Functional Responses of Dermal Fibroblasts to Low Nutrition and Pro-Inflammatory Stimuli Mimicking a Wound Environment in Vitro

doi:10.21203/rs.3.rs-1458136/v1

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Research Article

Functional Responses of Dermal Fibroblasts to Low Nutrition and Pro-Inflammatory Stimuli Mimicking a Wound Environment in Vitro

https://doi.org/10.21203/rs.3.rs-1458136/v1

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posted 21 Mar, 2022

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Dermal fibroblasts (DF) constitute one of key cells involved in wound healing. However, the functions they perform in wound conditions remain poorly understood. This study involved exposing DF to low nutrition and to low nutrition + LPS for 5 days as conditions representing the wound. Although DF exhibited increasing metabolic activity in time under all conditions including control, the proliferation did not change in both low nutrition and low nutrition + LPS. Only the low nutrition + LPS was found to potentiate the migration and pro-inflammatory phenotype (IL6 release) of DF. The potential of DF to contract collagen hydrogel declined only under the low nutrition as a consequence of low cell number. The expression of α-SMA was reduced under the both conditions independently of the cell number. The remodeling capability of DF was affected under the both conditions as documented by the enhanced MMP2 activity. Finally, the production of collagen type I was not affected by either condition. The study shows that low nutrition as the single factor is able to delay the healing process. Moreover, the addition of the mild pro-inflammatory stimulus represented by LPS may amplify the cell response in case of decreased α-SMA expression or excite DF to produce IL6 impairing the healing process.

dermal fibroblasts

inflammation

lipopolysaccharide

low nutrition

wound healing

Dermal fibroblasts constitute a heterogeneous population of mesenchymal cells that are present in the dermis of the skin. They play a major role in terms of tissue structural support due to their producing an extracellular matrix (ECM). In addition, they are able to secrete and respond to cytokines and growth factors. In this way they communicate both with each other and with other cell types. As a result, dermal fibroblasts actively participate in the regulation of skin physiology processes such as tissue development and differentiation, as well as remodeling and repair during the wound healing process [1].

Dermal fibroblasts play an important role under the stress conditions of healing wound environments in both the inflammatory and proliferative phases of the wound healing process. They make up key players in terms of maintaining skin homeostasis and orchestrating physiological tissue repair. During the early stages of the wound repair process, dermal fibroblasts become highly proliferative, migrate to the wound site from the regional connective tissue and produce various ECM components including glycosaminoglycans (GAG) and collagen so as to form provisional granulation tissue. This granulation tissue further provides a scaffold for the migration of resident cells from local tissue and blood circulation toward the wound [2]. The regulation of fibroblast proliferation and migration occurs through the local release of various cytokines, e.g. IL6 and IL8, which contribute toward enhancing inflammation in the wound [3]. Granulation tissue degradation occurs in the later stages of the wound healing process via the action of matrix metalloproteinases (MMPs) produced by fibroblasts and other cells. The balance between MMPs and metalloproteinase tissue inhibitors (TIMPs) is essential for the remodeling of the ECM into the scar tissue, thus resulting in normal wound resolution [4] either via the degradation of the ECM or indirectly via their ability to affect cell behavior [5]. Some of the fibroblasts differentiate into myofibroblasts during the wound healing process following stimulation exerted by transforming growth factor-beta (TGF-β). Myofibroblasts with stress fibers containing α-smooth muscle actin (α-SMA) contract and close the wound. In addition, TGF-β promotes the production of collagen by dermal fibroblasts [6]. The final phase of healing consists of the remodeling of the granulation tissue into more mature and stronger scar tissue. The signaling changes and the proliferative phase are suppressed in disrupted healing; the fibroblasts produce a higher level of pro-inflammatory cytokines and exhibit decreased migratory and mitotic activity, insufficient ECM production and imbalance between the secretion of MMPs and TIMPs [7].

Several in vitro models exist that simulate the wound environment with respect to dermal fibroblasts, the most common of which involve fibroblasts isolated directly from wounds [8–10]. Dermal fibroblasts derived from chronic wounds exhibit decreased adhesion, proliferation and ability to withstand oxidative stress compared to those derived from uninjured skin, thus resulting in the early onset of senescence. Moreover, they express lower levels of chemokine genes (CXCL-1, -2, -3, -5, -6, 12) and are unable to correctly express a stromal address code which leads to the requirement for the guidance of infiltrating leukocytes to the site of injury [8]. The ratio of differentiated myofibroblasts was higher in the chronic wound group compared to dermal fibroblasts derived from properly healing wounds. The examination of several cytokines led to the conclusion that the production of IL6 was enhanced in the chronic wound group while the production of IL8 was enhanced in the properly healing wound group [10]. Burn wounds exhibited the significantly higher production of IL8 than did the healed wounds and intact skin; no difference was determined in terms of the production of IL8 between the two latter groups [11]. On the other hand, although IL8 typically comprises a major cytokine expressed by many cells following LPS stimulation, certain fibroblast subtypes (e.g. lung fibroblasts) did not produce IL8 following LPS stimulation [12, 13].

A further wound environment in vitro concept consists of the culturing of dermal fibroblasts with inflammatory stimuli, e.g. bacterial lipopolysaccharide (LPS). The treatment of fibroblasts with LPS was found to cause a decrease in viability and the increased production of the MCP-1, MIP-2, CINC and RANTES chemokines [14, 15], which initiated the recruitment of hematopoietic cells which, in turn, led to the further activation of the fibroblasts via the secretion of TNF-α [16]. Moreover, fibroblasts stimulated with LPS exhibited a 300-fold increase in IL8 mRNA compared to non-affected fibroblasts [17]. LPS has also been studied in an ex vivo human skin organ culture and was found to enhance the production of IL6 and IL8 in a dose-dependent manner [18]. LPS has also been used in in vivo studies; 10 µg/ml of K. pneumoniae-derived LPS delayed wound closure, prolonged the inflammatory response, reduced collagen deposition, induced early cell death in granulation tissue and inhibited wound edge proliferation in mice [19]. A further in vivo study showed that the production of collagen and pro-inflammatory cytokines (e.g. IL6, IL1β) and the wound breaking force were higher in mice treated with LPS [20].

A further important factor present in delayed wounds comprises a lack of nutrition supply due to improper wound vascularization followed by peripheral arterial disease, which is manifested by tissue ischemia and which, in chronic mode, can lead to delayed wound healing [21]. In the in vitro set up, fetal bovine serum represents the main source of nutrition for cells, and the decreasing of the concentration thereof is used regularly as a stress factor [22].

Nevertheless, no complex description of the behavior of dermal fibroblasts in the wound environment has yet been performed. Importantly, since several stress factors present in the wound environment have been characterized, it is possible to speculate which factor or factors exert an effect and the extent to which they are able to influence dermal fibroblasts.

