Wound healing-promoting effects stimulated by extracellular calcium and calcium-releasing nanoparticles on dermal fibroblasts

Ormoglass nanoparticles of oxide based composition CaO:P₂O₅:Na₂O:TiO₂ 44.5:44.5:6:5 were prepared by controlled hydrolysis sol-gel method at inert atmosphere as described previously [30]. The precursors used were prepared as follows. Calcium (Ca) and sodium (Na) 2-methoxyethoxides precursor solutions were prepared by refluxing metallic calcium (Sigma-Aldrich, 98%) and sodium (Panreac, 99%) respectively in anhydrous 2-methoxyethanol (Sigma-Aldrich, 99%) at 124 °C during 24 h. Phosphorus (P) ethoxide precursor solution was prepared by refluxing P₂O₅ (Sigma-Aldrich, 99.99%) in absolute ethanol (Panreac, 99.9%) at 78 °C during 24 h. Titanium (Ti) alkoxides precursor solution was prepared by diluting Ti-tetraisopropoxide (Alfa Aesar, 97%) in absolute ethanol. Then, Ti, Ca and Na precursors were added in a balloon with 1 h of vigorous stirring, followed by the addition of previously distilled 1,4-dioxane until a 5% v/v concentration was reached. The last precursor solution, P alkoxides precursor, was added at a control rate of 2.5 ml h⁻¹ at 4 °C with an infusion pump. Then a basic catalyzing aqueous solution with a molar relation 20:0.1:4 H₂O:NH₃:EtOH and a Ti:H₂O ratio of 1:60 was added at 1 ml h⁻¹ and 4 °C. The solution was aged in a closed vial at 70 °C for 4 d under vigorous stirring. After aging, the produced particles were washed with ethanol and centrifuged five times for 5 min at 1000 g. Finally, SG5 particles were manually grinded in an agate mortar, dried at 90 °C, and stored in a desiccator for further use.

The size of SG5 particles was assessed by dynamic light scattering (DLS) on a Malvern Instruments Zetasizer Nano-ZS instrument. The solution used was absolute ethanol containing 1% Tween^® 20 (Sigma) to avoid particle degradation and agglomeration. Before the measurement, the solution containing SG5 was homogenized in a bath sonicator and transferred into a quartz cuvette. SG5 morphology was observed with scanning electron microscopy (SEM) (NOVA NanoSEM 230, FEI Company, USA) uncoated on a silicon plate.

Ca, P, Na and Ti ion concentrations released by the particles in complete culture medium (CCM) consisting of DMEM (Gibco) supplemented with 10% fetal bovine serum (FBS) (Sigma-Aldrich), 2 mM L-glutamine (Invitrogen), 100 U ml⁻¹ penicillin (Gibco) and 100 U ml⁻¹ streptomycin (Gibco), were measured as follows. Particle suspensions of 1.2 mg ml⁻¹ were prepared and incubated at 5% (v/v) CO₂ and 37 °C. After 24 h, suspensions were centrifuged for 10 min at 1000 g. To achieve total degradation, particles were mixed with 5% HCl in the same proportion (w/v) as for the release and were incubated at 37 °C in an automatic shaker (Thermomixer Comfort, Eppendorf) for 24 h. The supernatant was diluted 1/10 in 1% in HNO₃ and the Ca, P, Na, Ti levels were measured by inductively coupled plasma optical emission spectrometry (Optima 3200RL, Perkin Elmer).

