Animal experiments were carried out with the approval of the local ethics committee for animal care (Landesamt für Natur, Umwelt und Verbraucherschutz Nordrhein-Westfalen, permit number 84-02.05.50.15.010).

PSCs were isolated from healthy wild-type C57BL/6J mice aged 8–12 weeks as described previously (Fels et al., 2016). Briefly, murine pancreata were isolated, and then enzymatically digested with 0.1% collagenase P (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) for 25 min at 37°C. After centrifugation, the homogenized tissue was resuspended in cell culture medium (DMEM/Ham F12 1:1, supplemented with 10% FCS and 1% penicillin/streptomycin) and seeded onto FCS-coated tissue culture dishes for 2 h. Afterwards, non-adherent cells were forcefully washed off the tissue culture plate, resulting in a homogeneous PSC culture. PSCs were used for experiments after two passages.

The pRK9 plasmid containing Piezo1-IRES-GFP (size 13,701 bp) was kindly provided by Prof. Gary R. Lewin's laboratory. The IRES promoter allows GFP to be transcribed only when Piezo1 has been transcribed, too. To create a suitable control plasmid, we removed the Piezo1 and IRES coding DNA from the plasmid. For this, we applied the restriction enzymes NotI and MscI (New England Biolabs, Ipswitch, MA, USA) to the Piezo1-IRES-GFP plasmid for 1 h at 37°C according to the manufacturer's instructions. Thus, the resulting plasmid was a structurally analogous control plasmid (size 5,411 bp) coding GFP but not the Piezo1 or IRES. Plasmid fragments were separated by gel electrophoresis on 1% agarose gel, then excised and purified using the QIAquick Gel Extraction Kit (Qiagen, Hilden, Germany). Following purification, DNA ends were blunted using the Klenow DNA Polymerase I, and finally, the 3′ and 5′ ends were ligated using T4 DNA ligase following the manufacturer's manual (New England Biolabs, Ipswitch, MA, USA).

A total of 25,000 HEK293 cells were seeded in glass-bottom dishes (Cell E&G, San Diego, USA) and after overnight incubation in 10% FCS-supplemented DMEM at 37°C, cells were transiently transfected with either the pRK9 Piezo1-IRES-GFP (HEK^Piezo1+) or the pRK9-GFP control plasmid (HEK^GFP+) using 1 μg plasmid DNA and 40 μl Lipofectamine® 3000 (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) in Opti-MEM reduced serum media (Thermo Fisher Scientific, Inc., Waltham, MA, USA). Twenty-four to forty-eight hours after transfection, HEK^Piezo1+ and HEK^GFP+ cells could be identified in the GFP channel of the fluorescence microscope (Visitron Systems, Puhheim, Germany) and subsequently used for Mn²⁺ quench experiments.

For immunocytochemistry, 30,000 PSCs were plated onto collagen I (Corning, New York, NY, USA) coated glass coverslips and cultured for 48 h. Following incubation, PSCs were treated with 5 μM Yoda1 (Tocris Bioscience, Bristol, UK) or solvent (0.2% DMSO) for 1 h. To monitor the effects of blebbistatin, cells were additionally treated with 5 μM blebbistatin (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) for 15 min.

For visualizing Piezo1 membrane expression, live cell staining was performed as described previously by our group (Waschk et al., 2011; Storck et al., 2017). First, cells were blocked with Ringer's solution (122.5 mM NaCl, 5.4 mM KCl, 1.2 mM CaCl₂, 0.8 mM MgCl₂, 5.5 mM d-glucose and 10.0 mM HEPES, titrated to pH 7.4) supplemented with 1% bovine serum albumin (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) for 1 h on ice. This was followed by staining with the primary antibody against mPiezo1 (Proteintech, Manchester, UK, #15939-1-AP, 1:200) for 2 h at 4°C. This antibody recognizes an extracellular epitope of the channel protein so that cell permeabilization was not necessary. After washing five times, coverslips were fixed in 3.5% paraformaldehyde for 20 min at 4°C. Afterwards, Alexa 488-conjugated secondary antibody (Invitrogen, Carlsbad, CA, USA, 1:500) was applied for 1 h at room temperature. Finally, after washing 3 times with PBS, 0.1% DAPI (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) was applied in DAKO mounting solution (DAKO Deutschland GmbH, Hamburg, Germany) and coverslips were mounted onto slides.