The study involved the selection of two wound stress factors - low nutrition and pro-inflammatory stimuli represented by a decreased concentration of fetal bovine serum in the culture media and the addition of bacterial LPS to the cell culture media, respectively, and the study of the typical behavior of dermal fibroblasts in terms of wound healing under both conditions. Typical parameters for the description of the behavior of dermal fibroblasts include metabolic activity and the production of pro-inflammatory cytokines (IL6 and IL8) and matrix metalloproteinases (MMPs) as inflammatory phase behavioral patterns, and the proliferation, migration, contraction ability and production of collagen type I and α-SMA as characteristics of the proliferative phase.

2.1 Cell isolation and culture

Normal human dermal fibroblasts (NHDF) were isolated from skin residues via the digestion-migration method following plastic surgery interventions. Immediately following skin biopsy, the samples were immersed in physiological solution and transported to the cell culture lab for immediate isolation. The samples were washed in Hank’s balanced salt solution (HBSS) (Sigma Aldrich) containing penicillin (100 U/ml)/streptomycin (0.1 mg/ml) (Biochrom, United Kingdom) and gentamicin (50 µg/ml) (Biochrom). The samples were cut into 3 mm² pieces and digested overnight at 37°C in Petri dishes (Techno Plastic Products, Trasadingen, Switzerland) in HBSS containing collagenase type I (100 U/ml, Thermo Fisher Scientific, USA). On the following day, the suspension containing the digested tissue was shaken intensively employing a vortex for 30 sec and filtered through a 100 µm nylon cell strainer (Falcon™, Thermo Fisher Scientific). The cell suspension was then transferred to a 75 cm² cultivation flask (Techno Plastic Products) containing 10 ml of culture medium composed of low glucose Dulbecco’s Modified Eagle’s Medium (DMEM) (Thermo Fisher Scientific), 10% of heat-inactivated FBS (Thermo Fisher Scientific), penicillin (100 U/ml)/streptomycin (0.1 mg/ml) (Biochrom), 0.5% of L-glutamin (Biosera, France) and 1.0% of non-essential amino acids (Biosera). The NHDF were cultured at 37°C, 5% CO₂ up to 80% confluence and then passaged. Only those NHDF from the 3rd – 5th passages were used in the experiments. The media were changed every 2nd − 3rd day of cell culturing.

2.2 In vitro simulation of wound conditions

Two types of wound conditions were simulated. Low nutrition conditions were simulated by means of a culture medium containing only 2% of FBS, and low nutrition + LPS conditions were simulated by means of a culture medium containing 2% of FBS and 0.1 µg/ml of lipopolysaccharide (LPS) that originated in pathogenic E. coli strain type O111:B4 (Sigma Aldrich). The NHDF culture medium mentioned in Chap. 2.1 provided for the control conditions.

2.3 Metabolic activity

The metabolic activity was estimated using alamarBlue assay that converted blue resazurin to pink resorufin. The NHDF were seeded on 96-well plates (Techno Plastic Products) at a density of 6000 cells/cm² (for the metabolic activity and cell proliferation) in a culture medium and cultured overnight at 37°C and 5% CO₂. On the following day, the NHDF were treated as described above (2.2). The media were collected and frozen at -80°C at time points 0, 1, 2, 3 and 5 days following culture treatment and subjected to the performance of alamarBlue assay; 100 µl of alamarBlue solution (ThermoFisher Scientific) 10x diluted in the same media as used for the culture treatment was added to each well and the NHDF were incubated for 2 hours at 37°C, 5% CO₂. Subsequently, 100 µl of the media was transferred to a black 96-well test plate (ThermoFisher Scientific) and the fluorescence was measured at 530 nm (ex) and 590 nm (em) in a microplate reader (Synergy HT, Biotek, USA). The change in the metabolic activity over time was expressed as the ratio of the metabolic activity at days 1, 2, 3 and 5 to that at day 0 for each treatment. The results were expressed as the mean ± standard deviation from 8 independent NHDF donors.

2.4 Cell proliferation

Cell proliferation was estimated via a total nuclei count. Following the performance of alamarBlue assay, the NHDF were washed in phosphate buffer saline (PBS) and the nuclei were stained using Hoechst #33342 (Thermo Fisher Scientific) diluted at 1:2000 with the culture media for 5 min at 37°C, 5% CO₂. The NHDF were then washed in PBS and images were captured of the whole of the well areas (0.335 cm²) using an Olympus IX-83 (Japan). Nine sequential pictures were taken covering the whole of the well areas using an Olympus UPlanFL N 4x/0.13 objective followed by automatic whole-well area picture decomposition employing VisiView software (Visitron Systems, Germany). The number of nuclei in each well was determined via counting in Fiji software (National Institute of Health, Bethseda, USA). The change in the proliferation rate over time was expressed as the ratio of the cell count at days 1, 2, 3 and 5 to that at day 0 for each treatment. The results were expressed as the mean of the cell number ± standard deviation from 8 independent NHDF donors.

2.5 Scratch wound assay

The NHDF were seeded on 6-well test plates (Techno Plastic Products) at a density of 30 000 cells/cm² in the culture medium and cultured overnight at 37°C and 5% CO₂. On the following day, the cell monolayers were scratched by means of a 10 µl pipette tip so as to create four crosses (wounded areas) per well thus mimicking the “wound bed”. The detached cells were washed carefully in PBS and treated with 0.1 µg/ml of LPS in a serum free medium (low nutrition + LPS) in order to eliminate the cell proliferation effect, and cultured for 5 days. The serum-free culture medium (low nutrition) was applied as the control treatment. Immediately following treatment, images of the four wounded positions in each well were captured using an Olympus UPlanFL N 4x/0.13 objective and Olympus IX-83 (Japan) inverted microscope. The positions were saved using cellSens Dimension 1.12 microscope software. Images of the migrating cells were taken at time points 0, 1, 2, 3 and 5 days following treatment. The final images were analyzed using TScratch software (CSELab, Switzerland) via the quantification of the wounded areas (i.e. the areas without cells) as a percentage of the whole of the picture area. The change in the migration rate of the cells was expressed as the ratio of the wounded area at days 1, 2, 3 and 5 to the wounded area at day 0. The results were expressed as the mean ± standard deviation from 3 independent NHDF donors.

2.6 IL6, IL8 and collagen type I quantification

The culture media were collected at each time point, i.e. 0, 1, 2, 3 and 5 days, and stored at -80°C. The concentrations of IL6, IL8 and collagen type I were assessed by means of ELISA using the Human IL-6 ELISA Ready-SET-Go! Kit (eBioscience, USA), the Human IL-8 ELISA Ready-SET-Go! Kit (eBioscience, USA) and the Human Pro-Collagen I alpha 1 ELISA Kit (Abcam, UK), respectively. The assays were performed according to the manufacturer’s instructions. The change in the time for the production of IL6, IL8 and collagen type I was expressed as the ratio of picograms per cell (IL6 and IL8) or picograms per 1000 cells (collagen type I) at days 1, 2, 3 and 5 to picograms per cell (IL6 and IL8) or picograms per 1000 cells (collagen type I) at day 1. The results were expressed as the mean ± standard deviation from 6–8 independent NHDF donors.