Primary cultures of rat dermal fibroblast were used in all the experiments. These cells were isolated from skin patches from the abdominal area of 10 Wistar rats of 2–4 weeks old. Rats were anesthetized in a chamber with 5% isofluorane and sacrificed in CO₂ saturated atmosphere. After shaving their abdomen, skin patches of 1 cm² were isolated and placed in sterile CCM. Then, patches were cut into pieces of 1 mm² and were transferred to empty 90 mm Petri dishes. Each dish contained the pieces of two patches separated by approximately 5 mm. Following 15 min, 15 ml of CCM was added per dish and cells from the explants were allowed to grow until 80% confluence. Then, cells were trypsinized (TrypLE ^TM Express Enzyme, Thermo Fisher Scientific), seeded into T75 Nunclon^TM flasks (Thermo Fisher Scientific), and allowed to expand until 80% confluence. This population of cells was highly enriched with rat skin fibroblasts. At this point, cells were trypsinized again, pooled, and kept in liquid nitrogen at passage 2 in a solution of 10% DMSO (Sigma-Aldrich) in FBS. All protocols concerning animal care were approved by the Committee on Ethics and Animal Experiments of the Scientific Park of Barcelona (Permit No. 0006S/13393/2011). When needed, cells were thawed from the frozen stock, grown at 37 °C in 5% (v/v) CO₂ in CCM, and used within passage 4–6. Medium was changed every 3 d.

Media containing calcium from CaCl₂ (Ca–CaCl₂) or the ion release from the SG5 particles (Ca-SG5) was prepared. Ca–CaCl₂ media was obtained by dissolving CaCl₂ (>96%, Sigma-Aldrich) in calcium-free DMEM to obtain a solution of approximately 8 mM Ca²⁺, incubating overnight, and filter sterilizing it (0.22 μm). Ca-SG5 media was prepared as follows. A particle suspension of 0.6 mg ml⁻¹ in calcium-free DMEM supplemented as regular medium was prepared and incubated for 24 h at 37 °C in 5% (v/v) CO₂. After incubation, the suspension was centrifuged for 10 min at 1000 g, and the supernatant was collected and sterile filtered (0.22 μm). Calcium released in the media was measured using the quantitative colorimetric method 0-cresolphtalein complexone (Sigma) [31], reading absorbance at 570 nm on the Infinite M200pro microplate reader (Tecan). Finally, media was diluted to prepare specific concentrations needed for the assays. To measure the pH of these samples, an aliquot was kept in the CO₂ incubator at 37 °C for 30 min, pH was quickly measured with a Laquatwin pHmeter (B-712, Horiba), and values were compared to a control sample without particle release. In all experiments, prior to addition of conditioned media, cells were rendered quiescent by an overnight incubation in low-serum (2% FBS) and 0.1 mM calcium-containing medium. Then, experimental media was added containing 2% or 10% FBS, depending on the restrictions or needs of each experiment.

Cell growth in conditioned media containing 10% FBS was studied by measuring metabolic activity and DNA quantification at days 1, 3 and 6. For metabolic activity determination, 2.5 × 10³ cells/well were seeded in 48-well plates. At each time point, Alamar Blue (Alamar Blue kit, Thermo Fisher Scientific) was added following manufacturer's instructions and fluorescence was read at Ex/Em wavelength of 530/590. Six replicate wells were used per condition. Increase of cell number overtime was assessed by DNA quantification with the Picogreen dsDNA Assay Kit (Invitrogen). Briefly, 1 × 10⁴ cells/well were seeded in 24-well plates. Four replicate wells were used per condition. At the above-mentioned time points, cells were lyzed in cold Tris-EDTA buffer (Sigma-Aldrich) by three freeze-thaw cycles and quantification of DNA was performed following manufacturer's instructions. Finally, fluorescence was read at Em/Ex 480/520. Each assay was assessed three times. In both assays, fluoresce was measured on the Infinite M200pro (Tecan) microplate reader.

To determine the effect on cell migration of the different conditioned media, a scratch-wounded fibroblast monolayer model was employed. Prior to cell seeding, a line passing through the middle of each well was drawn on the external side of the plate. Briefly, 4 × 10⁴ cells/well were seeded into 12-well tissue culture plates and cultured to confluence. Four replicate wells were used per condition. The confluent layer of fibroblasts was scrape-wounded using a sterile p200 pipette tip by one linear scratch perpendicular to the drawn line. Wells were washed with PBS to eliminate floating and dead cells, and conditioned media containing 2% FBS was added. Wound closure was monitored by collecting digitized images of the upper and lower area adjacent to the drawn line, immediately after the scratch and 24 h afterwards. Images were acquired with an inverted microscope (Nikon TE200) with a digital camera (Olympus DP72) using the 4× phase contrast objective lens. Reduction of wound areas was analyzed with the ImageJ 1.48i software. The experiment was repeated three times.

