stable.15 It is also notable that in a diabetic state, disruption of the glycosylation of skin collagen and other proteins is a chronic process; it appears later than haemoglobin glycosylation and has a longer metabolic half-life.16,17 Therefore, a longer duration of the diabetic state may be more conducive to simulating the microenvironment in diabetic hard-to-heal wounds, which might make experimental results under such conditions more reliable.
Characteristic ASC surface antigen expression in nuclear cells after enrichment and amplification
CD105 is a sign of multiple differentiation potential and active endothelial cell proliferation,18,19 and as such is rarely expressed in skin tissue and subcutaneous fat.20 However, the positive expression of CD105 in enriched ANCs (P1) was 15.22%, which shows that CD105-positive ASCs may also be enriched; similar methods may be used clinically. The CD105 expression level in P3-enriched ANCs that were cultured in stem cell growth medium was 46.37%. After culturing ANCs of diabetic rats to the P3 generation, CD105-positive expression was lower than in group N which may indicate the decrease of proliferation and differentiation in d-ASC-rich cells (P3 ANCs).
CD105 expression did not increase in P4 and did not differ from that in P3; in addition, the expression of CD44 (positive 98.43%), CD31 (negative)21 and CD34 (negative)22 was stable in P3 cells, and therefore, we chose P3-enriched cells as the ASC transplantation cells.
CD44 is a cell surface adhesion molecule expressed on the surface of human adipose stem cells (hASCs), while some dying cells or aged cells do not express CD44. The expression of CD44 in d-ASC-rich cells is also a little lower than normal ASC-rich cells, but the differences are not significant. In addition, the cell passage time was prolonged, and the number of aged cells in P3 was higher. More apoptotic d-ASC-rich cells were indicated by AnnexinV/PI staining, than normal ASC-rich cells. These observations may indicate that a diabetic microenvironment can affect the proliferation of ASC-rich cells, reduce cell density, and accelerate the ageing process, in accord with previous experimental results for hASCs in an in vitro simulated diabetes microenvironment. Our previous experiment results23 indicated that d-ASC-rich cells are physiologically and biochemically significantly different from normal ASC-rich cells, which may be caused by the high glucose and irregular glycation in the diabetic environment. The glycation end products may adversely affect the functional properties of proteins, lipids and DNA, and then lead to the changes of d-ASC-rich cells.23,24,25
Limitations
This study is not without limitations. The ASC-rich cells we used were primary cells and consisted of several types of cell; 46.37% was the highest expression of CD105 we could get from stem cell environment culturing. We attempted flow cytometry sorting, but were unable to increase the CD105-positive expression, and the fluorescence labelling before cytometry sorting may have also influenced the cell proliferation. We need to find a simple and practical way of choosing ASCs from the nuclear cells to increase the CD105 expression in the subsequent experiment. Limited by our experimental method, we cannot analyse the effect of other cells in wound healing. However, we can conclude that the increase in cells with stem cell markers will lead to better wound healing than ANC transplantation in which the stem cells are not enriched. The cells with stem cell markers comprised the majority of the enriched cells and may play a critical role in wound healing. In future research, we intend to separate each type of cell in our ‘enriched stem cells’ group, and compare the advantages and disadvantages of those wound repair cells in wound healing.
The expression of CD31 and CD34 in d-ASC-rich cells was positive but relatively low. We also found the ASC-rich cells in vivo in a simulated diabetic environment expressed a low positive level of CD31 and CD34.23,24,25 We found 5% of ASC-rich cells expressed CD34 in our study. Some researchers consider that ASCs are able to express CD34. The CD34+ cells are more proliferative and have a higher ability to form colonies, while CD34− cells have a greater differentiation ability.26,27 The literature also suggests that the number of CD34+ stem cells increases in old thymus fat tissue,28 and that in the simulated fat damage environment29 cell levels of CD34+ expression also increase. Whether the diabetic environment has the same effect on ASCs to increase CD34+ cell levels remains to be determined. The average positive rate of CD31 is very low (2%). False positives cannot be ruled out, or that the CD31+ cells may have originated from the CD34+ cells. This question also needs further research.
Observation and analysis of wound healing in rats
The experimental method described here can produce sufficient cell quantities through amplification, but there is a conflict between laboratory methods and clinical ethics, as certain policies formulated by the Chinese Government forbid using amplified cells in clinical studies or treatments. The most common cell transplantation method typically involves the direct extraction and enrichment of ANCs, followed by introduction into the wound, to avoid ethical and legal problems.30,31 Transplanting enriched ANCs without in vitro culture has become a new treatment for bone defects,32 dental implants,33 cartilage repair,34 lower limb vascular occlusion35 and other diseases. To simulate clinical ANC transplantation, we designed the ANC-transplantation wound method described here. As we discovered in preliminary experiments, sufficient ANCs (at least 107 are needed)36 could not be obtained from the bilateral inguinal fat pads of a single diabetic rat, and so this study only analysed normal ANC wound transplantation in normal and diabetic rats.
Water gel dressing is an advanced dressing that is applied to ulcers and exudation and has a good moisturising effect on the wound.23,24,25 The use of water gel dressings on wounds after cell transplantation has three advantages:
- It ensures that the injected cells remain in the wound after injection by preventing cell suspension leakage or flow to another location
- It keeps the wound moist, preventing cell inactivation or contamination caused by extended exposure to air
- Because the ulcer dressing is thick and soft, it not only keeps the shape of the wound from contracting or deforming, but also increases the comfort of the rats such that they may not struggle as much and thereby affect experimental results.