2.7 MMP2 and MMP9 quantification

The culture media were collected at time points 1 and 5 days and stored at -80°C. The pure cell culture media containing either 10% or 2% of FBS were analyzed so as to determine the basal MMP activities of the culture media. The MMP2 and MMP9 were analyzed via gelatin zymography using an SDS-PAGE system (7.5% separating gel, 4% stacking gel) containing 0.05% of gelatin and using a PageRulerPlus Prestained Protein Ladder, 10 to 250 kDa (ThermoFisher Scientific, USA) as a marker of MW. Following electrophoresis, the gels were washed in 2.5% Triton X-100 for 30 min while being shaken so as to remove the SDS and to renature the MMPs in the gels. The gels were then incubated overnight in a gelatinase activation buffer (100 mM Tris-HCl, pH 7.4; 5 µM CaCl₂; 1 µM ZnCl₂), stained with Coomassie Brilliant Blue G-250 for 30 min and de-stained using de-staining solution. The gels were scanned using the ChemiDoc MP Imaging System (BioRad, USA). The gels (Fig. 4a) were then cropped from the original scan (Fig. 9) using free Inkscape software (https://inkscape.org/). The MMP2 and MMP9 activities were determined via the software quantification of the degraded bands in Fiji software. The MMP activity was determined in two stages. Firstly, the basal MMP activity values of the pure culture media were subtracted from those of the culture media of the analyzed samples; secondly, the MMP activity values were normalized to the total cell number. The results were expressed as the mean ± standard deviation from 4 independent NHDF donors.

2.8 α-SMA and collagen type I visualization and α-SMA quantification

The NHDF were seeded on a µ-Slide 4-well (Ibidi, Germany) at a density of 6000 cells/cm² (visualization) and on 96-well plate (quantification) in the culture medium and cultured overnight at 37°C and 5% CO₂. On the following day, the NHDF were treated as described above (2.2). 5 days following treatment, the NHDF were fixed with fresh 4% paraformaldehyde. The cells were then permeabilized with cold 0.1% Triton X-100 solution for 15 minutes. After rinsing with PBS, the non-specific bonds were blocked with 5% BSA in PBS for 30 minutes at room temperature. Subsequently, the samples were incubated overnight at 4°C with the following primary antibodies diluted in PBS: anti-alpha smooth muscle actin antibody (ab7817, Abcam) diluted to a final concentration of 1 µg/ml and collagen I alpha I antibody (MAB 6220, R&D Systems) diluted to a final concentration of 125 µg/ml. Following rinsing with PBS, secondary goat anti-mouse IgG H&L antibody conjugated with Alexa Fluor 488 (ab150113, Abcam) was applied to the samples (diluted in PBS to a final concentration of 1 µg/ml) for 1 hour at room temperature in the dark. The nuclei were stained with Hoechst #33342 diluted at 1:2000 in the final 10 minutes. The samples were then rinsed with PBS and scanned using an Olympus IX-83 (Japan). An Olympus UPlanFL N 20x/0.50 objective was used for the visualization of the α-SMA. The visualization of the collagen type I was conducted using an Olympus LUCPlanFL N 40x/0.60 objective. With respect to the quantification of the α-SMA, nine sequential images covering the whole well area were captured by means of an Olympus UPlanFL N 4x/0.13 objective followed by automatic whole well area picture decomposition using VisiView software. The number of nuclei and the area of the α-SMA were determined from the same well by means of Fiji software. The results were expressed as the mean of the α-SMA area at day 5 (mm²) normalized to 1000 cells at day 5 ± standard deviation from 3 independent NHDF donors.

2.9 NHDF contraction ability

The NHDF were seeded in hydrogels produced from rat tail collagen type I. The lyophilized rat tail collagen was dissolved in 0.1% acetic acid to a concentration of 5 mg/ml followed by the preparation of a neutralized collagen suspension containing NHDF (final collagen concentration 3 mg/ml, NHDF density 30 000 cells/ml). 2 ml of the collagen suspension with NHDF was added to the wells of 24-well plates and left at 37°C and 5% CO₂ to polymerize. After 1 hour, once the hydrogels were completely stiff, 1 ml of the culture medium was added to the hydrogels. On the following day, the NHDF were treated as described above (2.2). An equal volume of fresh culture medium was replaced three times per week. The diameter of the hydrogels was measured using a caliper at time points 1, 6 and 13 days. The results were expressed as the mean ± standard deviation from 6 independent NHDF donors.

2.10 Statistical analysis

All the data was expressed as the mean ± standard deviation from 3–8 independent NHDF donors depending on the type of analysis conducted. The Shapiro-Wilk test was used to ascertain data normality and the Leven’s test was employed for the assessment of the equality of the variances for a variable calculated for two or more groups. The statistical significance of the differences between the control, low nutrition and low nutrition + LPS groups was determined via one-way ANOVA (p < 0.05) followed by the Fisher’s LSD test for the quantification of the α-SMA and MMP2 and by the Kruskal-Wallis test for the metabolic activity, proliferation, contraction and quantification of the IL6, IL8, collagen type I and MMP9. The Mann-Whitney U test (p < 0.05) was used for the scratch wound assay for migration since only two groups were compared. The statistical significance of the differences between the time points within each group was determined using the Mann-Whitney U test (p < 0.05). All the statistical analyses were performed in Statistica v12 software (Tibco Software, Palo Alto, California, USA).

3.1 Metabolic activity is not affected by low nutrition and low nutrition + LPS

The effect of low nutrition and low nutrition + LPS conditions on the NHDF metabolic activity was estimated by means of alamarBlue assay. The NHDF were cultured for 5 days under control, low nutrition and low nutrition + LPS conditions, and the metabolic activity was estimated after 0, 1, 2, 3 and 5 days of treatment (Fig. 1a). Under all three conditions, the cultured NHDF exhibited increasing tendencies in Fig. 1a terms of metabolic activity during the whole of the cultivation period (Mann Whitney U test; p < 0.05). The comparison of the metabolic activity of the NHDF cultured under the various conditions revealed no significant difference between the groups, with the exception of the occurrence of a significantly higher NHDF metabolic activity under the low nutrition (4.08 ± 0.76) conditions than that of the control (2.60 ± 0.59) at day 3 (Kruskal-Wallis test; p < 0.05).