The activities of MMP2 and MMP9 of cell culture supernatants were measured by a gelatin zymogram protease assay. Cells (4 × 10⁴ cells/well) were seeded in 24-well plates. Following starvation, conditioned media (2% FBS) was added and, after 24 h, media was collected and centrifuged at 10 000 g for 10 min at 4 °C. Aliquots of supernatants were mixed with 3× non-reducing sample buffer (0.3% SDS; 15% 1M Tris-HCl pH 6.8, 30% glycerol and 0.075% bromophenol blue) and 30 μl were loaded on 8% SDS-polyacrylamide gels containing 0.1% gelatin (Gelatin from porcine skin BioReagent, Type A, Sigma). Then, prepared samples were subjected to electrophoresis under a constant voltage of 125 mV for 90 min. A protein marker (Kaleidoscope^TM Prestained SDS-PAGE Standards, Bio-Rad) was run in parallel for molecular weight identification. Following electrophoresis, gels were washed in 2% Triton X-100 (Sigma) for 30 min at room temperature to remove SDS. Gels were then incubated at 37 °C for 18 h in 1× Development buffer (10× Zymogram Development Buffer, Bio-Rad) and stained with 0.5% Coomassie Blue R250 (Bio-Rad) in 50% methanol and 10% glacial acetic acid for 1 d. After destaining, gelatinolytic activities were identified as clear bands against the blue background. To quantify the amount of gelatinase activity, gels were scanned on a densitograph (LAS4000 Imaging System) and the intensities of the digitalized bands were measured with ImageJ software. All density values were normalized to the intensity of the 0.1 mM sample and to the protein level of the cell extract, which was measured with the BCA protein assay (Thermo Fisher Scientific). The experiment was repeated three times including three replicates per condition. For each experiment, two gels were run including different samples.

Collagen secreted in media by fibroblasts cultured under the different conditioned media was measured using the Sircol Collagen Assay (Biocolor, UK). Cells (2.5 × 10⁵ cells/well) were seeded in T25 flasks with complete DMEM. After starvation, cells were treated with 3.5 ml/flask conditioned media containing 2% FBS for 24 h. Then, media from the flasks was transferred in 2 ml low protein binding microcentrifuge tube, and total collagen was quantified following manufacturer's instructions. Absorbance signal was measured at 555 nm on the Infinite M200pro (Tecan) microplate reader and values were normalized to the protein content from the cell extract measured with the BCA protein assay (Thermo Fisher Scientific). The experiment was carried out three times using triplicates per condition.

The stressed-relaxed FPCL model was used as previously described [32]. Briefly, collagen lattices were polymerized in 6-well tissue culture plates. Two hundred microliters of the mix of OptiCol^TM Rat Collagen Type I (Cell guidance systems) at a concentration of 1.2 mg ml⁻¹ containing 2 × 10⁴ rat dermal fibroblasts were carefully added in each well and allowed to gel for 1 h in a 5% (v/v) CO₂ humidified atmosphere at 37 °C. Then, 2.7 ml of the different conditioned media was added. Following 5 days of culture, gels were imaged using a Leica MZ16 F stereomicroscope. Then, gels were detached from the surface of the wells by rimming the lattice with a sterile spatula and images of the floating lattices were captured at 0.5, 1, 2 and 4 h. To quantify contraction, FPCL surface area was measured with ImageJ software and values were normalized to the area of the lattices before being released. The experiment was repeated three times including three replicates per condition.