For other immunofluorescence assays, fixation was performed on ice using 3.5% paraformaldehyde for 1 h, followed by washing with PBS 3 times. Cells were subsequently permeabilized using 0.25% Triton-X-100 in PBS, then blocked using 10% goat serum for 1 h at room temperature. Primary antibodies against αSMA (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany, #A2547, 1:200), phosphorylated myosin light chain 9 (P-MYL-9; Invitrogen, Carlsbad, CA, USA, 1:100), and non-muscle myosin IIa (NMIIA) (Novus Biologicals, Littleton, CO, USA, 1:100) were applied overnight at 4°C. After washing three times with PBS, Alexa 488-conjugated secondary antibodies against mouse (Invitrogen, Carlsbad, CA, USA, 1:500) and rabbit (Invitrogen, Carlsbad, CA, USA, 1:500) were applied for 1 h at room temperature. Lastly, after washing three times with PBS, 0.1% DAPI was applied in DAKO mounting solution and coverslips were mounted onto slides.

Fluorescent images were acquired using a Zeiss Axiovert 200 inverted fluorescence microscope (Zeiss, Oberkochen, Germany) at 40× or 100× magnification. Quantification of Piezo1 channel density was performed manually by counting bright spots representing channels in 10 rectangular regions of 7.6 μm × 7.6 μm per cell and subtracting the mean number of spots in equivalent extracellular regions. For P-MYL9 and NMIIA immunocytochemistry fluorescence ratios of P-MYL9/NMIIA were assessed in an unpaired manner, as both primary antibodies originate from the same species (rabbit). First, the total cellular P-MYL9 and NMIIA intensities of 30 cells were measured using similar exposure and illumination settings. Afterwards, the amount of P-MYL9 was normalized to the total NMIIA of a randomly assigned cell. To evaluate the effects of blebbistatin, a scoring system was used to measure the stress fibers, the dendrite-like processes and cellular concavities (Figure 1).

Western blots of Piezo1, GAPDH, MYL2, and phospho-MYL2 (P-MYL2) were performed as described previously (Bulk et al., 2017). Total protein of PSCs was extracted using radioimmunoprecipitation assay (RIPA) buffer [50 mmol/l Tris, 150 mmol/l NaCl, 0.1% SDS, 0.5% sodium deoxycholate, 1% NP-40, and 1% Complete Mini protease inhibitor (Roche, Mannheim, Germany)]. Protein concentration of the samples was determined with Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, Inc., Waltham, MA, USA). Fifteen microgram of denatured total cellular protein was applied to each lane of a 15% polyacrylamide gel for electrophoresis at 80 mV. After overnight transfer to PVDF membranes at 4°C, we blocked the membrane with PBS containing 5% skim milk for 1 h, then incubated the blots with primary antibodies against mPiezo1 (Proteintech, Manchester, UK, #15939-1-AP, 1:100), P-MYL2 (1:500, MYL-Ser19 mouse mAb, #3675 Cell Signaling Technology, Danvers, Ma, USA), or MYL2 (1:500, MYL2 rabbit Ab, #3672 Cell Signaling Technology, Danvers, Ma, USA), and for housekeeping control GAPDH (1:2,000, GAPDH mouse mAb, #ab125247, Abcam, Cambridge, UK) overnight at 4°C. After washing three times with PBS, we applied HRP-conjugated goat-anti rabbit (1:10,000, Goat Anti-Rabbit IgG H&L, #ab6721, Abcam, Cambridge, UK) in case of Piezo1 and P-MYL2; and goat anti mouse secondary antibody (1:10,000, Goat Anti-Mouse IgG H&L, #ab6708, Abcam, Cambridge, UK) in case of MYL2. Chemiluminescence was detected using a commercial detection system (Chemidoc MP, Bio-Rad, Hercules, CA, USA), and band intensities were evaluated with ImageJ.