3.2 The dermal fibroblasts did not proliferate under either low nutrition or low nutrition + LPS conditions

The effect of low nutrition and low nutrition + LPS conditions on NHDF proliferation was evaluated by means of the total cell count (Fig. 1b). The NHDF were found not to have proliferated after 5 days under either low nutrition or low nutrition + LPS conditions. The comparison of the proliferation of the NHDF under the various conditions and at particular time points revealed no significant difference between the low nutrition and low nutrition + LPS conditions at any of the time points. However, at days 3 and 5 the proliferation of the NHDF was observed to be significantly lower under both the low nutrition (1.86 ± 0.78 for day 3; 1.67 ± 0.41 for day 5) and the low nutrition + LPS (1.89 ± 0.79 for day 3; 2.17 ± 1.11 for day 5) conditions than under the control conditions (3.30 ± 0.69 for day 3; 4.90 ± 0.90 for day 5) (Kruskal-Wallis test; p < 0.05).

3.3 LPS stimulated the migration of dermal fibroblasts

The effects of low nutrition and low nutrition + LPS conditions on NHDF migration were evaluated by means of scratch wound assay. The NHDF migrated for 5 days and the wounded area was measured 0, 1, 2, 3 and 5 days following wounding (Fig. 2). Interestingly, the low nutrition + LPS conditions, particularly with respect to the addition of LPS, were observed to promote NHDF migration significantly from day 2. After 2 days of migration, the wounded area under low nutrition conditions was determined at 71.5 ± 12.4% and under low nutrition + LPS conditions at 56.7 ± 7.0%. At the subsequent time points, the wounded areas were determined at: 62.7 ± 11.0% (low nutrition) versus 42.2 ± 4.7% (low nutrition + LPS) after 3 days and 56.8 ± 13.3% (low nutrition) versus 36.5 ± 9.7% (low nutrition + LPS) after 5 days (Mann-Whitney U test; p < 0.05). The healthy physiological condition of NHDF throughout scratch wound assay is demonstrated in Fig. 2b. Third control group containing 10% FBS was avoided in this assay to prevent the effect of proliferation on wound area closure as used in other study [23, 24].

3.4 Low nutrition + LPS promoted the pro-inflammatory behavior of the dermal fibroblasts

The pro-inflammatory response of the NHDF under low nutrition and low nutrition + LPS conditions was estimated via the quantification of IL6 (Fig. 3a) and IL8 (Fig. 3b) in the cell culture media and normalized to the cell number at each day and, subsequently, to the value at day 1. The production of IL6 by the NHDF was significantly higher under the low nutrition + LPS conditions than under the low nutrition and control conditions over time from day 2 (Kruskal-Wallis test; p < 0.05). The production values were determined at 1.35 ± 0.50 (low nutrition + LPS), 0.85 ± 0.18 (low nutrition) and 0.78 ± 0.20 (control) at day 2; 2.16 ± 1.04 (low nutrition + LPS), 0.83 ± 0.49 (low nutrition) and 0.78 ± 0.55 (control) at day 3, and 5.47 ± 2.07 (low nutrition + LPS), 1.07 ± 0.60 (low nutrition) and 1.16 ± 1.02 (control) at day 5. The production of IL8 by the NHDF exhibited a greater increasing tendency over time when cultured under low nutrition + LPS conditions than it did under the low nutrition and control conditions. However, the concentration of IL8 did not differ significantly between the groups at any of the time points (Kruskal-Wallis test; p > 0.05).

3.5 Both low nutrition and low nutrition + LPS stimulated MMP2 activity and did not affect the MMP9 activity in the dermal fibroblasts

The effects of low nutrition and low nutrition + LPS conditions on the NHDF secretion of MMP2 and MMP9 was evaluated via gelatin zymography at the 1- and 5-day time points. The signals from the MMP2 and MMP9 activities in the pure cell culture media without cells were subtracted from the signals of the tested samples and normalized to the cell number (Fig. 4a-c and Fig. 9). In both, the low nutrition and low nutrition + LPS conditions, the activity of MMP2 per cell at day 1 was significantly promoted (Fig. 4b), with MMP2 per cell counts of 1.25 ± 0.60 (low nutrition + LPS) and 0.58 ± 0.26 (low nutrition) versus − 0.68 ± 0.37 (control). The MMP9 increased significantly under the low nutrition conditions at day 1, evincing an MMP9 per cell count of 0.38 ± 0.17 (low nutrition) versus − 0.37 ± 0.33 (control). The negative values of the control conditions were the result of the higher activity in the control cell culture media without cells (Fig. 4a). We decided to use negative values instead of simple zero values. Similarly, at day 5 (Fig. 4c), the activity of MMP2 per cell was increased significantly under both the low nutrition and low nutrition + LPS conditions, with MMP2 per cell counts of 1.48 ± 0.50 (low nutrition + LPS) and 1.54 ± 0.82 (low nutrition) versus 0.35 ± 0.17 (control). The level of MMP9 per cell was not found to have changed at day 5 under either the low nutrition or low nutrition + LPS conditions; however, the level of MMP9 per cell under the control conditions was observed to attain the values of the low nutrition and low nutrition + LPS (One-way ANOVA; p < 0.05 followed by Fisher’s LSD for the MMP2; Kruskal-Wallis test; p < 0.05 for the MMP9).

3.6 Both the low nutrition and low nutrition + LPS conditions reduced the NHDF contraction ability

The impacts of the low nutrition and low nutrition + LPS conditions on the contraction ability of the NHDF were evaluated via the diameter measurement of collagen hydrogels seeded with the NHDF at the 3-, 6- and 13-day time points (Fig. 5 and Fig. 10). After 1 day and 6 days of culturing under the control, low nutrition and low nutrition + LPS conditions, the contraction ability of the NHDF was found not to have changed. After 13 days of culturing, the highest NHDF contraction ability was observed in the control conditions, with scaffold diameters of 10.27 ± 1.3 mm (control) versus 11.60 ± 0.99 mm (low nutrition) and 11.89 ± 1.33 mm (low nutrition + LPS) (Kruskal-Wallis test; p < 0.05).

3.7 Both the low nutrition and low nutrition + LPS conditions decreased the area of the α-SMA positive NHDF, with the more profound effect of the low nutrition + LPS

The area of α-SMA positive NHDF per the total cell number was visualized and quantified after 5 days of culturing under the control, low nutrition and low nutrition + LPS conditions (Fig. 6a-d). The area of α-SMA decreased under both the low nutrition and low nutrition + LPS conditions after 5 days of culturing compared to the control conditions. Moreover, the low nutrition + LPS conditions acted to decrease the area of α-SMA to a greater extent than did the low nutrition conditions, with areas of α-SMA 0.17 ± 0.04 mm²/1000 cells (control), 0.14 ± 0.02 mm²/1000 cells (low nutrition) and 0.07 ± 0.04 mm²/1000 cells (low nutrition + LPS) (Fig. 6a) (One-way ANOVA; p < 0.05 followed by the Fisher’s LSD test). The differences between the control, low nutrition and low nutrition + LPS conditions are apparent from the selected representative microscopy images shown in Fig. 6b-d and Fig. 11.