The expression of alpha-smooth muscle actin (α-SMA) by fibroblasts incubated in the collagen lattices under different calcium concentrations was quantified by immunoblot analysis. After the last picture acquisition, each gel used in the FPCL experiment was washed in PBS and transferred into microcentrifuge tubes. Cell lysate was obtained by incubating the gels with the radioimmunoprecipitation lysis and extraction buffer supplemented with a protease inhibitor cocktail (Santa Cruz Biotechnology) for 20 min in ice. To ensure complete cell lysis, gels were also sonicated (Ultrasonic Processor UP50H) twice for 5 s at 100% potency in cold. Finally, samples were centrifuged at 10 000 g for 10 min at 4 °C and supernatants were transferred into clean microcentrifuge tubes. Equal sample loading into the electrophoresis was ensured by measuring DNA concentration of each sample with the Picogreen dsDNA Assay Kit (Invitrogen), and 40 μg per sample were mixed with 6× Laemmli buffer. Then, samples were run in SDS-polyacrylamide (10%) gels and transferred to nitrocellulose membranes (0.45 μm, Bio-Rad) by electrophorosesis. Membranes were blocked in 5% skim milk in Tris-buffered saline containing 0.1% Tween^® 20 (TTBS) at room temperature for 1 h, and they were sequentially probed with a goat anti-α-SMA polyclonal antibody (1:500 dilution, PA5-18292, Thermo Fisher Scientific) overnight at 4 °C, and then GAPDH (1:500, G8795, Sigma) for 1 h at RT. After three washes in TTBS, membranes were sequentially incubated with secondary antibodies conjugated with horseradish peroxidase: mouse anti-goat (1:1000 dilution, 2354, Santa Cruz Biotechnology) and donkey anti-mouse (1:3000 dilution, SA1-100, Thermo Fisher Scientific) for 1 h in 5% milk-TTBS, and finally developed by Clarity^TM Western ECL Substrate (Bio-Rad). The chemiluminescent signal was read with a densitograph (LAS4000 Imaging System), and the intensities of the digitalized bands were measured with ImageJ software. Density values were normalized to their correspondent GAPDH band. Three membranes were analyzed running samples from three different experiments.

Differential gene expression of cells treated with 0.1 and 3.5 mM calcium concentration from CaCl₂ and from the release of the particles was assessed using the RT² Profiler^TM PCR Array Rat Wound Healing (PARN-121ZC-2, Qiagen). Cells (8 × 10⁴ cells/well) seeded on 6-well plates were treated with conditioned media (10% FBS) for 24 h, and RNA was extracted using the RNeasy PlusMinikit (Qiagen) following the manufacturer's instructions. RNA (0.5 μg) of each condition were reversely transcribed with a RT² First Strand Kit (Qiagen) and the cDNA was used with the RT² SYBR Green Mastermix (Qiagen) in array plates containing a panel of 84 wound healing-related genes, 5 housekeeping (HK) control genes, a rat genomic contamination control and RT-controls. Quantitative RT-PCR was performed with the StepOnePlus RT-PCR system (Appleid Systems). Cycling parameters were 10 min at 95 °C (activation) followed by 40 cycles consisting of 15 s at 95 °C (denaturation) and 1 min at 60 °C (extension). The CT threshold for all the conditions was matched and mRNA expression for each gene was normalized to the HK genes of the array. Relative amounts of RNA for each gene and fold increase were calculated by using the 2 − ΔCT method or the 2 − ΔΔCT method. The genes selected were the ones with, at least, two-fold higher or lower expression than the control sample.

Data are presented as means ± standard deviation (S.D). The results were subjected to one or two-way ANOVA, and statistical differences between the groups were analyzed using post hoc Tukey's test at a significance level of 5%. The statistical software used was GraphPad Prism 6.0 (San Diego, CA, USA).

