Arch Dermatol Res (2014) 306:809–821 DOI 10.1007/s00403-014-1494-2

ORIGINAL PAPER

Expression and vitamin D-mediated regulation of matrix metalloproteinases (MMPs) and tissue inhibitors of metalloproteinases (TIMPs) in healthy skin and in diabetic foot ulcers Nallely Lo´pez-Lo´pez • Irma Gonza´lez-Curiel • Marcela Beatriz Trevin˜o-Santa Cruz Bruno Rivas-Santiago • Valentı´n Trujillo-Paez • Jose´ Antonio Enciso-Moreno • Carmen J Serrano

•

Received: 5 February 2014 / Revised: 12 June 2014 / Accepted: 20 August 2014 / Published online: 29 August 2014 Ó Springer-Verlag Berlin Heidelberg 2014

Abstract Diabetic foot ulcers (DFUs) are chronic wounds with high matrix metalloproteinase (MMP) activity, and are a frequent complication on diabetics. This work studied the expression of selected MMP and tissue inhibitor of metalloproteinases (TIMP) gene family members in DFU and normal skin biopsies, and in vitamin D-treated keratinocytes cultured from those biopsies. We report for the first time the expression of some of these genes in healthy skin. Our results suggest that vitamin D may modulate the expression of some MMP gene family members in keratinocytes. Gene expression in DFU and in non-diabetic healthy skin (control) biopsies was evaluated by RT-qPCR for MMP-1, MMP-3, MMP-8, MMP-9, MMP-10, MMP-19, TIMP-1 and TIMP-2, and also by immunohistochemistry for MMP-1 and MMP-9. Primary keratinocytes cultured from DFU and healthy skin biopsies were used for gene expression analyses of selected MMPs and TIMPs by RT-qPCR, both in the presence and absence of calcitriol. The expression of MMP-1, MMP-8, MMP-9, MMP-10, and TIMP-2 in healthy skin is reported here for the first time. DFUs showed increased MMP-1, MMP-9 and TIMP-1 expression, compared to healthy skin. Calcitriol down-regulated MMP-1 and MMP-10 expression in DFUN. Lo´pez-Lo´pez  I. Gonza´lez-Curiel  B. Rivas-Santiago  V. Trujillo-Paez  J. A. Enciso-Moreno  C. J. Serrano (&) Medical Research Unit Zacatecas, Mexican Institute of Social Security-IMSS, Zacatecas, Mexico e-mail: [email protected] N. Lo´pez-Lo´pez  I. Gonza´lez-Curiel  V. Trujillo-Paez Department of Immunology, Faculty of Medicine, Autonomous University of San Luis Potosı´, San Luis Potosı´, Mexico M. B. Trevin˜o-Santa Cruz Science Department, Edison State College, Fort Myers, Florida, USA

derived keratinocytes but not in those derived from healthy skin. Our data demonstrate the expression of certain MMPs that had not been previously described in healthy skin, and further support previous reports of MMP and TIMP upregulation in DFUs. Our results point to calcitriol as a potential modulator for the expression of certain MMP members in DFUs. Keywords Type 2 diabetes  Non-insulin dependent diabetes mellitus (NIDDM)  Matrix metalloproteinases (MMPs)  Tissue inhibitors of metalloproteinases (TIMPs)  Diabetic foot ulcers (DFUs)  Keratinocytes  Vitamin D  Calcitriol

Introduction Diabetes is a chronic disease affecting 347 million people worldwide as of 2013; type 2 diabetes (also called noninsulin dependent diabetes mellitus, or NIDDM) accounts for 90 % of those cases [70]. One of the most frequent complications in NIDDM is the formation of diabetic foot ulcers (DFUs), a problem that affects the patients’ quality of life and the economy of many countries worldwide [34, 70]. A DFU is a sort of chronic wound characterized by delayed or absent closure. There are several associated factors contributing to the permanence of these wounds, including local inflammation; decreased production of growth factors; and uncontrolled proteolysis, which brings about an imbalance between the accumulation of extracellular matrix (ECM) proteins, and their degradation by proteases such as matrix metalloproteinases (MMPs) [4, 63].

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MMPs are a family of extracellular proteases produced by various cell types such as fibroblasts, keratinocytes, endothelial cells, neutrophils, macrophages and eosinophils. A total of 24 related, but distinct, MMPs have thus far been described in vertebrates, 23 of which are known to be expressed in humans [43]. The different MMPs are either anchored to the plasma membrane or secreted from the cell [43]. On the basis of main substrate specificity, sequence similarity, and domain organization, vertebrate MMPs can be divided into six groups: collagenases, gelatinases, stromelysins, matrylisins, membrane-type, and others with less well-defined specificity [43]. In vitro enzyme studies have shown considerable overlap in the substrates that the different MMPs can cleave, particularly among the ECM substrates [57]. For example, fibronectin, laminins, elastin and type IV collagen can be degraded by various MMPs in vitro. In a setting such as inflammation, in which essentially all MMPs are present, the shared substrate potential would seemingly provide biochemical redundancy. However, substrate selectivity may be honed by two processes: compartmentalization and enzyme–substrate affinity. Kinetic studies using model substrates have shown that specific MMPs degrade some substrates more efficiently than others [18, 36]. Regulation of MMP activity is very tight, including control points at the level of transcriptional regulation, and of inhibition of the active enzyme mediated by the specific binding to tissue inhibitors of metalloproteinases (TIMPs) [35]. Four different TIMPs (TIMP-1 through TIMP-4) have thus far been described in mammals, and all MMPs are inhibited by all TIMPs, albeit to variable extent [43]. In normal tissue, MMPs and TIMPs are generally believed to be expressed at very low levels, if at all, but their expression and activation can be rapidly induced when active tissue remodeling is needed [38]. Constitutive expression in intact skin has thus far only been documented for MMP-3, MMP-7, MMP-19 and MMP-28 [51, 53, 54], as well as for TIMP-1 [23]. Despite extensive studies, the molecular processes underlying wound repair are still not fully understood [50]. Knowledge of such processes is a crucial step in the development of improved treatments for chronic wounds. Several research groups have shown that chronic wound fluid (CWF) has an increased proteolytic activity compared to acute wound fluid (AWF) [29, 31, 60], a condition largely attributed to the presence and activity of MMPs [68]. Wound repair proteome studies have shown protein expression differences in poor healing, chronic wounds (including DFUs), when compared to normally healing wounds; most of the involved proteins are related to inflammation, angiogenesis, and cell mortality, with MMP1, MMP-2 and MMP-8 being among the up-regulated proteins [10, 28, 32, 37, 42, 49]. Additional evidence