3.8 Neither the low nutrition or low nutrition + LPS conditions exerted an effect on the synthesis and production of collagen type I by the NHDF

The synthesis of intracellular collagen type I by the NHDF was visualized using fluorescent microscopy after 5 days of cultivation under the control, low nutrition and low nutrition + LPS conditions. The production of collagen type I by the NHDF was evaluated by means of ELISA from the culture media. The synthesis of intracellular collagen type I was found to be comparable under all the tested conditions (Fig. 7b-d). The production of collagen type I increased during culturing in the low nutrition + LPS, low nutrition and control media (Fig. 7a); however, no difference was detected between the groups at any of the time points (Kruskal-Wallis test; p > 0.05).

The dermal fibroblasts cultured under the low nutrition conditions exhibited a slightly higher metabolic activity level than those cultured under the control conditions; however, the increase was significant at day 3 only. A quiescence phenotype and pathways that generated NADPH were induced when the dermal fibroblasts were cultured under low nutrition conditions represented by a medium with 0.1% of FBS [25]. AlamarBlue containing resazurin, the substrate used in this study for the estimation of the metabolic activity is able to be reduced by NADPH [26]. Therefore, we hypothesized that low nutrition conditions used in our study induces a quiescence phenotype and that the metabolic activity of the dermal fibroblasts under low nutrition and low nutrition + LPS conditions is, paradoxically, increased due to the generation of increased levels of NADPH produced by the activated metabolic pathways. The results of the metabolic activity response of the fibroblasts to the LPS in the culture media obtained in other studies were inconsistent. 0.2 µg/ml of LPS (the authors did not state the LPS origin) in a serum-free medium that contained other components that supported proliferation and plating efficiency was observed not to affect the metabolic activity of the dermal fibroblasts [27]. 10 µg/ml of LPS (E. coli) in a medium with 10% of FBS acted to decrease the metabolic activity of gingival fibroblasts [15], and 25 µg/ml of LPS (derived from E. coli O111:B4) in a medium containing 10% of FBS was seen to increase the metabolic activity of gingival fibroblasts [28]. The inconsistency in the data of our and other studies was presumably the result of the differing sources of the fibroblasts and differences in the experimental set ups. Based on our results and on experience of other researchers we can conclude that the metabolic activity of the fibroblasts depends primarily on the place of the wound occurrence.

The experimental results indicated that the dermal fibroblasts did not proliferate under either the low nutrition or the low nutrition + LPS conditions. Similarly, the serum-free medium exhibited a sharp decrease in the cell count [29]. With respect to the addition of LPS, 1 µg/ml of LPS in a medium containing 10% of calf serum led to an increase in the proliferation of mouse lung fibroblasts [30], while, conversely, 10 µg/ml of LPS in a medium containing 10% of FBS acted to decrease the proliferation of gingival fibroblasts [15]. Based on the information provided in the studies mentioned above, we propose that the 0.1 µg/ml of LPS used in this study was too low a concentration to exert an impact on the proliferation of the dermal fibroblasts. Thus, we conclude that the dermal fibroblasts in our study did not proliferate due to the low nutrition conditions rather than the presence of LPS, which is supported by the results obtained by Ejiri et al. Importantly, the comparison of the metabolic activity and proliferation presented in our paper provides evidence that while the two methods are sometimes erroneously interchanged [23, 28, 31], the metabolic activity does not equate to proliferation. Whereas the dermal fibroblasts under both the low nutrition and low nutrition + LPS conditions exhibited enhanced metabolic activities, they did not proliferate, which suggests that the culturing conditions of the experiment had the potential to increase the metabolic activity of the cells as a reflection of the cellular activity that focused primarily on surmounting the “uncomfortable” culturing conditions, accompanied by the non-proliferative state of the cells.

Scratch wound assay was found to provide an appropriate method for the assessment of cell migration into the wounded area [22]. We demonstrated that dermal fibroblasts cultured in the presence of LPS migrate to a greater extent than do dermal fibroblasts cultured without LPS. The enhanced migration rate of mouse adventitial fibroblasts was observed in a culture medium with 10 µg/ml of LPS [23]. Similarly, treatment with 0.4 µg/ml of LPS increased the migration of mouse embryo cell line fibroblasts through the positive feedback between β-catenin and COX-2 in concentration in a time-dependent manner [24]. The results of both studies are consistent with our results, i.e. that LPS exerts a stimulatory effect on dermal fibroblast migration. It is important to note, that 10% (v/v) of FBS in culture media, which is the compound of control condition in our study, promote cell proliferation. However, the purpose of scratch wound assay is to monitor the covering of wounded area by cell migration not by cell proliferation. For this reason, we avoided this control condition (10% FBS culture medium) from scratch wound assay, which is consistent with other studies [32, 33].

The production of IL6 was significantly enhanced when the dermal fibroblasts were cultured under low nutrition + LPS conditions independent of the cell count. Similarly, other researchers have shown that the secretion of IL6 increased when gingival fibroblasts were cultivated with 0.1 µg/ml of LPS from E. coli in a medium containing 10% of fetal calf serum. The NFκB signaling pathway was found to play an essential role in the regulation of the expression of IL6 by LPS-stimulated fibroblasts [34]. Similarly, 10 µg/ml of LPS from P. aeruginosa significantly enhanced the production of IL6 by nasal polyp-derived fibroblasts [35]. Therefore, we propose that the presence of LPS exerts a primary effect on the production of IL6. In addition, a complex of IL6 with the IL6 receptor suppresses IL1β, TNFα and PDGF-AA-induced dermal fibroblast proliferation [36]. These findings correlate well with the non-proliferating fibroblasts in our experiment. An increased level of IL6 following LPS treatment was also observed in vivo. A gel containing 10 mg/g of LPS from S. typhi applied to incision-wounded mice was observed to enhance the production of IL6 [20]. Lung fibroblasts cultured with 1 µg/ml of LPS in a 10% calf serum medium led to the production of increased levels of IL6 and IL8, which resulted in a decrease in the metabolic activity of the fibroblasts in an autocrine manner [31]. However, the origin of the fibroblasts and the origin and concentration of the bacterial LPS appear to be of significant importance. For instance, 10 µg/ml of LPS from E. coli enhanced the production of IL8 in human nasal but not lung fibroblasts [13]. Similarly, LPS from E. coli and S.typhimurium led to increases in the levels of IL6 and IL8 after 48 hours of exposure (0.1 µg/ml) in an ex vivo human skin organ culture, while LPS that originated in S. enteritidis failed to do so [18]. Moreover, no increase in the level of IL8 was observed following the application of 1 µg/ml of LPS from the E. coli stimulation of pulmonary fibroblasts over 24 hours [12]. Similarly, 0.05 and 0.1 µg/ml of LPS from P. aeruginosa did not affect the production of IL6 and IL8 by corneal fibroblasts; however, in this case, it is necessary to take into account the age of the patients that provided the corneal samples, i.e. the samples were provided by elderly persons as opposed to the mostly young donors of the dermal fibroblasts used in this study. Interestingly, when the same LPS treatment was performed using ulcerated corneal fibroblasts, the production of IL6 and IL8 increased significantly [37]. Biopsies from burn wounds produced increased levels of IL8 compared with those from healed wounds and intact skin [11]. The production of IL8 by dermal fibroblasts increased significantly following treatment with 0.2 µg/ml of LPS in a serum-free medium that, nevertheless, contained components that supported proliferation and plating efficiency (it should be noted that the authors did not state the origin of the bacterial species of the LPS) [27]. Similarly, 10 µg/ml of LPS from P. aeruginosa significantly enhanced the production of IL8 by nasal polyp-derived fibroblasts [35]. Our results suggest that dermal fibroblasts are able to produce a higher level of IL8 under low nutrition + LPS conditions, especially after 5 days of culturing. However, our IL8 assay data exhibited a high degree of variability, thus rendering the differences between the experimental groups insignificant. Moreover, interleukin values were observed in our study for 5 days of cell culturing while in the studies mentioned above the researchers observed interleukin production up to just 3 days of cell culturing. Therefore, we attribute the discrepancies between our results and those of other research teams to the differing time points of the analysis. Moreover, we used the primary dermal fibroblasts isolated from different human donors, which make this study unique. The use of dermal fibroblasts from up to 8 donors provides more representative samples selected from the whole population, however is affected by higher data variability. The analysis of IL8 production by dermal fibroblasts under the same conditions as used in this study has not been reported to date in the literature. Therefore, we can only speculate on the general impact of LPS on the production of IL8 by dermal fibroblasts.