Particle size was measured with DLS and confirmed with SEM images. DLS indicated that the average size of SG5 particles was 357.3 ± 38.6 nm (figure 1(a)) and SEM images showed particles of this size (figure 1(b)). Because we are interested in studying the effect of the ions released by SG5, the ion profile of the release of the particles after 24 h in CCM was obtained (figure 1(c)). We also measured the ion release after complete degradation of the particles to compare how much of the ion content had been released. The measurement showed that, at the analyzed time point, all the calcium (Ca) and phosphorous (P) had been released from the particles, generating concentrations in the mM range. As expected by the theoretical oxide composition of the particles, the total concentration of P released doubled the total concentration of Ca. On the other hand, titanium (Ti) was only partially liberated and in much lower concentrations, within the μM range. Due to the high concentration of sodium (Na) in cell media, Na released from the particles could not be properly quantified, but measurement of the total release of Na indicated that the maximum that can be liberated is in the mM range.

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Cell viability and growth was studied by metabolic activity assessment and dsDNA quantification. A wide range of calcium concentrations, from 0.1 up to 7.5 mM, were used, and the effect was evaluated along 6 d. The lowest concentration used, to which the effect of the different concentrations were compared to, was 0.1 mM because it is optimal for keratinocyte growth while does not promote fibroblast proliferation [20]. In addition, two types of solutions were employed: media containing calcium from CaCl₂ (Ca–CaCl₂) or the ion release from the SG5 particles (Ca-SG5).

Metabolic activity was stimulated with specific calcium concentrations, and differences were detected between Ca–CaCl₂ and Ca-SG5 media. Stimulation by Ca–Cl₂ was observed on day 3 by the 2.5 and 3.5 mM conditions, but concentrations of 5 mM or higher maintained activity static or even decreased viability (figure 2(a)). On the other hand, stimulation by Ca-SG5 was detected on day 1 by the 3.5 mM-SG5 condition, but a significant decrease was observed on day 6.

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The increase in metabolic activity triggered by Ca–Cl₂ and Ca-SG5 at specific time points and concentrations did not correlate with an increased cell growth, as shown by total DNA quantification (figure 2(b)). Indeed, the different calcium concentrations tested did not augment cell growth compared to the lowest concentration used, 0.1 mM, at each time point. These results differ from previous studies which showed that mitogenic activity incremented upon calcium exposition [21, 22]. While these previous studies were performed in low-serum conditions, we used media containing 10% FBS because we wanted to analyze the effect of calcium for a long period of time and in a more physiological environment. Thus, we posit that the high content of growth factors and other proteins from serum in media might have masked the expected calcium-stimulated proliferative effect.

In addition, 2.5 mM and 3.5 mM Ca-SG5 showed a slower growth rate compared to their Ca–CaCl₂ counterpart on day 6 (figure 2(b)). This result, together with the previously mentioned on metabolic activity, indicate that long term exposure of fibroblasts to the SG5 release provoked cytotoxicity in the in vitro system. To discard pH effect, we measured the pH of the solutions containing SG5 release and found that the pH of these samples remained equivalent to non-conditioned media (data not shown). Therefore, one or more of the species liberated by the particles caused this effect. Since the method employed for the production of SG5, based on sol-gel without posterior high temperature treatment, favors the presence of non-decomposed organic compounds in the structure of the particles, when released, these could induce some degree of cytotoxicity. However, it should be considered that in an open system such as in an in vivo situation, the cytotoxic effect should diminish, as previously observed [28, 30, 33].

Fibroblast migration into the newly formed fibrin matrix at the wound site is an essential event for the progression of the healing. To test whether the different Ca²⁺ treatments affect fibroblast migration, a wound scratch assay was performed (figure 3(a)). The experiment was carried out in low-serum conditions and stopped after 24 h to reduce the effect of cell proliferation. Cell migration was evaluated by measuring the area of the initial wound covered by the cells at the final time point, as shown in figure 3(b).