We applied the Mn²⁺ quench technique to monitor calcium influx into the cells (Merritt et al., 1989; Fels et al., 2016; Nielsen et al., 2017). Mn²⁺ largely mimics Ca²⁺ as it enters cells via similar pathways. In contrast to Ca²⁺ ions, Mn²⁺ ions quench the fluorescence emission of the calcium sensitive dye Fura-2. These experiments are performed at the Ca²⁺ insensitive, isosbestic excitation wavelength of Fura-2. The Mn²⁺-induced drop of the Fura-2 fluorescence is largely proportional to the transmembrane influx of Ca²⁺.

After staining the cells with the Ca²⁺-sensing dye Fura-2-AM (#F1221, Thermo Fisher Scientific, Inc., Waltham, MA, USA; 6 μM) for 20 min at 37°C in HEPES buffered Ringer's solution, cells were visualized using a ionic imaging setup consisting of a fluorescence microscope, a high-speed shutter and a polychromator (Visitron Systems, Puchheim, Germany). The proper isosbestic excitation wavelength, at which the emission is Ca²⁺ independent, was determined using the Ca²⁺ ionophore ionomycin (not shown). Thereafter, measurements were conducted at an excitation wavelength of 357 nm and fluorescence emission was recorded at 510 nm. Images were acquired in 10 s intervals. The experimental protocol started with an initial 3 min incubation in Ringer's solution. This was followed by another 3 min incubation in a Ca²⁺-containing, Mn²⁺ supplemented Ringer's solution for the experiments depicted in Figures 2C, 5A. For all other measurements we used a Ca²⁺-free, Mn²⁺ supplemented Ringer's solution (Mn²⁺ Ringer's). The Mn²⁺ concentration in Mn²⁺ Ringer's was 400 μM in case of PSCs and 1 mM in case of HEK293 cells. Also, in HEK293 cells, GFP caused negligible interference with the Fura-2 signals (not shown).

Data analysis was performed by measuring fluorescence intensities over the whole cell area and correcting it for background fluorescence. The extracted fluorescence intensities F were normalized to the initial fluorescence intensities F₀ determined under control conditions in the presence of Ringer's solution (F/F₀). For each cell, slopes of linear regression (ΔF/F₀/min) were calculated before and after Mn²⁺ application during intervals of 30 s. Subsequently, the slope after Mn²⁺ application was subtracted by the slope in the presence of Ringer's solution to correct for potential photobleaching. Lastly, for easier interpretation, the inverse value of the Mn²⁺ quench was determined which directly correlates with Ca²⁺ influx.

We assessed the intracellular pH (pH_i) of HEK293 cells using the fluorescent pH indicator BCECF-AM. Prior to the experiment, HCO3--buffered cell culture medium was exchanged for the HEPES-buffered Ringer's solution for 15 min to equilibrate pH_i at 37°C. BCECF staining was performed using 1 μM BCECF-AM for 3 min in Ringer's solution (pH 7.4) at 37°C. Afterwards, cells were washed and continuously superfused with Ringer's solution for 3 min, followed by 3 min superfusion of acidified Ringer's solution (pH 6.6) or Ringer's solution supplemented with 30 mM sodium propionate (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany). To maintain osmolarity the latter replaced 30 mM NaCl. Lastly, for calibration purposes, cells were permeabilized for protons using 1 μM nigericin (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) and superfused with a modified Ringer's solution (KCl 125 mM, MgCl₂ 1 mM, CaCl₂ 1 mM, and HEPES 20 mM) titrated to pH 7.5 and pH 6.5. Data were acquired with the same ionic imaging setup used for Ca²⁺ influx measurements measuring with dual excitation wavelengths of 440 and 490 nm and emission wavelength of 510 nm.

During data analysis, fluorescence intensities were measured over the whole cell area and corrected for background fluorescence. Subsequently, the 440 nm_ex/490 nm_ex fluorescence intensity ratios were calculated. As BCECF fluorescence ratio follows a linear trend between pH 6.5 and pH 7.5, two-point calibration was performed using linear regression.