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suggests that while MMP activity is necessary during the normal healing process, some MMPs may have a deleterious effect when present in excessive amounts. For example, MMP-9 knock-out mice display delayed wound closure, and keratinocyte migration was reported to be dependent on this MMP in in vitro wound assays [20]. But on the other hand, studies of DFU exudates showed that an excess of MMP-9 was deleterious to wound healing [42], while a high MMP-9/TIMP-1 ratio was associated with poor ulcer healing [32]. Another MMP that has been associated with proper wound healing is MMP-1; in a prospective study, a high MMP-1/TIMP-1 ratio in DFU exudates was as predictor of wound healing [42]. Besides MMP-9 and MMP-1, there are other MMPs whose expression is induced as a response to wounding; among them are MMP-3 and MMP-10, which participate in epidermal wound healing [38]. MMP-3 is known to have a major role in regulating wound contraction [5]; it can activate several pro-MMPs, and it increases the bioavailability of many cytokines [65, 67]. MMP-19 is detected in proliferating epithelium during cutaneous wound healing in humans, and its overexpression in a keratinocyte cell line promotes cell proliferation and migration [21, 54]. Lastly, MMP-8 is also known to promote cutaneous wound healing in mice [16, 19]. On the other hand, TIMPs can regulate cell migration in wound healing by modulating the activity of specific MMPs. TIMP-1 has been detected in epithelial cells of healing excisional and burn wounds in humans. Moreover, TIMP-1 is also present in wound fibroblasts, especially around blood vessels [64]. Exogenous addition of TIMP-2 impairs cell migration in vitro, possibly by inhibiting MMP-14’s cleavage of syndecan-1 from the cell surface [11]. Additional studies suggest that TIMP-2 increases migration of human epidermal keratinocytes in vitro and favors mice wound healing in vivo [61]. TIMP-3 appears to be involved in the ECM remodeling phase, as TIMP-3 null mice have abnormal collagen and fibronectin remodeling [14]. Expression of TIMP-4 has not been detected in human acute wounds [64]. Efforts to describe the expression of MMPs and TIMPs in healthy skin and in DFUs have been far from exhaustive [41]. While the presence of MMP-1 and MMP-9 in DFUs is now clear, there are no reports describing their expression in healthy, intact skin. Moreover, the expression patterns of MMP-10, MMP-19 have not been studied in either healthy skin or DFUs. A more complete description of the spatiotemporal expression patterns of MMPs and TIMPs, both in DFUs and in intact skin, should help us better understand the molecular events occurring in chronic wounds, as well as those associated with healing. Calcitriol is the active form of vitamin D, and it is produced in humans by a cascade of reactions beginning

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with the photochemical synthesis of vitamin D3 in the skin. The epidermis is unique among calcitriol-producing organs in that it contains the full enzymatic complex needed for calcitriol generation, as well as its nuclear receptor, VDR (vitamin D receptor). In skin, vitamin D3 has several wellrecognized biological activities, including modulation of the proliferation and differentiation of keratinocytes [3, 13], as well as suppression of MMP-9 production in HaCaT cells [1] and of MMP-2, MMP-3, TIMP-1, and TIMP-2 production in vitro in cholesteatoma keratinocytes [26]. The present work aimed to describe the expression of selected MMP and TIMP gene family members, including some whose expression in healthy skin and/or in DFUs had not been previously studied. Given the need for therapeutic agents to treat diabetic foot, this paper also explored the effect of exogenous calcitriol on the expression of a group of MMP and TIMP genes in primary keratinocyte cultures from DFU and healthy skin biopsies.