The presence of MMPs in wounded tissue is essential with respect to cell migration, tissue remodeling and the regulation of the level of cytokines [5, 38]. The control conditions applied in this study contained more FBS (10%) than did the low nutrition and low nutrition + LPS conditions (2%). Our results revealed that the FBS contained MMPs regardless of those produced by the cells (Fig. 4a; culture media without cells). Nevertheless, the results also indicated higher levels of MMP2 under the low nutrition and low nutrition + LPS conditions than that of the control conditions over time and independent of the cell count. We propose that the increases in the MMP2 levels in both the stress media were due primarily to low nutrition rather than the presence of LPS. The level of MMP9 remained unchanged under both stress conditions. After 48 hours of the exposure of bovine dermal fibroblasts to LPS (5 µg/ml) from E. coli, increased levels of MMP2 and MMP9 were observed [39]. Our study involved the culturing of dermal fibroblasts in 0.1 µg/ml of LPS. Our findings and those of Akkoc et al. indicate that LPS is capable of enhancing the production of MMP2 and MMP9 by dermal fibroblasts in concentrations of higher than 0.1 µg/ml. The release of MMPs is dependent not only upon the origin of the LPS but also on that of the fibroblasts [40]. An increased level of MMP9 in nasal polyp-derived fibroblasts was determined following exposure to 10 µg/ml of LPS (P. aeruginosa) for 12 hours [35]. We conclude that, with respect to our experimental set up, the dermal fibroblast secretion of MMP2 was induced primarily by low nutrition, and that the LPS concentration of 0.1 µg/ml was too low to enable the release of the active form of MMP9.

Our study revealed that the contraction ability of dermal fibroblasts decreased under both the low nutrition and low nutrition + LPS conditions, most likely due to the decreased number of cells under the low nutrition conditions. These results are consistent with those obtained from a previous in vivo study; 10 µg of LPS subcutaneously injected prior to wounding delayed the wound closure process [19] (in this case the LPS originated from K. pneumoniae rather than E. coli as used in our study). However, other researchers have observed the opposite effect. Gels containing 0.2, 2 and 10 mg/g of LPS from S. typhi were applied to the incision-wounded skin of mice. The wound closure process was enhanced in the presence of LPS and was dose-dependent [20]. A similar effect was also observed in vitro. 2 and 5 µg/ml of LPS (origin not specified) in a culture medium with 1% of FBS was found to enhance the contraction of intestine fibroblast-mediated collagen in a dose dependent manner [41]. Whether the differences in the types of fibroblasts and/or origin and concentrations of LPS were responsible for the discrepancies in the results remains open to debate.

α-SMA constitutes one of the typical characteristics of myofibroblasts, a contractile type of fibroblast that appears later in the proliferative phase of the wound healing process [42–44]. We observed a decrease in the area of α-SMA positive cells normalized to 1000 cells under the low nutrition conditions and a more pronounced decrease under the low nutrition + LPS conditions, which was accompanied by a decrease in the contraction ability, thus indicating decreased myofibroblast differentiation. The impact of LPS on the expression of α-SMA by dermal fibroblasts has not previously been reported in the literature. However, a decreased α-SMA protein level has been observed in human cardiac fibroblasts following LPS treatment (1 µg/ml) [45].

We observed that the dermal fibroblasts produced the same amount of collagen type I under all the test conditions. The production of collagen type I by dermal fibroblasts was verified in this study via the application of two independent methods – quantification in a cell culture medium by means of ELISA and the immunocytochemistry staining of intracellular collagen. Another study demonstrated that the production of collagen type I and hyaluronic acid in dermal fibroblasts is dependent on growth factor and cytokine changes. The research team determined that neither IL6 nor IL8 exerted changes with concern to the production of either of the molecules compared to the control [46]. These findings correspond to our results, i.e. that the production of collagen by dermal fibroblasts is not affected by pro-inflammatory conditions modeled in the form of increased levels of LPS or IL6/IL8 in the culture media.

This study describes a number of new observations on the behavior of dermal fibroblasts via the analysis of the typical functions thereof under wound stress conditions. The results led to the conclusion (summarized in Fig. 8) that although both low nutrition and low nutrition + LPS conditions do not affect the naturally increasing metabolic activity of cells, this phenomenon is not accompanied by increasing proliferation, as evinced by the absence of a change in the cell number under either of the conditions. Further, low nutrition + LPS served to potentiate the migration and pro-inflammatory phenotype of the cells as evidenced by enhanced in vitro wound closure and the increased production of IL6, respectively. The potential of dermal fibroblasts to contract collagen hydrogel declined under the low nutrition conditions as the result of the low cell number. Moreover, the expression of α-SMA decreased under the low nutrition conditions as well as (to a greater extent) in the low nutrition + LPS conditions. The remodeling ability of the cells was influenced by the both wound conditions, as documented by the enhanced MMP2 and unchanged MMP9 activities. Finally, the production of collagen type I was not affected by either of the conditions. We believe that, taken overall, our results provide a comprehensive functional insight into the response of dermal fibroblasts to in vitro modeled wound stress conditions.