Migration of PSCs was monitored using time-lapse video microscopy as described previously (Schwab et al., 2005; Fels et al., 2016). PSCs in cell culture medium were seeded in pre-coated 12.5 cm² dishes, Here, the coating matrix mimics the desmoplastic stroma and contained the following: 40 μg/ml laminin (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany), 40 μg/ml fibronectin (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany), 800 μg/ml collagen I (Corning, New York, NY, USA), 12 μg/ml collagen III (Corning, New York, NY, USA), and 5.4 μg/ml collagen IV (BD Biosciences, Heidelberg, Germany). Cell migration was recorded in temperature-controlled chambers (37°C) with CMOS cameras for 12 h at 5 min intervals using the MicroCamLab 3.1 software (Bresser, Rhede, Germany). Quantitative analysis of single cell migration was based on the segmentation of PSC contours using the Amira software suite (Thermo Fisher Scientific, Inc., Waltham, MA, USA). From the segmented cell contours, cell velocity (μm/min) and translocation were calculated as described before (Dieterich et al., 2008). Migration velocity was defined as the displacement of the cell centroid as a function of time. Mean translocation represents the net distance covered during the entire experiment.

For spheroid formation, 5,000 PSCs were inserted into a hanging drop of 40 μL containing 0.31% methylcellulose (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) in cell culture medium. Spheroids formed spontaneously within 48 h. They were transferred into a modified matrix suitable for spheroid embedding containing 75 μg/ml laminin, 75 μg/ml fibronectin, 4.3 mg/ml collagen I, 27 μg/ml collagen III, and 12 μg/ml collagen IV. The matrix also contained 1 × 1,010 microbeads per ml (FluoSpheres 1 μm, Invitrogen, Carlsbad, CA, USA) to monitor matrix traction. Eighty microliter of matrix containing a spheroid was allowed to polymerize in 12.5 cm² tissue culture dishes for 2 h at 37°C in 5% CO₂, and subsequently 3 mL cell culture medium was added to the dishes supplemented with 50 ng/ml platelet-derived growth factor (PDGF). The experiments started after an equilibration period of another 2 h at 37°C in a 5% CO₂/air atmosphere.

The fate of PSC spheroids in the three-dimensional matrix was monitored using time-lapse video microscopy for a total of 48 h, similarly to the single cell migration assay described above. As a readout of spheroid invasion, the equatorial cross-sectional area of a given spheroid after 24 h was divided by the initial spheroid area at t = 0 h. The number of cells detached from the spheroid after 24 h was counted. Spheroids apply a remarkable amount of traction toward the matrix, as visible in Video 1. The traction of the matrix was evaluated by quantifying the movement of 10 beads within a maximal radius of 500 μm neighboring the spheroid during a 12 h period. The mean velocity of the beads (μm/h) is a surrogate of the traction of the spheroid toward the surrounding matrix.

PSC spheroids were incubated for 24 h at pH 7.4 and pH 6.6 in the presence of 20 μM Yoda1 or vehicle (0.1% DMSO), respectively. Afterwards, the collagen matrix containing spheroids was scraped off the dishes using a scalpel. Spheroids in the collagen matrix were fixed in 4% paraformaldehyde, 15% saturated picric acid in 100 mM phosphate buffer overnight at 4°C, dehydrated via ascending ethanol series, and embedded in paraffin. For histology, thin sections (5–7 μm) were cut with a rotary microtome (Leica RM 2135), rehydrated, and a routine hematoxylin and eosin stain (HE) was performed. The hematoxylin stains cell nuclei in a blue color, whereas eosin stains the ECM as well as the cytoplasm pink.

To evaluate the consistence of PSC spheroids, first, total cross-sectional area of spheroids was measured in multiple sections of each spheroid. Afterwards, by thresholding according to Li (Li and Tam, 1998), the acellular area within each spheroid could be measured in ImageJ. Spheroid fragmentation was calculated by dividing the acellular spheroid area by total spheroid area.