Table 1 Gene-specific oligonucleotide sequences used for RT-qPCR assays

Materials and methods

RNA extraction and RT-qPCR assays

Subjects

Biopsy sections placed in RNAlater were homogenized with an Ultra-Turrex disperser (Ika, Wolmington, NC, USA); RNA was isolated using RNeasy fibrous tissue kit (Qiagen, Du¨sseldorf, Germany) and then quantified using a spectrophotometer ND-1000 (Nanodrop, USA). Reverse transcription of mRNA was performed using 5 lg of total RNA, 2 lM oligodT (Invitrogen, Carslbard, CA), 10 units of RNase inhibitor (10 U/lL) (Invitrogen, Carslbard, CA), 0.5 mM of each dNTP, and 4 U of Omniscript Reverse Transcriptase (Qiagen, Inc, Mexico) in a total volume of 20 lL. cDNA was spectrophotometrically quantified using an ND-1000 (Nanodrop, USA). A LightCycler 2.0 thermocycler (Roche, Germany) was used for qPCR assays, using the LightCycler Taqman Master Mix (Roche, Germany); 100 ng of each cDNA sample were used in each qPCR. Gene-specific oligonucleotide pairs for qPCR experiments were designed for six MMP, two TIMP genes, and for the hypoxanthine–guanine phosphoribosyltransferase (HPRT) gene using Roche’s online Universal Probe Library Assay Design Center; the gene-specific primers are shown in Table 1; they were synthetized by Invitrogen, USA. The specific probes (Roche, Germany) used for each oligonucleotide pair were also designed with the Roche freeware. Relative gene expression in each sample was calculated with the 2-DDCt method, with HPRT expression used as the internal reference [33].

The study was approved by the National Ethics Committee and the National Commission for Scientific Research of the Mexican Social Security Institute (project numbers R-2008-785-066 and R-2013-785-019). All participants signed a written informed consent. None of the subjects included in the study had any systemic or autoimmune diseases, nor were they being treated with steroids or hormonal therapy. All patients were HIV negative. For the experimental group, eleven type 2 diabetes patients with diabetic foot ulcer (DFU) grade II or III according to Wagner´s scale [22] were recruited at the Hospital General de Zona (HGZ) #1 IMSS (Zacatecas, Mexico). In all patients, foot ulcers were debrided and thoroughly washed, and this was followed by systemic antibiotic therapy for 7 days prior to foot biopsy. Control biopsies were taken from eight non-diabetic donors during an orthopedic surgery. Biopsies of diabetic foot ulcers were obtained from patients receiving angiology services and were selected under the following inclusion criteria: diagnosis of type 2 diabetes, fasting glucose plasma levels [100 mg/dL, postprandial glucose [200 mg/dL, as reported in their current medical records. Exclusion criteria were as follows: individuals with diabetes mellitus type 1 (DM1), treated with immunosuppressant steroidal anti-inflammatory drugs, or not signing informed consent. All skin samples were obtained as a triangular biopsy measuring 1 cm per side. Each biopsy was divided into three sections; one of them was stored in RNAlater

Gene

Forward primer (50 –30 )

Reverse primer (50 –30 )

MMP-1

gctaaaccctgaaggtgat

ggcgtgtaattttcaatcctg

MMP-3

caaaacatatttctttgtagaggacaa

ttcagctatttgcttgggaaa

MMP-8

tctttgtaaatgaccaattctgga

ggaaaggcacctgatatgct

MMP-9

gaaccaatctcaccgacagg

gccacccgagtgtaaccata

MMP-10

tggccctctcttccatcata

tgatggcccagaactcattt

MMP-19

gaagatatcaccgaggctctga

gatcctctaggccacaacga

TIMP-1 TIMP-2

ctgttgttgctgtggctgat tggaggaaagaaggaatatctca

aacttggccctgatgacg ttctgggtggtgctcagg

HPRT

tgaccttgatttattttgcatacc

cgagcaagacgttcagtcct

(Qiagen, Du¨sseldorf, Germany) for RT-qPCR assays, the second one was embedded in paraffin for immunohistochemistry assays, and the third one was used for keratinocyte primary culture.

Primary epidermal keratinocyte cultures from biopsies Biopsy sections were mechanically disaggregated with a scalpel to obtain explants, which were then seeded into

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25-mL cell culture bottle flasks (Corning flask 30–56, NY, USA) filled with a 1:1 mixture of DMEM and HAM-F12 media (Safc Biosciences, Kansas, USA). This mixture results in a medium with lowered Ca2? concentration (1.05 mM), recommended for epidermal primary keratinocytes, and it was supplemented with 10 % fetal bovine serum (FBS), penicillin and streptomycin 1 % each (all from Gibco BRL, CA, USA). Explants were incubated at 37 °C in a 5 % CO2 atmosphere until adherence. Once adhered, the cells were cultured in a 1:1 mixture of DMEM and HAM-F12 media, supplemented with the full content of one keratinocyte growth kit PCS-200-040 (ATCC). The final concentration for each component in the culture medium is as follows: bovine pituitary extract (BPE) 0.4 %, recombinant human transforming growth factoralpha (rh TGF-alpha) 0.5 ng/mL, L-glutamine 6 mM, hydrocortisone 100 ng/mL, insulin 5 lg/mL, epinephrine 1.0 lM, apo-transferrin 5 lg/mL. The keratinocyte growth medium is formulated to inhibit fibroblast growth, and the low calcium concentration (0.06 mM) slows down differentiation. Hydrocortisone prevents fibroblast proliferation by controlling the synthesis of several growth factors required for it [9]. Once cell cultures reached confluence, they were immediately used for vitamin D induction experiments. Bacteriological controls were performed by culturing samples on blood agar for 24 h at 37 °C (MCDLAB, Mexico). Two approaches were employed to confirm that primary epidermal cells were keratinocytes and not fibroblasts; both are based on detection of cytokeratin-5 (KRT5), a protein specifically expressed in keratinocytes from the basal layer [56], and were carried out as follows. (1) The percentage of keratinocytes in cell cultures was determined by fluorescence-activated cell sorting (FACS) analysis using antiKRT5 antibody (cytokeratin pam antibody, Thermofisher Scientific, USA), obtaining 95 % purity (n = 2). Viability was checked by the Guava Viacount Assay (Millipore, MA, USA) showing 95 % of viability (n = 2) (results not shown). (2) KRT5 was detected by western blot (WB) as follows: cell lysates were prepared by homogenizing the cells in RIPA buffer (NaCl 150 mM, Tris–HCl 50 mM pH 7.4, EDTA 1 mM, Triton X-100 1 %, SDS 0.1 %), supplemented with phenylmethanesulphonylfluoride (PMSF) 1 mM (Sigma-Aldrich, USA) and 5 % v/v protease inhibitor (complete protease inhibitor cocktail, previously dissolved in RIPA buffer; Roche Diagnostics, Germany); equal amounts of total protein, as quantified by the bicinchoninic acid method (n = 2) [55], were loaded for WB analysis with an anti-rabbit polyclonal anti-KRT5 (Santa Cruz Biotechnology, Inc., CA, USA); b-actin was used as an expression control with anti-b-actin mouse ascites fluid (clone AC-74, Sigma-Aldrich, USA) (results not shown).