The use of a splinted wound model can prevent wound contraction and create wounds which heal through granulation and re-epithelialisation in a quantifiable fashion.37 The methods of the splint wound model are various. The use of silicone ring fixation is the most common.38,39 Additionally, many researchers use skin suture, adhesive dressing36 and metal splinting40 to fix the wound.
In our experiment, we used a sticky water gel dressing which can prevent injected cell leakage, prevent wound contraction on the first day, and which may have a similar effect in keeping the wound from contracting as the traditional splint wound model. After 24 hours, in order to minimise the impact on the skin around the wound, we removed the water gel dressing and replaced it with an ordinary dressing, this being more like the clinical treatment after surgery, because a water gel dressing is used only in a wound or ulcer with much exudate. The tension and mechanical stretch of the wound surface will not be changed,41 so the proliferation of local fibroblasts is not affected, and will not influence the stem cell transplantation.42
We use cell injection to transplant ASCs; it can simplify operation steps, reduce the impact on the wound and minimise the effect on surrounding tissues. The cell injection method can only reflect the treatment effect of cells on the wound, which is consistent with the objective of our study. Our research objective was to clarify if diabetic stem cells can treat the diabetic wound and the changes of ASCs in the diabetic environment. Many researchers have used biomaterial scaffolds combined with stem cells to repair wounds;43,44 the use of cell sheets can make the ASC treatment more effective,45 and we can also add some cell factors into the sheet46 to increase the treatment effect.
Studies have shown that injection can preserve cells in dermis and the surrounding skin, and engrafted into the newly formed tissue and closely associated with other cells in the wound.47 In addition, there have been examples of using stem cell injections to treat wounds in clinical practice.48,49
In group D, the healing rates of d-ASC, ANC and control wounds were not significantly different. This finding indicates that d-ASC-rich cells and ANC transplantation did not promote diabetic wound healing and that the effects of the diabetic microenvironment were considerable; merely increasing the concentration of wound-repair cells did not change the healing process. On day three and day 12, the ASC-rich cell transplanted wounds healed faster, indicating that ASC-rich cells could promote diabetic wound healing. In diabetic patients, ASC-rich cells have been affected by high glucose, advanced glycation end-products (AGEs), and other microenvironmental factors, and their physiological and biochemical characteristics are significantly different from those of normal ASC-rich cells.23 D-ASC-rich cell transplantation may not promote diabetic wound healing, whereas normal ASC-rich cell transplantation, as a type of allograft transplantation, may face ethical problems, and immunological rejection may be unavoidable.
The healing rates of the four types of wounds were different in group N. The ASC wound healed the fastest, indicating that ASC-rich cells may promote wound healing. The healing rate of d-ASC wounds was slower than that of ASC wounds, but it was still faster than those of ANC and control wounds. The results showed that d-ASC-rich cells could still function to promote the healing of normal wounds, but their promoting effect was weakened by their former diabetic microenvironment. ANC transplantation promoted wound healing at specific time points (days three and seven), which indicates that similar cell transplantation might be an effective clinical treatment for wound healing.
The length, weight and total body surface area of the diabetic rats were different than for the sibling normal rats; therefore, we only compared the wound-healing rates among different types of wounds in one group and not between group N and group D.
Skin histological changes in diabetic rats
The subcutaneous fat layer is considered to be a source of ASCs. Under the fat layer, there is a layer of smooth muscle termed the dartos fascia, which has a specific structure in rodent animal skin.50 The dartos fascia can contract the skin and reduce wounding after injury. A layer of loose connective tissue under the dartos fascia connects the skin and the deep fascia. This connective tissue layer is another source of ASCs.51 This layer in the rat inguinal position becomes thick and develops into the inguinal fat pad. The skin of diabetic rats became thinner, the subcutaneous fat layer was lost, the loose connective tissue layer under the dartos fascia became thin, and the number of ASCs in the skin decreased. These changes in the skin structure may cause diabetic skin to be more easily damaged and more difficult to heal.
After 24 hours, three days and seven days, the granulation tissue of ASC and d-ASC wounds was thicker than that of ANC and control wounds in both group N and group D, indicating that ASC-rich cells and d-ASC-rich cells may promote granulation and accelerate the wound-healing process. The regeneration of epithelial and basal loose connective tissue in the ASC and d-ASC wounds was more obvious than in the other wounds on day 12; this difference may have been caused by ASC differentiation52 or paracrine signalling.53
Conclusions
The results presented here show that ASC-rich cells can promote wound healing in diabetic and normal rats. In the adipose tissue of diabetic rats, the density of ASCs was lower than in normal rats. D-ASC-rich cells could not promote wound healing in diabetic rats, suggesting that caution is warranted in the clinical use of diabetic adipose stem cell transplantation for the treatment of diabetic wounds. ANC transplantation may have also promoted wound healing, but the therapeutic effect was less than that for ASC-rich cell transplantation.
Reflective questions
- How can adipose stem cells (ASC) migrate to the wound and differentiate into the repair cells and its molecular biological mechanism?
- How might one increase the treatment effect of diabetic (d)-ASCs and how could one change the microenviroment of the diabetic hard-to-heal wound?
- How might one establish a quantitative and standard pathway of ASC transplation treatment?