MTT assays were performed with matrix-embedded PSC spheroids cultured at pH 7.4 and pH 6.6 with 20 μM Yoda1 or vehicle (0.1% DMSO), respectively, for 24 h. First, spheroids were transferred to a 96-well plate, medium was exchanged for one containing 0.5 mg/ml MTT reagent (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany). Spheroids were incubated at 37°C in 5% CO₂ for 2 h. Then medium was discarded and 100 μL DMSO were added in order to dissolve formazan crystals. Absorbance was measured at 546 nm and corrected for absorbance at 650 nm using a multi-well spectrophotometer (Beckman Coulter, Brea, CA, USA). MTT reduction was normalized to absorbance of spheroids cultured at pH 7.4 with vehicle.

Annexin V staining was performed with trypsin-digested PSC spheroids according to manufacturer's instructions. Briefly, PSC spheroids were extracted from the collagen gels manually, washed with PBS and then inserted into tubes containing 0.25% Trypsin-EDTA solution (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany) and shaken at 250 rpm at 37°C for 10 min. After centrifugation at 200 g for 5 min, cells were resuspended in Annexin V binding buffer (Mybiosource, San Diego, CA, USA) and stained with Annexin V conjugated with Alexa 555 (A35108, Invitrogen, Carlsbad, CA, USA, 1:20) at RT for 15 min. Lastly, cells were washed with Annexin V binding buffer and visualized using a fluorescence microscope at 10× magnification in the red fluorescence channel and DIC (Zeiss Axio Observer, Zeiss, Oberkochen, Germany). Annexin V positivity was evaluated by manual thresholding in ImageJ for 3 visual fields per spheroid. Annexin V positive cells were counted and summed for all visual fields and divided by the total number of cells counted in the respective DIC images.

Data are presented as mean values ± S.E.M and following test for normality unpaired Student's t-tests or one-way ANOVA were performed with Tukey's post-hoc test where applicable using Graphpad Prism 7. Statistical significance was assumed when p < 0.05.

We performed Western blot to investigate Piezo1 expression in whole-cell lysates of cell populations investigated in our study: namely in untransfected HEK293 cells, in HEK293 cells transiently transfected with control plasmid or Piezo1 and in PSCs (Figure 2A). We found that Piezo1 is abundantly expressed in PSCs, whereas overall Piezo1 was scarcely expressed in both untransfected and transiently transfected HEK293 cells. To visualize membrane expression of Piezo1 in PSCs, we labeled the channel using immunocytochemistry (Figure 2B). Piezo1 is expressed homogeneously in the cell membrane at a density of 0.52 ± 0.08 channels/μm² (n = 300 regions of 30 cells, N = 3 experiments). Moreover, to assess functionality of the channels, we measured Piezo1-mediated Ca²⁺ influx using the Mn²⁺ quench technique in the presence or absence of the Piezo1 agonist, Yoda1. Here, we found that 20 μM Yoda1 almost doubles Ca²⁺ influx into PSCs, from 0.29 ± 0.02 to 0.52 ± 0.03 ΔF/F₀/min (p < 0.0001, N = 3 experiments with n = 72 PSCs) (Figure 2C).

To validate the specificity of the Yoda1-induced Mn²⁺ quench assay, we compared Ca²⁺ influx into untransfected HEK293 cells with that into HEK293 cells transfected with a control plasmid or with Piezo1 (Figure 2E). As overall Piezo1 expression in the transiently transfected HEK293 cell population is low, we only measured Mn²⁺ quench in transfected GFP⁺ HEK293 cells (Figure 2D). GFP intensities of GFP⁺ control (1,724 ± 79, n = 523) and Piezo1 transfection (2,266 ± 61, n = 816) were proportionately increased compared to wild-type HEK293 autofluorescence with intensities < 100 (Figure 2D). Untransfected and control transfected HEK293 cells exhibit the same (p = 0.97) Yoda1-induced Mn²⁺ quench rate: 0.27 ± 0.02 ΔF/F₀/min (n = 50, N = 3) and 0.28 ± 0.02 ΔF/F₀/min (n = 82, N = 3), respectively (Figure 2E). In contrast, Piezo1-transfection leads to a 2-fold higher Mn²⁺ quench rate in HEK293 cells (0.55 ± 0.02 ΔF/F₀/min, n = 106, N = 3). Interestingly and in line with the Western blot results shown in Figure 2A, we needed a considerably higher Mn²⁺ concentration for HEK293 cells than for PSCs to elicit comparable quench rates. Piezo1-mediated Ca²⁺ influx is distinctly amplified using 5 μM Yoda1 (0.81 ± 0.03 ΔF/F₀/min, n = 60, N = 3) compared to solvent (0.025% DMSO) superfusion (Figure 2E). Collectively, these results provide evidence that PSCs express Piezo1 channels and that their function can be assessed with the Mn²⁺ quench technique.