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Vitamin D induction assay The cell concentration of keratinocytes cultured from biopsies was determined by direct microscopic count of a 10-lL culture aliquot (stained with trypan blue, final concentration 0.04 % v/v) using a Neubauer chamber (n = 2). Keratinocytes were then seeded into 24-well plates (Corning Incorporated, USA, COSTAR Number 3524) at 2 9 105 cells/well, suspended in 1 mL DMEM:HAM-F12 supplemented with 1 % (v/v) FBS, and incubated in a 5 % CO2 atmosphere at 37 °C for 24 h. For Vitamin D induction assays, cells were incubated under the same conditions for an additional 18 h, after addition of either 1 mL of culture medium (untreated control), or 1 mL of culture medium containing 10 ll of a 1 9 10-4 mM calcitriol (1,25-dihydroxyvitamin D3, Roche, USA) stock solution (in absolute ethyl alcohol; stored at -20 °C) (SigmaAldrich, USA;) for a final calcitriol concentration of 1 nM—i.e., the concentration previously reported by Bahar-Shany et al. [1]. Immunohistochemistry Biopsies were fixed by immersion (for at least 24 h and a maximum of 2 months) in buffered formaldehyde (formaldehyde (CH2O) 1.3 M, monobasic sodium phosphate (NaH2PO4) 0.02 M, dibasic sodium phosphate (Na2HPO4) 0.045 M, pH 7.0–7.5) and then embedded in paraplast X-tra tissue embedding medium (McCormick Scientific, USA). 4-lm thick sections were cut using a Slee Mainz Cut 5062 microtome (Orgontec Company, Germany), and mounted on KlingOnSlides (Biocare Medical, CA, USA). Sections were deparaffinized by heating the slide at 58 °C for 18 h; endogenous peroxide was then quenched with 250 lL hydrogen peroxide/methanol 3 % (v/v) for 45 min at room temperature, and non-specific protein binding sites were blocked through 12 washes (using a squirt bottle) with HNC buffer (Hepes 0.02 M, NaCl 0.3 M, CaCl2 0.00529 M) supplemented with 7 % pooled human serum. Polyclonal primary antibodies for either MMP-1 or MMP-9 (both from Santa Cruz Biotechnology, CA, USA) were used at 1:20 and 1:30 dilutions, respectively, applying 70 lL on each slide, and incubating at room temperature for 24 h. Negative controls were incubated with HNC buffer supplemented with 2 % pooled human serum without the primary antibody, under the same conditions used for the primary antibodies. Slides were then washed 12 times (using a squirt bottle) with a 1:1 mixture of HNC/ PBS at room temperature. A biotinylated goat anti-mouse antibody (Santa Cruz Biotechnology, CA, USA), 200 lL at 1:550 dilution was applied to each slide, incubating at room temperature for 1.5 h.

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Secondary antibody binding was detected using 100 lL per slide of the streptavidin–HRP system (Biocare Medical, CA, USA), developed with 200 lL per slide diaminobenzidine (DAB) for 30 s, and counterstained with hematoxylin (SEALAB, Mexico). Finally, samples were dehydrated by subsequently immersing the slides (20 times each) in 50, 70, 96 % ethyl alcohol (Sigma, USA) and finally in absolute ethyl alcohol, then 20 times each in ethyl alcohol/xylol (1:1) and in absolute xylol (Sigma, USA). Excess xylol was drained prior to mounting with 200 lL Entellan (Merck Chemicals, Chemical, Massachusetts, USA). Biopsy sections were examined with an inverted microscope (Zeiss, Axiostar, NY, USA) and a semi-quantitative analysis was performed to determine the area positive for either MMP-1 or MMP-9, using the AxionVisionRel 4.8 program (Carl Zeiss International Company, Germany). Sample analysis was carried out by measuring the sample’s total epidermis area and then the epidermal area positive for either MMP-1 or MMP-9, to calculate the percent area positive for either MMP-1 or MMP-9. Statistics analysis To compare the proportion of individuals expressing a gene, a Fischer test was used. Differences were considered statistically significant at p \ 0.05. Relative gene expression values were analyzed using a Grubb’s test to detect outliers, using Quickcals freeware by GraphPad, USA. Then, to compare relative mRNA levels of each gene between groups, the mean, standard deviation (SD) or median and interquartile range (IQR) values were calculated and compared using the Student t test and the Mann–Whitney U test for parametric and non-parametric data, respectively. Differences were considered statistically significant at p \ 0.05. All calculations were made using GraphPad Prism software v. 5.0 (San Diego, CA, USA).