The unique benefit of this study concerns the provision of evidence that increased metabolic activity does not necessarily reflect cellular proliferation under particular stress conditions; rather it reflects the energy required to trigger a cellular response to alarming or changing conditions in general. Further, the study shows that even a single stress factor is able to retard the healing process, since the low nutrition conditions alone acted to reduce the cell proliferation and the contractile capacity and to enhance the production of MMPs by the dermal fibroblasts. The nutrition supply thus appears to be the one key factor with respect to the support of dermal fibroblasts in the healing process. Moreover, the pro-inflammatory behavior of dermal fibroblasts in the healing process should also be taken into account since they actively produce pro-inflammatory cytokines in the pro-inflammatory conditions of wounds. Last but not least, when discussed with literature the origin of dermal fibroblasts and the origin and the concentration of LPS are overlooked. These variables bring different results followed by different conclusions made from them. In conclusion, we can state that the differing experimental set ups simulating wound conditions in vitro led to inconsistent results making the generalization of dermal fibroblasts functions upon these conditions challenging.

Funding

The study was supported by project No. CZ.02.1.01/0.0/0.0/16_019/0000787 “Fig

hting INfectious Diseases”, awarded by the Ministry of Education, Youth and Sports of the Czech Republic and financed from the European Regional Development Fund; by project No. NU20-02-00368 awarded by Czech Health Research Council and by the Ministry of Health of the Czech Republic, by project GA UK No. 128417 awarded by the Charles University; and the Charles University Research Fund (Progres Q39).

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analysis or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

Author Contribution

Anna Zavadakova and Lucie Vistejnova contributed to the study conception and design. Anna Zavadakova and Pavla Tonarova performed the experiments and collected the data. Data analysis was performed by all authors. The first draft of the manuscript was written by Anna Zavadakova and all authors commented on previous versions of the manuscript. All authors have read and approved the final manuscript.

Data availability

The data that support the findings of this study are available from the corresponding author on reasonable request.

Ethics approval

The study followed the approval by the local ethics committee of the University Hospital in Pilsen, E. Benese 13, 305 99 Pilsen, Czech Republic, decision of November 5^th, 2015. Guidelines were followed as set out in the Declaration of Helsinki.

Consent to participate

Written informed consent was provided prior to intervention by the parents of the children who participated in the study.