Aiming for a readout of the physiological functions of Piezo1 in PSCs, we investigated whether cellular migration is altered by the Piezo1 activator Yoda1. Indeed, 5 μM Yoda1 stimulate the migration of PSCs seeded on a matrix resembling the desmoplastic matrix found in the PDAC stroma (Figure 3A). Velocity and translocation rise from 0.25 ± 0.01 μm/min and 20.4 ± 2.2 μm (n = 83, N = 3) under control conditions (0.075% DMSO) to 0.29 ± 0.01 μm/min (p = 0.0002) and 41.2 ± 4.3 μm (p = 0.0089, n = 52, N = 3), respectively. The fact that the increase of translocation is more pronounced than that of the velocity indicates a Yoda1-mediated increase in directionality (Figure 3B). Further increasing the concentration of Yoda1 to 15 μM lessens the stimulatory effect on PSC migration (velocity 0.27 ± 0.01 μm/min, p = 0.04; translocation 32.5 ± 3.6 μm, p = 0.62; n = 47, N = 3 experiments). In the presence of 50 μM Yoda1, PSCs are no longer accelerated (velocity 0.22 ± 0.01 and translocation 17.0 ± 1.4 μm; n = 36, N = 3) when compared to equivalent vehicle-treated PSCs (0.25 % DMSO; velocity 0.25 ± 0.02 μm/min, p = 0.09; translocation 18.6 ± 1.9 μm, p = 0.48; n = 45, N = 3; Figure 3B).

We next tested whether the Piezo1 activator Yoda1 also affects the migratory behavior of PSCs in a more physiological three-dimensional environment (Figure 3C). To this end we monitored the emigration of PSCs from a spheroid placed in a 3D desmoplastic collagen matrix. The normalized equatorial cross sectional area of PSC spheroids after 24 h increases similarly under control conditions (0.1% DMSO; by 3.5 ± 0.2-fold, n = 51 spheroids; N = 8) and in the presence of 20 μM Yoda1 (by 3.0 ± 0.2-fold, n = 54 spheroids, N = 8; p = 0.06). However, 20 μM Yoda1 lead to the detachment of more cells from the spheroid (11.0 ± 1.3 cells, n = 45 spheroids; N = 3) compared to control cells (6.5 ± 0.7 cells, n = 40 spheroids; N = 3) (Figure 3C).

When monitoring PSC emigration from spheroids with live-cell imaging, it was conspicuous that the surrounding matrix was pulled toward the spheroids. We quantified this phenomenon by tracking the movement of microbeads incorporated into the matrix (Figure 3D). As judged from the bead movement, spheroids pull on the matrix more vigorously in the presence of 20 μM Yoda1 (11.0 ± 0.4 μm/h, n = 105 beads, N = 3) compared to DMSO-treated spheroids (9.5 ± 0.4 μm/h, n = 90 beads, N = 3) (Video 1).