Results Patient data Eleven patients with DFUs (9 male, 2 female; mean age 57 ± 11 years, age range 44–72 years) were recruited for the experimental group. The reported mean blood glucose level in this group was 233 ± 116 mg/dL; their biopsied DFUs were either grade II or grade III (Wagner´s scale). The control group consisted of nine healthy, non-diabetic individuals (3 male, 6 female; mean age 59 ± 18 years, age range 25–84 years) undergoing orthopedic surgery, during which skin biopsies were taken; the reported mean blood glucose level in this group was 94.26 ± 14.75 mg/ dL. Statistically significant differences (p \ 0.05) between

813 Table 2 Percentage of individuals in the control (healthy skin biopsies) and experimental groups (DFU biopsies) expressing MMPs and TIMPs Gene

Percentage of individuals in the control group expressing the genea (healthy skin biopsies)

Percentage of individuals in the experimental group expressing the genea (DFU biopsies)

Fisher’s exact test p \ 0.05

MMP-1

75

91.6

0.5368

MMP-3 MMP-8

100 50

100 58.33

NA 1.0

MMP-9

62.5

100

0.049*

MMP-10

50

75

0.356

MMP-19

100

91.6

1.0

TIMP-1

100

91.6

1.0

TIMP-2

62.5

83.3

0.347

a

A positive expression subject was considered when an RT-qPCR using 100 ng cDNA (from the reverse transcription of 5 lg total RNA) showed a Ct value \40. The percentage of individuals expressing each gene in the control and experimental groups was compared using Fisher’s exact test (p \ 0.05). NA Not applicable, i.e. no percentage difference between groups. GraphPad InStat software, USA

the experimental and the control groups were only seen in their mean blood glucose levels. Several MMPs and TIMPs are expressed in healthy human skin and in DFUs Thus far, the expression of MMP-1, MMP-3, MMP-8, MMP-9, MMP-10, MMP-19, TIMP-1, and TIMP-2 has been associated with various stages of wound healing; however, the expression of most of these genes had not been explored in healthy skin with MMP-3, MMP-19 and TIMP-1 being the exceptions [23, 51]. We thus set out to explore the presence of transcripts from these genes both in intact, normal skin, and in DFUs. To this end, RT-qPCR assays were performed for all biopsies taken from both the control group and the experimental group subjects. Transcripts for all tested genes were found in both healthy skin and DFUs biopsies, although with varying percentages of individuals within each group expressing each of the specific genes (Table 2). The percentage of subjects expressing each individual gene in the control and experimental groups was compared. Only in the case of MMP-9 was there a significant difference between the groups, with the percentage of subjects expressing this gene being greater in the experimental group (p \ 0.05; Table 2); the rest of the genes appear to be constitutively expressed both in DFUs

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Fig. 1 Relative transcription levels of MMP-1 (a), MMP-3 (b), MMP-8 (c), MMP-9 (d), MMP-10 (e), and MMP-19 (f) in DFUs compared to healthy skin, calculated from RT-qPCR experiments. For each gene, mRNA levels were first normalized using HPRT expression as the internal reference, and the relative mRNA levels (fold change) in DFUs with respect to those in healthy skin were then calculated according to the 2-DDCt method. Data are presented as median and interquartile range. A Grubbs’ test was run for each biopsy group to detect outliers, which were not included on the performed comparisons. Comparisons between the control and experimental biopsy groups were performed using the paired Student t test (for MMP-10) or the Mann–Whitney U test (for all other MMPs). HS = healthy skin biopsies (n = 8), DFU diabetic foot ulcer biopsies (n = 11 for MMP-1, MMP-9, and MMP-19; n = 12 for MMP-3; n = 7 for MMP-8; n = 10 for MMP-10). *p \ 0.05, **p \ 0.01, ***p\ 0.001. All statistical tests were performed with GraphPad Prism 5 software, USA

and in healthy skin, as at least 50 % of the individuals tested in each group expressed those genes. mRNA levels of MMP-1, MMP-9 and TIMP-1 are upregulated while those of MMP-8 and MMP-10 are down-regulated in DFUs Once basal transcription of the selected MMPs and TIMPs was confirmed in healthy skin biopsies (control group), their mRNA levels were used as a reference to compare their expression in DFU biopsies (experimental group). Figures 1 (MMPs) and 2 (TIMPs) show the gene expression analyses derived from the RT-qPCR experiments, indicating the relative mRNA levels (fold change) in DFUs compared to the basal levels measured in healthy skin. Given that not all genes were detected in all biopsies, the number of samples included in the analysis of each gene varies accordingly. Figures 1 and 2 show that expression of

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MMP-1, MMP-9, and TIMP-1 is up-regulated in DFU biopsies, with significant differences of p \ 0.01, p \ 0.05 and p \ 0.01, respectively. Furthermore, expression of MMP-8 and MMP-10 was down-regulated in DFUs, with significant differences of p\ 0.05 and p \ 0.001, respectively. However, no significant expression differences were seen between healthy skin and DFU biopsies for MMP-3, MMP-19, and TIMP-2. Semi-quantitative immunohistochemical analyses reveal up-regulation of MMP-9, but not of MMP-1 in DFU biopsies Out of the six MMP genes we tested, only MMP-1 and MMP-9 were up-regulated in DFUs as measured by RTqPCR (Fig. 1). To determine if such increases in transcript accumulation actually result in increased protein levels, we investigated by immunohistochemistry the presence of