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All calcium concentrations, from both Ca–CaCl₂ and Ca-SG5, significantly stimulated cell migration compared to the lowest concentration. In addition, the intensity of the effect seemed to be concentration-dependent and the maximum migration was reached using 2.5 mM Ca²⁺. Similar stimulation of cell migration was found when media with Ca-SG5 was used. Quantification of the increase in perimeter of the cell front also revealed that concentrations of 2.5 mM and above stimulated a more invasive migration behavior (figure 3(c)).

Other studies had shown migration stimulation with calcium on fibroblasts [34] but, to our knowledge, the concentration-dependent behavior had not been reported before. There is not a single proposed mechanism by which extracellular Ca²⁺ can affect cell migration due the plethora of events in which Ca²⁺ is implicated and the complexity of cell motility, but it can be linked to the activation of calcium-binding proteins. Studies have shown how increasing extracellular Ca²⁺ leads to increase in intracellular Ca²⁺ [35] that results in the activation of calcium-dependent proteins involved in cell migration, such as calpains [36].

During the remodeling and repair phase of the healing process, fibroblasts play an essential role in transforming the new matrix that is being created. Remodeling requires both degradation and synthesis of new ECM, but in chronic wounds the deposition of new tissue is prevented or impaired by numerous factors [37]. To study the effect that calcium and the ion release of SG5 have on the remodeling ability of fibroblasts, gelatinase activity and collagen synthesis were analyzed.

Gelatin zymography was used to assess MMP2 and MMP9 activity, two key players in ECM degradation. The results show that 3.5 mM Ca–CaCl₂ increased the active form of MMP2 compared to the other Ca²⁺ concentrations (figures 4(a) and (c)). An increase in MMP9 activity was also observed by incubating cells with 3.5 mM Ca–CaCl₂ (figures 4(a) and (b)). Increased expression of MMP2 and MMP9 by Ca²⁺ had been previously described [34], but the clear activation of MMP2 by specific calcium concentrations had not been reported. Interestingly, in the presence of 3.5 mM Ca-SG5, no increase of MMP activity was detected. The additional compounds released by the particles, such as phosphate, may cause this different behavior. This result may be beneficial in the context of chronic wound healing, since MMP2 and MMP9 seem to be abnormally expressed in these type of wounds [38] and are considered partly responsible for the chronic condition [37, 39].

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Stimulation of collagen synthesis by extracellular calcium and the ion release of SG5 was evaluated by quantifying total collagen secreted in the different test media. Ca²⁺ concentrations of 2.5 and 3.5 mM Ca–CaCl₂ significantly stimulated collagen synthesis, and a similar effect was observed in the Ca-SG5 condition (figure 4(d)). Although the stimulating effect of calcium on collagen synthesis by fibroblasts has been suggested by others [23, 40], by testing a wide range of calcium concentrations we were able to identify an optimal concentration to promote collagen production by dermal fibroblasts. The de novo deposition of collagen stimulated by both CaCl₂ and SG5 could contribute to an improved healing, since expression of the collagen gene in fibroblasts is suppressed in some types of chronic wounds [41].

Fibroblasts play a key feature in the latest events of the healing process by providing the contractile forces that bring the wound edges together. In our study, the stressed-relaxed FPCL model was used to quantify the contractile capacity of fibroblasts cultured with different Ca²⁺ concentrations and the ionic release of SG5.

Media containing Ca–Cl₂ and Ca-SG5 with equivalent calcium concentrations affected fibroblast's contractile capacity similarly (figure 5(a)). Fibroblasts cultured in lower Ca²⁺ concentrations (0.1 mM and 1.25 mM) were able to contract the collagen matrix faster and in greater extent compared to higher Ca²⁺ concentrations (2.5 and 3.5 mM). Differences observed among conditions showed a concentration-dependence behavior. Only for the highest calcium concentration (3.5 mM) of Ca-SG5, fibroblasts showed reduced contractile capacity compared to their equivalent Ca–CaCl₂ (figure 5(a)). One or more of the molecules liberated by SG5 is triggering these differences. Exposure to either the organic content or the high phosphate concentration released by the particles may be causing this effect.