In order to assess how extracellular and/or intracellular acidification affect Piezo1 function, Piezo1-transfected HEK293 cells were treated with pH 6.6 Ringer's solution (n = 64, N = 3) and/or with 30 mM sodium propionate (n = 68, N = 3), respectively (Figure 4A). Our pH_i measurements indicate that 30 mM sodium propionate acidify HEK293 cells from pH_i 7.24 to pH_i 6.77 (ΔpH_i=-0.47±0.01) within 1 min. pH_i then reaches a steady state. In contrast, pH_i acidification occurs much more slowly upon treatment with pH 6.6 Ringer's solution, as pH_i only drops by 0.1 ± 0.002/min. Thus, when assessing Piezo1 function by means of the Mn²⁺ quench technique at t = 1 min after the start of the acidification stimulus, we can clearly distinguish between the effect of an intracellular or extracellular acidification.

Acidification of pH_e decreases Ca²⁺ influx into Piezo1-transfected HEK293 cells (Figure 4B). The Mn²⁺ quench rate falls from 0.55 ± 0.02 under control conditions (pH_e7.4) to 0.42 ± 0.01 ΔF/F₀/min (n = 196, N = 5; p < 0.0001). The acidification of pH_i by sodium propionate elicits a similar effect: 0.45 ± 0.02 ΔF/F₀/min (n = 81, N = 3; p < 0.0001). These effects are additive in Piezo1-transfected HEK293 cells, so that the most pronounced inhibition of Ca²⁺ influx occurs upon dual acidification of the intra- and extracellular space by co-applying pH 6.6 Ringer's solution with 30 mM sodium propionate: 0.31 ± 0.01 ΔF/F₀/min, n = 77, N = 3; p < 0.0001). This combined acidification diminishes the Mn²⁺ quench to a rate that is only 20% higher than that of control plasmid-transfected HEK293 cells (Figure 4D).

When Piezo1-transfected HEK293 cells are treated with 5 μM Yoda1, pH_e acidification (0.66 ± 0.02 ΔF/F₀/min, n = 151, N = 4) as well as pH_i acidification (0.5 ± 0.02 ΔF/F₀/min, n = 129, N = 4) still impair Ca²⁺ influx (Figure 4C). Interestingly, in case of Yoda1 stimulation, intracellular acidification is more effective in inhibiting Ca²⁺ influx than extracellular acidification (p < 0.0001), and there appears to be no additional effect of dual acidification (0.48 ± 0.04 ΔF/F₀/min, n = 34, N = 2).

As it is not known whether Piezo1 is functionally coupled to cytoskeletal tethers in PSCs, we aimed to investigate this by disrupting the actomyosin architecture using the myosin-inhibitor blebbistatin. Upon pre-incubating PSCs with 5 μM blebbistatin for only 15 min, 5 μM of the Piezo1 activator Yoda1 leads to an increase of only 28% in Mn²⁺ quench (0.37 ± 0.02 ΔF/F₀/min, n = 175, N = 3) (Figure 5A). This effect is not due to decreased membrane recruitment of the channel. Immunofluorescence staining of Piezo1 in the plasma membrane showed that the channel density after 5 μM blebbistatin treatment (0.47 ± 0.01 channels/μm) does not differ from that of untreated PSCs (p = 0.91, Figure 5B). Blebbistatin also causes substantial morphological changes in PSCs, that we evaluated by the criteria shown in Figure 1. Compared to control incubation with 0.1% DMSO (membrane concavities: 1.3 ± 0.1; stress fibers: 2.7 ± 0.1; dendrite-like protrusions: 1.3 ± 0.1; n = 30, from N = 3), PSCs treated with blebbistatin have more membrane concavities (2.7 ± 0.1) and denrite-like protrusions (2.5 ± 0.1), but less stress fibers (1.5 ± 0.1; n = 30, N = 3) (Figure 5C).

To elucidate whether Piezo1 modulates cellular contractility, we measured the phosphorylation status of the regulatory myosin light chain MYL9, which is known to have a regulatory role in smooth muscle cells (Figure 6A). We performed immunocytochemistry experiments and normalized the staining of P-MYL9 to that of the non-muscle myosin NMIIA. The treatment with 5 μM Yoda1 increased the ratio of P-MYL9/NMIIA from 0.90 ± 0.03 (vehicle-treated cells) to 1.31 ± 0.04 (n = 90, N = 3 for both conditions; p < 0.0001). For comparison, we also measured the phosphorylation status of MYL2, another regulatory myosin light chain that exerts its main function in cardiac muscle. The Western blot analysis comparing the amount of P-MYL2 to MYL2 in PSC cell lysates shows that the phosphorylation of MYL2 is not significantly different in Yoda1-treated cells (1.4 ± 0.4, n = 5, N = 5 control lysates vs. 3.2 ± 1.2, n = 4, N = 4 lysates from Yoda1-treated PSCs; N = 5 each; p = 0.17) (Figure 6B).