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Fig. 2 Relative transcription levels of TIMP-1 (a) and TIMP-2 (b) in DFUs compared to healthy skin, calculated from RT-qPCR experiments. For each gene, mRNA levels were first normalized using HPRT expression as the internal reference, and the relative mRNA levels (fold change) in DFUs with respect to those in healthy skin were calculated according to the 2-DDCt method. Data are presented as median and interquartile range. A Grubbs’ test was run for each

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biopsy group to detect outliers, which were not included on the performed comparisons. Comparisons between the control and experimental biopsy groups were performed using the Mann–Whitney U test. HS healthy skin biopsies (n = 8), DFU diabetic foot ulcer biopsies (n = 11). *p \ 0.05, **p\ 0.01, ***p \ 0.001. All statistical tests were performed with GraphPad Prism 5 software, USA

Fig. 3 Immunohistochemistry of MMP-1 and MMP-9 in healthy skin and DFU biopsies. Representative images for MMP-1 immunodetection in healthy skin (a) and DFU (b) biopsies, and for MMP-9 immunodetection in healthy skin (c) and DFU (d) biopsies. Sections contain dermis and epidermis tissues; they were stained using an IHC method that produces a brown stain, and then counterstained with hematoxylin. All cells positive for the corresponding MMP (brown immunostain) are epidermal keratinocytes. Size bar is valid for all micrographs shown. Micrographs were taken at 2009 magnification with an inverted microscope

MMP-1 and MMP-9 in DFU and healthy skin biopsies containing dermis and epidermis. Both MMPs localized solely to the epidermal keratinocytes from DFU and

healthy skin biopsies; representative micrographs are shown in Fig. 3a, b (MMP-1), c and d (MMP-9). According to the semi-quantitative data provided by this method, a

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Fig. 4 Semi-quantitative analysis of immunohistochemical data for MMP-1 and MMP-9 in healthy skin and DFU biopsies. The percent of MMP-1 (a) and MMP-9 (b) immbiopsies exhibited a significantunostained area within the epidermis was calculated for each healthy skin (HS) and DFU biopsy section analyzed. Values shown are mean and

standard deviation. Statistical analysis was performed using the paired Student t test with GraphPad Prism 5 software, USA. n = 10 (HS) and n = 8 (DFU) for MMP-1; n = 11 (HS and DFU) for MMP-9. *p \ 0.05, **p \ 0.01, ***p\ 0.001

significant increase of MMP-9 was detected in epidermal keratinocytes of DFU biopsies, compared to those of healthy skin (p \ 0.01; Fig. 4b). No significant differences in MMP-1 levels were observed between the epidermal keratinocytes of DFU and healthy skin biopsies (Fig. 4a).

(Fig. 6a). Similar results were observed in keratinocytes derived from DFUs for MMP-3 (Fig. 5d) and for TIMP-1 and TIMP-2 (Fig. 6b, d). However, keratinocytes cultured from DFU biopsies exhibited a significant reduction (p \ 0.05) in MMP-1 and MMP-10 mRNA levels when treated with vitamin D, compared to the untreated control (Fig. 5b, e).

Exogenous calcitriol (1,25-dihydroxyvitamin D3) application down-regulates transcription of MMP-1 and MMP-10 in keratinocytes cultured from DFU biopsies Exogenous application of the active form of vitamin D (calcitriol) is known to down-regulate the transcription and activity of MMP-9 in the HaCaT keratinocyte cell line, with a maximum effect at 1 nM [1]. We thus evaluated the effect of this calcitriol dosage on the transcription of selected MMPs and TIMPs, 18 h after its addition to primary keratinocytes cultured from either healthy skin or DFU biopsies. Initially, all MMPs and TIMPs previously assayed by RT-qPCR in skin biopsies were also surveyed by RTqPCR in these cultures. Transcripts for nearly all tested genes were detected in all cultures, with the exceptions being MMP-8 and MMP-9, which were not found in any of them, and MMP-19, which was only detected in 4 out of 9 healthy skin-derived cultures, and in 1 out of 11 DFUderived cultures (data not shown). MMP-8, MMP-9 and MMP-19 were thus not included in the vitamin D induction assays. Figures 5 (MMP-1, MMP-2 and MMP-3) and 6 (TIMP-1 and TIMP-2) show the relative transcript levels present in vitamin D-treated keratinocytes, calculated with respect to those measured in untreated (control) keratinocytes, both for healthy skin- and DFU-derived cultures. In keratinocytes cultured from healthy skin, mRNA levels did not change significantly in response to exogenous vitamin D for any of the tested MMPs (Fig. 5a, c, e) and TIMPs

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Discussion It has been proposed that basal levels of MMPs and TIMPs maintain the skin´s homeostasis, and that they may be part of a quick response to infections and mild lesions, since MMP and TIMP modulate keratinocyte migration to the wounded site [38]. However, only the expression of a few MMPs has thus far been reported in normal tissues. These include MMP-3, whose presence in healthy skin biopsies has been reported [53]; MMP-7 and MMP-19, which are constitutively expressed in sebaceous glands of healthy skin [51, 54]; MMP-28 and MMP-19, which are expressed in muscle´s basal keratinocytes and in endothelial cells of veins and arteries [54]; and TIMP-1 in aged human skin [23]. Our results show that also MMP-1, MMP-8, MMP-9, MMP-10, and TIMP-2 are expressed in healthy intact skin. To our knowledge, this is the first time that the expression of MMP-1, MMP-8, MMP-9, MMP-10, and TIMP-2 is reported in healthy skin. Our results show that MMP-1 and MMP-9 are up-regulated in DFUs, which is consistent with related literature reports. In the case of MMP-1, while its expression is induced as a rapid response to wounding [52], it must decrease to allow reepithelialization [58], and it is, therefore, not surprising that a high level of MMP-1 seems to be essential for wound healing in DFUs [42]. On the other hand, high MMP-