Stunova A, Vistejnova L (2018) Dermal fibroblasts-A heterogeneous population with regulatory function in wound healing. Cytokine Growth Factor Rev 39:137–150. https://doi.org/10.1016/j.cytogfr.2018.01.003
Tracy LE, Minasian RA, Caterson EJ (2016) Extracellular Matrix and Dermal Fibroblast Function in the Healing Wound. Adv Wound Care 5:119–136. https://doi.org/10.1089/wound.2014.0561
Zgheib C, Xu J, Liechty KW (2014) Targeting Inflammatory Cytokines and Extracellular Matrix Composition to Promote Wound Regeneration. Adv Wound Care. https://doi.org/10.1089/wound.2013.0456
Caley MP, Martins VLC, O’Toole EA (2015) Metalloproteinases and Wound Healing. Adv Wound Care. https://doi.org/10.1089/wound.2014.0581
Martins VL, Caley M, O’Toole EA (2013) Matrix metalloproteinases and epidermal wound repair. Cell Tissue Res 351:255–268. https://doi.org/10.1007/s00441-012-1410-z
Desmouliere A, Geinoz A, Gabbiani F, Gabbiani G (1993) Transforming growth-factor-beta-1 induces alpha-smooth muscle actin expression in granulation-tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J Cell Biol 122:103–111. https://doi.org/10.1083/jcb.122.1.103
Greaves NS, Ashcroft KJ, Baguneid M, Bayat A (2013) Current understanding of molecular and cellular mechanisms in fibroplasia and angiogenesis during acute wound healing. J. Dermatol. Sci.
Hehenberger K, Kratz G, Hansson A, Brismar K (1998) Fibroblasts derived from human chronic diabetic wounds have a decreased proliferation rate, which is recovered by the addition of heparin. J Dermatol Sci. https://doi.org/10.1016/S0923-1811(97)00042-X
Wall IB, Moseley R, Baird DM, et al (2008) Fibroblast dysfunction is a key factor in the non-healing of chronic venous leg ulcers. J Invest Dermatol. https://doi.org/10.1038/jid.2008.114
Schwarz F, Jennewein M, Bubel M, et al (2013) Soft tissue fibroblasts from well healing and chronic human wounds show different rates of myofibroblasts in vitro. Mol Biol Rep. https://doi.org/10.1007/s11033-012-2223-6
Iocono JA, Colleran KR, Remick DG, et al (2000) Interleukin-8 levels and activity in delayed-healing human thermal wounds. Wound Repair Regen. https://doi.org/10.1046/j.1524-475X.2000.00216.x
Rolfe MW, Kunkel SL, Standiford TJ, et al (1991) Pulmonary fibroblast expression of interleukin-8: a model for alveolar macrophage-derived cytokine networking. Am J Respir Cell Mol Biol. https://doi.org/10.1165/ajrcmb/5.5.493
Xing Z, Jordana M, Braciak T, et al (1993) Lipopolysaccharide induces expression of granulocyte/macrophage colony-stimulating factor, interleukin-8, and interleukin-6 in human nasal, but not lung, fibroblasts: evidence for heterogeneity within the respiratory tract. Am J Respir Cell Mol Biol. https://doi.org/10.1165/ajrcmb/9.3.255
Xia Y, Pauza ME, Feng L, Lo D (1997) RelB regulation of chemokine expression modulates local inflammation. Am J Pathol
Basso FG, Soares DG, Pansani TN, et al (2015) Effect of LPS treatment on the viability and chemokine synthesis by epithelial cells and gingival fibroblasts. Arch Oral Biol. https://doi.org/10.1016/j.archoralbio.2015.04.010
Smith RS, Smith TJ, Blieden TM, Phipps RP (1997) Fibroblasts as sentinel cells. Synthesis of chemokines and regulation of inflammation. Am. J. Pathol.
Abbott A (2003) Biology’s new dimension. Nature
Gvirtz R, Ogen-shtern N, Cohen G (2020) Kinetic cytokine secretion profile of LPS-induced inflammation in the human skin organ culture. Pharmaceutics. https://doi.org/10.3390/pharmaceutics12040299
Crompton R, Williams H, Ansell D, et al (2016) Oestrogen promotes healing in a bacterial LPS model of delayed cutaneous wound repair. Lab Investig. https://doi.org/10.1038/labinvest.2015.160
Kostarnoy A V., Gancheva PG, Logunov DY, et al (2013) Topical bacterial lipopolysaccharide application affects inflammatory response and promotes wound healing. J Interf Cytokine Res. https://doi.org/10.1089/jir.2012.0108
Li WW, Carter MJ, Mashiach E, Guthrie SD (2017) Vascular assessment of wound healing: a clinical review. Int Wound J. https://doi.org/10.1111/iwj.12622
Liang CC, Park AY, Guan JL (2007) In vitro scratch assay: A convenient and inexpensive method for analysis of cell migration in vitro. Nat Protoc. https://doi.org/10.1038/nprot.2007.30
Cai XJ, Chen L, Li L, et al (2010) Adiponectin inhibits lipopolysaccharide-induced adventitial fibroblast migration and transition to myofibroblasts via AdipoR1-AMPK-iNOS pathway. Mol Endocrinol. https://doi.org/10.1210/me.2009-0128
Li X-J, Huang F-Z, Wan Y, et al (2018) Lipopolysaccharide Stimulated the Migration of NIH3T3 Cells Through a Positive Feedback Between β-Catenin and COX-2. Front Pharmacol. https://doi.org/10.3389/fphar.2018.01487
Lemons JMS, Coller HA, Feng XJ, et al (2010) Quiescent fibroblasts exhibit high metabolic activity. PLoS Biol. https://doi.org/10.1371/journal.pbio.1000514
Rampersad SN (2012) Multiple applications of alamar blue as an indicator of metabolic function and cellular health in cell viability bioassays. Sensors (Switzerland). https://doi.org/10.3390/s120912347
Eleftheriadis T, Liakopoulos V, Lawson B, et al (2011) Lipopolysaccharide and hypoxia significantly alters interleukin-8 and macrophage chemoattractant protein-1 production by human fibroblasts but not fibrosis related factors. Hippokratia
Xi Z De, Xie CY, Xi Y Bin (2016) Macrophage migration inhibitory factor enhances lipopolysaccharide-induced fibroblast proliferation by inducing toll-like receptor 4. BMC Musculoskelet Disord. https://doi.org/10.1186/s12891-016-0895-0
Ejiri H, Nomura T, Hasegawa M, et al (2015) Use of synthetic serum-free medium for culture of human dermal fibroblasts to establish an experimental system similar to living dermis. Cytotechnology. https://doi.org/10.1007/s10616-014-9709-0
He Z, Gao Y, Deng Y, et al (2012) Lipopolysaccharide induces lung fibroblast proliferation through toll-like receptor 4 signaling and the phosphoinositide3-kinase-Akt pathway. PLoS One. https://doi.org/10.1371/journal.pone.0035926
Zhang J, Wu L, Qu JM (2011) Inhibited proliferation of human lung fibroblasts by LPS is through IL-6 and IL-8 release. Cytokine. https://doi.org/10.1016/j.cyto.2011.02.018
Monsuur HN, Boink MA, Weijers EM, et al (2016) Methods to study differences in cell mobility during skin wound healing in vitro. J Biomech. https://doi.org/10.1016/j.jbiomech.2016.01.040
Grada A, Otero-Vinas M, Prieto-Castrillo F, et al (2017) Research Techniques Made Simple: Analysis of Collective Cell Migration Using the Wound Healing Assay. J. Invest. Dermatol.
Jin J, Sundararaj KP, Samuvel DJ, et al (2012) Different signaling mechanisms regulating IL-6 expression by LPS between gingival fibroblasts and mononuclear cells: Seeking the common target. Clin Immunol. https://doi.org/10.1016/j.clim.2012.01.019
Cho JS, Kang JH, Um JY, et al (2014) Lipopolysaccharide induces pro-inflammatory cytokines and mmp production via TLR4 in nasal polyp-derived fibroblast and organ culture. PLoS One. https://doi.org/10.1371/journal.pone.0090683
Mihara M, Moriya Y, Ohsugi Y (1996) IL-6-soluble IL-6 receptor complex inhibits the proliferation of dermal fibroblasts. Int J Immunopharmacol. https://doi.org/10.1016/0192-0561(95)00106-9
Wong Y, Sethu C, Louafi F, Hossain P (2011) Lipopolysaccharide regulation of toll-like receptor-4 and matrix metalloprotease-9 in human primary corneal fibroblasts. Investig Ophthalmol Vis Sci. https://doi.org/10.1167/iovs.10-5459
Howard EW, Crider BJ, Updike DL, et al (2012) MMP-2 expression by fibroblasts is suppressed by the myofibroblast phenotype. Exp Cell Res. https://doi.org/10.1016/j.yexcr.2012.03.007
Akkoc A, Kahraman MM, Vatansever A, et al (2016) Lipopolysaccharide (LPS) induces matrix metalloproteinase-2 and – 9 (MMP-2 and MMP-9) in bovine dermal fibroblasts. Pak Vet J
Lindner D, Zietsch C, Becher PM, et al (2012) Differential expression of matrix metalloproteases in human fibroblasts with different origins. Biochem Res Int. https://doi.org/10.1155/2012/875742
Burke JP, Cunningham MF, Watson RWG, et al (2010) Bacterial lipopolysaccharide promotes profibrotic activation of intestinal fibroblasts. Br J Surg. https://doi.org/10.1002/bjs.7045
Darby I, Skalli O, Gabbiani G (1990) Alpha-smooth muscle actin is transiently expressed by myofibroblasts during experimental wound-healing. Lab Investig 63:21–29
Tomasek JJ, Gabbiani G, Hinz B, et al (2002) Myofibroblasts and mechano: Regulation of connective tissue remodelling. Nat. Rev. Mol. Cell Biol.
Hinz B, Phan SH, Thannickal VJ, et al (2007) The myofibroblast: One function, multiple origins. Am J Pathol. https://doi.org/10.2353/ajpath.2007.070112
Bolívar S, Santana R, Ayala P, et al (2017) Lipopolysaccharide Activates Toll-Like Receptor 4 and Prevents Cardiac Fibroblast-to-Myofibroblast Differentiation. Cardiovasc Toxicol. https://doi.org/10.1007/s12012-017-9404-4
Kim MS, Song HJ, Lee SH, Lee CK (2014) Comparative study of various growth factors and cytokines on type I collagen and hyaluronan production in human dermal fibroblasts. J Cosmet Dermatol. https://doi.org/10.1111/jocd.12073

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Functional Responses of Dermal Fibroblasts to Low Nutrition and Pro-Inflammatory Stimuli Mimicking a Wound Environment in Vitro

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Abstract

Figures

1. Introduction

2. Material And Methods

2.1 Cell isolation and culture

2.2 In vitro simulation of wound conditions

2.3 Metabolic activity

2.4 Cell proliferation

2.5 Scratch wound assay

2.6 IL6, IL8 and collagen type I quantification

2.7 MMP2 and MMP9 quantification

2.8 α-SMA and collagen type I visualization and α-SMA quantification

2.9 NHDF contraction ability

2.10 Statistical analysis

3. Results

3.1 Metabolic activity is not affected by low nutrition and low nutrition + LPS

3.2 The dermal fibroblasts did not proliferate under either low nutrition or low nutrition + LPS conditions

3.3 LPS stimulated the migration of dermal fibroblasts

3.4 Low nutrition + LPS promoted the pro-inflammatory behavior of the dermal fibroblasts

3.6 Both the low nutrition and low nutrition + LPS conditions reduced the NHDF contraction ability

4. Discussion

5. Conclusion

Declarations

References

Competing Interests

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