To get a better understanding of how PSC physiology is affected in the acidic tumor core, we inspected how changes in intra- and extracellular pH affect Piezo1-elicited responses of PSCs. Our results show that Mn²⁺ entry rate in case of pH_e acidification alone (0.23 ± 0.02 ΔF/F₀/min, n = 57, N = 5) remains analogous to the control pH 7.4 Ringer's perfusion (0.24 ± 0.02 ΔF/F₀/min, n = 48, N = 5; p = 0.99) (Figure 7A). On the contrary, intracellular acidification decreases Mn²⁺ entry by 40% (0.14 ± 0.01 ΔF/F₀/min, n = 45, N = 5; p = 0.003) similarly to combined intra- and extracellular acidification (0.16 ± 0.01 ΔF/F₀/min, n = 46, N = 5; p = 0.03). Thus, Ca²⁺ influx and thereby Piezo1 function appears to be predominantly inhibited by an intracellular acidification of PSCs.

As PSCs require Ca²⁺ to exert ECM traction, we expected that inhibited Ca²⁺ influx in acidic conditions would impair bead traction in PSC spheroids (Figure 7B). Interestingly, it turned out bead displacement toward PSC spheroids at pH_e6.6 (9.5 ± 0.4 μm/h, n = 57, N = 5) is comparable to the displacement at pH_e7.4 (11.0 ± 0.4 μm/h, n = 108, N = 5). Even more intriguingly, in the presence of 20 μM Yoda1 at pH_e6.6, bead displacement is decreased by 40% (5.8 ± 0.3 μm/h, n = 37, N = 5). To obtain knowledge regarding the mechanism of these phenomena, we investigated the composition of spheroids at different conditions by hematoxylin and eosin staining (Figure 7D). Spheroids treated with 20 μM Yoda1 at pH_e6.6 are largely fragmented (19.3 ± 1.5 % spheroid cross sectional area, n = 52 sections, N = 3 spheroids) compared to pH_e6.6 alone (6.1 ± 0.5 % area, n = 35, N = 3, p < 0.0001) (Figure 7C). In contrast, at pH_e7.4 there is no difference between vehicle (5.1 ± 0.4 % area, n = 58, N = 3) and 20 μM Yoda1 treatment (4.5 ± 0.4% area, n = 42, N = 3, p = 0.99). Lastly, we investigated the cellular viability in order to find out whether spheroid fragmentation is due to impaired cell metabolism and viability. MTT reduction normalized to control spheroids cultured at pH_e7.4 (1.00 ± 0.07, n = n = 6, N = 3) is almost halved at pH_e6.6 in the presence of 20 μM Yoda1 (0.52 ± 0.05, n = 6, N = 3, p = 0.03) (Figure 7E). For comparison, MTT reduction in the presence of 20 μM Yoda1 at pH_e7.4 (1.19 ± 0.07, n = 6, N = 3, p = 0.56) and pH_e6.6 cultured spheroids with vehicle treatment (0.82 ± 0.12, n = 6, N = 3, p = 0.6) is not different. However, loss of MTT reduction can also be attributed to decreased proliferation and decreased cell metabolism and is not a specific readout for cell viability in general. Therefore, we also performed Annexin V staining to detect apoptosis (Figure 7F). Here, cells derived from spheroids that were treated with Yoda1 and pH 6.6 have approximately twice as many Annexin V positive cells (31.0% ± 4.4, n = 6, N = 3) than under control conditions (16.3% ± 2.3, n = 6, N = 3, p = 0.0046).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.