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Fig. 5 Effect of exogenous calcitriol on mRNA levels of selected MMPs in keratinocytes cultured from healthy skin and DFU biopsies. The relative transcript levels (fold change) of MMP-1 (panel A and B), MMP3 (panel C and D) and MMP-10 (panel E and F) in response to 1 nM calcitriol application (?Vit. D) were calculated with respect to the untreated control (U) according to the 2-DDCt method, both in keratinocytes cultured from healthy skin (HSK, panel A, C, and E) or from DFUs (DFUK, panel B, D, and F). HPRT expression was used as internal reference. A Grubbs’ test was run for each biopsy group to detect outliers, which were not included on the performed comparisons. Comparisons between groups were performed using the Mann–Whitney U test. HSK U samples: n = 9 for all MMPs.HSK ? Vit. D samples: n = 9 for MMP-3; n = 7 for MMP-1 and MMP-10; DFUK U samples: n = 9 for all MMPs. DFUK ? Vit. D samples: n = 8 for MMP-3; n = 6 for MMP-1 and MMP-10. *p \ 0.05, **p\ 0.01, ***p \ 0.001. All statistical tests were performed with GraphPad Prism 5 software, USA

9 levels in DFU exudates have been associated with poor healing [32]. Additionally, current evidence suggests that diabetics have a MMP and TIMP imbalance compared to non-diabetics. For example, higher active levels of MMP-1 and MMP-9, and lower TIMP-1 levels were seen in human skin organ cultures derived from intact skin of diabetics, compared to non-diabetics [71]. In fact, diabetic patients whose ulcers failed to heal had higher MMP-9 expression in biopsies and higher MMP-9 serum levels when compared with those whose ulcers healed [8]. The same study concluded that while neuropathy and vascular factors are associated with the development of DFUs, the main factors associated with a failure to heal these ulcers include preexisting increased serum levels of MMP-9, inflammatory cytokines, and various growth factors.

While we observed that MMP-1 transcription was upregulated in DFUs compared to healthy skin, such effect was not detected when MMP-1 protein levels were compared by immunohistochemistry (IHC). There may be various reasons behind this discrepancy, one of them being the detection method itself, since IHC is a semi-quantitative method, less sensitive than qPCR. Another explanation may be related to the fact that MMP-1 levels decrease when wound closure is being reached [58] and, given the heterogeneity of our samples and the associated variability in their damage/regeneration stages, there may be varying translational regulatory processes at play, causing the average protein levels to be less meaningful. For example, studies of unfolded protein response (UPR) on cell lines showed MMP-9 was among the translationally repressed

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Fig. 6 Effect of exogenous calcitriol on mRNA levels of selected TIMPs in keratinocytes cultured from healthy skin and DFU biopsies. The relative transcript levels (fold change) of TIMP-1 (panel A and B) and TIMP-2 (panel C and D) in response to 1 nM calcitriol application (?Vit. D) were calculated with respect to the untreated control (U) according to the 2-DDCt method, both in keratinocytes cultured from healthy skin (HSK, panel A and C) or from DFUs (DFUK, panel B and D). HPRT expression was used as internal

reference. A Grubbs’ test was run for each biopsy group to detect outliers, which were not included on the performed comparisons. Comparisons between groups were performed using the Mann– Whitney U test. HSK U samples: n = 9 for TIMP-1 and TIMP-2. HSK ?Vit. D samples: n = 6 for TIMP-1 and n = 8 for TIMP-2. DFUK U samples: n = 9. DFUK ?Vit. D samples: n = 8 for TIMP-1 and n = 7 for TIMP-2. *p\ 0.05. All statistical tests were performed with GraphPad Prism 5 software, USA

proteins [48], which may be relevant in our case as UPR has been linked to diabetes and its associated neuropathy [30, 44]. In the present work, transcription of MMP-8 and MMP10 was found to be down-regulated in DFUs compared to healthy skin. Such down-regulation of MMP-8 is in agreement with a previous study using a diabetic mice wound model, in which the specific chemical inhibition of MMP-8 caused delayed wound healing by reducing reepithelialization [15]. Similarly, previous evidence points to a role for MMP-10 in wound healing, as its expression is induced in keratinocytes at the wound’s leading edge during epidermal healing in both human and mouse wounds [53]. Moreover, a tightly regulated expression level of MMP-10 is a pre-requisite for controlled matrix degradation at the wound site, thereby controlling keratinocyte migration [27]. Therefore, a decreased expression of MMP-8 and MMP-10 in DFUs may be contributing to the ulcers’ chronicity.

