The Mechanical Environment Modulates Intracellular Calcium Oscillation Activities of Myofibroblasts

Myofibroblast contraction is fundamental in the excessive tissue remodeling that is characteristic of fibrotic tissue contractures. Tissue remodeling during development of fibrosis leads to gradually increasing stiffness of the extracellular matrix. We propose that this increased stiffness positively feeds back on the contractile activities of myofibroblasts. We have previously shown that cycles of contraction directly correlate with periodic intracellular calcium oscillations in cultured myofibroblasts. We analyze cytosolic calcium dynamics using fluorescent calcium indicators to evaluate the possible impact of mechanical stress on myofibroblast contractile activity. To modulate extracellular mechanics, we seeded primary rat subcutaneous myofibroblasts on silicone substrates and into collagen gels of different elastic modulus. We modulated cell stress by cell growth on differently adhesive culture substrates, by restricting cell spreading area on micro-printed adhesive islands, and depolymerizing actin with Cytochalasin D. In general, calcium oscillation frequencies in myofibroblasts increased with increasing mechanical challenge. These results provide new insight on how changing mechanical conditions for myofibroblasts are encoded in calcium oscillations and possibly explain how reparative cells adapt their contractile behavior to the stresses occurring in normal and pathological tissue repair

Restricting cell size decreases [Ca²⁺]_i frequency.

<p>A) SCMF were grown on microcontact-printed FN square islets of 100–10,000 μm², immunostained for F-actin, and imaged by confocal microscopy. A composite was produced by stitching images from different cells on the same substrate, containing all square sizes. Scale bar = 500 μm. B) Cells spreading on FN islets of 2,500 and 4,900 μm² were immunostained for F-actin (red), vinculin (green), and FN (blue). Scale bar = 250 μm. C) Representative fluorescence ratios (Em₃₄₀/Em₃₈₀) were recorded over time on Fura-2-loaded cells, stimulated with increasing concentrations of endothelin-1 (ET-1). D) Distribution fits of [Ca²⁺]_i oscillation periods are displayed for cells grown on 2,500 and 4,900 μm² islands (n_exp = 18–25, n_cells = 24–29) and treated with 50 nM ET-1.</p

Increasing the E-modulus of silicone substrates increases [Ca²⁺]_i oscillation frequency.

<p>SCMF were cultured on FN-coated silicone substrates produced with E-moduli of 5, 15, and 50 kPa for 2 days. A) Cells were immunostained for α-SMA (red), F-actin (green) and nuclei (blue). Scale bar = 50 μm. B) Representative fluorescence ratios (Em₃₄₀/Em₃₈₀) were recorded over time on Fura-2-loaded cells of each stiffness group. C) The dominant periods of regular oscillations were determined and pooled into a histogram that was fitted following a generalized extreme value distribution. D) [Ca²⁺]_i period distribution fits of 5 kPa, 15 kPa and 50 kPa groups are displayed and maxima highlighted with dotted lines (n_exp≥14, n_cells≥44). E) Period distribution fit maxima were translated into oscillation frequency (peaks/min) and expressed as a function of the Young's E-modulus of silicone substrates.</p

Disrupting actin stress fibers decreases [Ca²⁺]_i oscillation frequency.

<p>SCMF were grown for 2 days on FN-coated coverslips. A) Cells fixed before (left panel) and 30 min after Cytochalasin D treatment were immunostained for F-actin (red), vinculin (green) and nuclei (blue). Scale bar = 50 μm. B) Representative fluorescence ratio (Em₃₄₀/Em₃₈₀) of a Fura-2-loaded cell over time is shown. Fluorescence was recorded for 15 min before and 15 min after 30 min treatment with Cytochalasin D (15 μM) or vehicle (DMSO) only (C). D) [Ca²⁺]_i oscillation period was calculated before Cytochalasin D treatment and plotted against the period after treatment for the same cell. Any point above the diagonal indicates a period decrease after addition of the drug (n_exp =9, n_cells = 25).</p

Decreasing cell adhesion decreases [Ca²⁺]_i oscillation frequency.

<p>SCMF were grown for 2 days on glass coverslips, coated with PLL at 5.0 μg/cm², 0.5 μg/cm², or with FN. A) Cells were immunostained for F-actin (red), vinculin (green) and nuclei (blue). Scale bar = 50 μm. B) Representative fluorescence ratios (Em₃₄₀/Em₃₈₀) were recorded over time on Fura-2-loaded cells. C) [Ca²⁺]_i period distribution fits are displayed and maxima highlighted with dotted lines (n_exp = 29–34, n_cells = 68-93). D) SCMF area (n = 3; mean±SD) and E) the length of vinculin-positive focal adhesions were quantified from fluorescence staining (n = 3; mean±SEM, *p≤0.05, ***p≤0.001), F) Period distribution fit maxima were translated into oscillation frequency (peaks/min) and expressed as a function of the mean SCMF focal adhesion lengths on differently adhesive substrates.</p

The E-modulus of collagen gels modulates [Ca²⁺]_i oscillation frequency.

<p>SCMF were grown in gels of 1.0, 1.5, 2.0, or 2.5 mg/ml collagen for 2 days. A) Confocal reflection microscopy imaging of cell-free gels demonstrates collagen fiber density. B) The Young's E-modulus of cell-free gels was measured using a micro-indentation approach. C) Representative fluorescence ratios (F/F₀) were recorded over time on Fluo-4-loaded cells grown in collagen gels. D) [Ca²⁺]_i period distribution fits are displayed and maxima highlighted with dotted lines (n_exp = 18–41, n_cells = 60–93). E) Period distribution fit maxima were translated into oscillation frequency (peaks/min) and expressed as a function of the Young's E-modulus of collagen gels. F) Myofibroblasts in collagen gels were stained after 2 days for F-actin (green) and collagen ECM was overlaid with confocal reflection imaging (red). Scale bars = 50 μm. The collagen density was measured by applying an integrated density function to confocal images of gels either in G) cell-free regions of the gels or H) in the vicinity of SCMF. I) [Ca²⁺]_i oscillatory frequencies are expressed as a function of measured collagen densities (Fig. 2I). (n = 3; mean±SD, **p≤0.01).</p

Prestress in the extracellular matrix sensitizes latent TGF-β1 for activation.

Integrin-mediated force application induces a conformational change in latent TGF-β1 that leads to the release of the active form of the growth factor from the extracellular matrix (ECM). Mechanical activation of TGF-β1 is currently understood as an acute process that depends on the contractile force of cells. However, we show that ECM remodeling, preceding the activation step, mechanically primes latent TGF-β1 akin to loading a mechanical spring. Cell-based assays and unique strain devices were used to produce a cell-derived ECM of controlled organization and prestrain. Mechanically conditioned ECM served as a substrate to measure the efficacy of TGF-β1 activation after cell contraction or direct force application using magnetic microbeads. The release of active TGF-β1 was always higher from prestrained ECM as compared with unorganized and/or relaxed ECM. The finding that ECM prestrain regulates the bioavailability of TGF-β1 is important to understand the context of diseases that involve excessive ECM remodeling, such as fibrosis or cancer