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Since expression of α-SMA is known to increase fibroblast contractile activity, [45, 46] α-SMA expression levels were quantified to test whether the differences in contractile capacity could be originated due to differences in α-SMA expression. The immunoblot analysis performed showed correlation between α-SMA levels of expression and fibroblast contractile capacity, unveiling clear differences in stimulation among Ca²⁺ concentrations (figure 5(b)). At 0.1 mM the expression of α-SMA was the highest among the conditions tested, while at 2.5 and 3.5 mM the expression was significantly lower or even absent (figure 5(b)). SG5 conditioned media stimulated similar levels of expression as their equivalent Ca–CaCl₂ concentration (figure 5(b)). According to this result, extracellular Ca²⁺ would play a crucial role in fibroblasts activation to myofibroblasts. The study of the influence and control of extracellular Ca²⁺ concentration in the latest stages of the healing process as an strategy to avoid scar formation could be a new concept to explore in the field.

Expression of genes involved in wound healing by fibroblasts exposed to calcium was analyzed by Real-time transcriptase polymerase chain reaction (RT-PCR) using an array of 84 key genes central to the wound healing response. Cells were cultured for 24 h in 10% FBS media containing the low Ca²⁺ concentration used through the assays, 0.1 mM, or a concentration proved to stimulate many relevant effects, 3.5 mM, either from CaCl₂ or from the ionic release of SG5.

Our results indicate that 38 genes including ECM components, remodeling enzymes, cellular adhesion, cytoskeleton, inflammatory chemokines and cytokines, growth factors, and signal transduction factors were upregulated at least 2-fold in fibroblasts cultured with 3.5 mM Ca–CaCl₂ compared to 0.1 mM (figure 6(a)). Even though discrepancies may exist between mRNA and protein levels, the results obtained correlated with the previously exposed biological effects, and provided insight into possible mechanisms generating such effects. The increased expression of mRNA of proteins involved in cell migration in the presence of calcium, such as several integrin subunits and the cytoskeleton regulator Rhoa, was in line with the increased cell migration observed. We hypothesize that these proteins might be partly involved in the stimulation of cell migration reported. In addition, augmented expression of ECM components including different types of collagen as well as gelatinases MMP2 and MMP9 were detected, in accordance with the zymography assay. Even though we did not find increased cell proliferation when quantifying dsDNA, signal transduction factors involved in cell proliferation, such as mitogen-activated protein kinase 3 (MAPK3), were also increased. Thus, extracellular calcium might be triggering specific mitogenic pathways. In addition, increased expression of inflammatory cues, especially neutrophil-recruiting CXC chemokines, were identified, meaning that high extracellular Ca²⁺ could contribute to the inflammatory state of the wound by increasing expression of inflammatory chemokines and cytokines of, at least, fibroblasts.

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Similar results were observed in fibroblasts cultured with 3.5 mM Ca-SG5, for which the upregulated genes were 36 and only one gene was downregulated (Ccl12) (figure 6(a)). Nevertheless, by comparing the expression between 3.5 mM Ca–CaCl₂ and Ca-SG5 we found downregulation in the expression of inflammatory cytokines and chemokines in cells stimulated with SG5 (figure 6(b)). Again, exposure to either the organic content or the high phosphate concentration released by the particles may be causing these differences. Since stimulation of inflammatory cytokines in chronic wounds should be avoided [38, 42], the fact that less inflammatory cues are expressed with the ion release of SG5 than with calcium from Ca–Cl₂ makes SG5 more suitable for treating this type of wounds.

Overall, these findings suggest that our developed calcium-releasing particles may be used to effectively improve wound healing. However, due to the complexity and plethora of players involved in the wound healing process, a deeper understanding of the effect of the release should be acquired, especially on keratinocytes, inflammatory cells, and angiogenesis. In addition, trustworthy measurements of the levels of calcium in chronic wounds are needed in order to adjust the dose delivered to the optimal one, as it is known that its concentrations can fluctuate through the healing process [1, 43, 44].