TIMP-1 has been detected in epithelial cells of healing excisional wounds and burn wounds in humans. Additionally, TIMP-1 transcripts were present in wound fibroblasts, especially around blood vessels [64]. We found significantly higher TIMP-1 mRNA levels in DFUs compared to healthy skin. Given that a high MMP-1/TIMP-1 ratio was associated to wound healing in DFUs [42], it is conceivable that elevated TIMP-1 levels may be delaying wound healing in the analyzed DFU biopsies; however, this would need to be confirmed through measurements of actual MMP-1 and TIMP-1 protein levels. Although we did not evaluate TIMP-3 expression, it is important to mention that it appears to play a homeostatic role in the skin’s health, and that a decrease in its levels has been associated to organ complications in diabetics. Regarding the liver, activation of TNF-a-converting enzyme (TACE) is central to the pathogenesis of nonalcoholic steatohepatitis (NASH), and it is mediated, among others factors, by a down-regulation of Sirtuin1/

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TIMP-3 pathways [24]. TIMP-3 is also important for maintaining kidney homeostasis: studies on kidney biopsies from patients with diabetic nephropathy showed a reduction in TIMP-3, compared to controls [12]. Combined with data obtained through studies of Timp-3 -/knock-out mice, the authors suggested that loss of TIMP3 is a hallmark of diabetic kidney disease both in human and in mouse. Furthermore, on experimental models, even the unwounded skin of diabetic rats have decreased type 1 collagen (COL1) and TIMP-3 concentrations, compared to healthy controls [25]. Moreover, insulin or metformin treatment of diabetes type 1 and 2, respectively, in such diabetic rats, resulted in increased TIMP-3 and COL1 concentrations in unwounded skin as compared to those found in untreated diabetic rats [25]. In addition, ischemic DFUs exhibited higher MMP-9 activity and lower TIMP-3 mRNA expression, compared to neuropathic ulcers [40], suggesting that the increased proteolytic environment may represent a causative factor in the ulcer’s progression [40]. In an effort to explore the use of vitamin D as a potential therapeutic agent, we investigated calcitriol’s effect on the expression of MMPs and TIMPs in primary keratinocyte cultures derived both from DFU and healthy skin biopsies. In the absence of exogenous calcitriol, both types of cultured keratinocytes were found to express MMP-1, MMP-3, MMP-10, MMP-19, TIMP-1 and TIMP-2. The present report documents for the first time the expression of MMP19 and TIMP-2 in cultured keratinocytes from human DFU and healthy skin biopsies, although MMP-19 transcripts were not found in every single sample analyzed. On the other hand, MMP-8 and MMP-9 expression was not detected in either type of keratinocyte culture. This was expected, since MMP-8 is expressed mainly by neutrophils, and at low levels in keratinocytes and fibroblasts [38], while MMP-9 has been detected in injured epithelia [2] and in migrating keratinocytes, T cells, neutrophils and macrophages [38, 63]. The expression of MMP-1, MMP-3, MMP-10, TIMP-1 and TIMP-2 was analyzed in both types of keratinocyte cultures after exogenous calcitriol application. The only vitamin D-mediated effect observed was a decrease in MMP-1 and MMP-10 expression in DFU-derived keratinocytes. However, keratinocytes were stimulated using a single vitamin D dose (1 nM) and exposure time (18 h); and given that vitamin D modulation is known to be concentration and time dependent [17, 26], we cannot rule out that calcitriol does in fact modulate MMP-3, TIMP-1 and TIMP-2 expression. As mentioned before, MMP-1 is expressed exclusively in basal keratinocytes at the migrating epithelial edge of wounds [45, 52]; and after a rapid increase in response to

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wounding, its expression must decrease to favor reepithelialization [58]. Likewise, MMP-10 is known to be expressed at the wound edge, co-localizing with MMP-1; a tight regulation of its expression level is a pre-requisite for controlled matrix degradation at the wound site, thereby controlling keratinocyte migration [27]. Therefore, the observed MMP-1 and MMP-10 down-regulation by exogenous vitamin D in vitro may be considered as a favorable response, as it could signify favorable tissue remodeling in vivo. Although the mechanism for vitamin D-mediated modulation of MMP-1 and MMP-10 expression has not been described, there are various reports about vitamin D-mediated regulation of MMPs. For example, calcitriol inhibited MMPs expression and activity in leucocytes infected with Mycobacterium tuberculosis [6], and it attenuated MMP-9 expression in a keratinocyte cell line, in turn inhibiting c-Jun N-terminal kinase (JNK) and nuclear factor kappa-light-chain-enhancer of activated B cells (NF-jB) signaling cascades [1]. In chondrocytes, calcitriol activated MMP-13 expression through the p38 MAPK pathway [7]; and, as calcipotriol (a synthetic derivative of calcitriol), it reduced both transcript and protein levels of MMP-9 and MMP-13 through inhibition of extracellular signal-regulated kinases (ERK) and p38 phosphorylation, respectively, in a squamous cell carcinoma cell line [39]. Further experiments will be needed to describe in detail the vitamin D-mediated regulation of specific MMPs in keratinocytes. The development of type 2 diabetes involves impaired pancreatic b cell function, insulin resistance, and inflammation. Although mechanistically unclear, it has been suggested that both environmental and genetic factors seem to be involved in type 2 diabetes development [59]; also, human and experimental data support the role of vitamin D in these pathways [46, 69]. Vitamin D is not only a regulator of calcium and phosphate homeostasis, but it also has numerous extra-skeletal effects [66]. Vitamin D deficiency may predispose to glucose intolerance, altered insulin secretion and type 2 diabetes [47], either through a direct action via vitamin D receptor (VDR) activation or indirectly via calcemic hormones, and also via inflammation [62]. Given the relationship between type 2 diabetes and vitamin D, the effects observed in our keratinocyte culture experiments may be explained by the compensation of a vitamin D deficiency. Therefore, to better clarify the role of diabetes-related factors in vitamin D-induced effects, it would be important to perform the same experiments using biopsies from chronic wounds with different etiologies. Conflict of interest The authors declare no conflicts of interest. All authors have read and approved the final version of the article.

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