MC3T3-E1 cells (sub-clone 14) were maintained in α-MEM media (Sigma Aldrich) supplemented with 10% Fetal Bovine Serum (FBS, HiMedia) and 100 units/mL Penicillin + 100 μg/mL Streptomycin solution (HiMedia). For preparing differentiation media, 10 mM β-glycerophosphate, 10 nM Dexamethasone and 100 μg/ml L-ascorbic acid (purchased from Sigma Aldrich) were added to the above-mentioned media.

Sprague-Dawley (SD) rat pups (0–2 days old) were procured from NISER animal house facility and sacrificed by cervical dislocation. Calvaria was carefully taken out, washed with PBS, muscles and blood were removed. The calvarial fragments were then subjected to enzymatic digestion by incubating in 10 mg/mL collagenase + dispase solution (purchased from Sigma) in 37°C water bath for 1.5 h with gentle shaking. Cells were collected after passing the enzyme digested tissue suspension through a 70 μm cell strainer. Cells were cultured in α-MEM supplemented with 10% FBS. For the isolation of mouse bone marrow-derived mesenchymal stem cells (MSC), we have followed the protocol as described before (Acharya et al., 2021).

Transient transfection was performed by Lipofectamine 3,000 Plus reagent (Invitrogen) according to the manufacturer’s protocol. Typically, 24–36 h after transfection, the cells were used for live-cell imaging or immunocytochemistry.

Alkaline Phosphatase Assay (ALP assay) media was prepared by adding 100 μg/ml L-ascorbic acid and 10 mM β-glycerophosphate to normal culture media. The ALP assay was performed by seeding equal number of cells (RCOs and MC3T3-E1) in 96-well plates (Corning) and culturing in ALP-media for 2 days in presence or absence of drugs. The RCOs were treated with different concentrations of 4αPDD (10 pM, 100 pM, 1 nM, 10 nM, 100 nM, 1 µM) and RN1734 (10 pM, 100 pM, 1 nM, 10 nM, 100 nM, 1 µM) along with ALP media. Similarly, the MC3T3-E1 cells were treated with (0.1 nM, 1 nM, 10 nM, 100 nM, 1 µM) and RN1734 (0.1 nM, 1 nM, 10 nM, 100 nM, 1 µM) with ALP media. After 2 days, media was aspirated, and cells were washed with PBS. Cells were lysed by subjecting the plate to repeated freeze-thaw cycle and ALP substrate (purchased from Sigma Aldrich, cat#N2770) was put in equal quantity in each plate. After 2–3 h of incubation, dark color develops. Subsequently the spectrophotometric reading was taken in 405 nm in an ELISA plate reader (VarioSkan, Thermo scientific).

Mineralization media was prepared by adding 10 mM β-glycerophosphate, 100 μg/ml L-ascorbic acid and 10 nM Dexamethasone (Sigma) to normal culture media (Siddiqui et al., 2010; Siddiqui et al., 2011). Mineralization assay was performed by seeding 1 × 10⁵ number of cells (RCOs) in 12-well culture plates and cultured in mineralization media for 21 days in the presence of TRPV4 channel agonist 4αPDD (10 nM). After 21 days, media was aspirated, cells were washed with PBS and an equal amount of Alizarin Red-S (ARS) dye (Sigma Aldrich, cat#A5533) was added to each well. After 30 min of incubation with ARS, the dye was washed away with water and micrographs were taken to visualise the extent of mineralization in different conditions as required in individual experiments. For quantifying the extent of mineralization, Cetyl Peridynium Chloride (CPC, purchased from MP Biomedicals) were added in each well. The dye was extracted in CPC and spectrophotometric reading was taken at 550 nm. In another condition mouse bone marrow derived MSCs were treated with 4αPDD (1 µM) only and RN1734 (1 µM) + 4αPDD (1 µM) for 15 days along with osteogenic differentiation media. In a similar manner, for mineralization of MSCs, we used GSK1016790A (10 nM, 100 nM, 1 µM) as another specific agonist of TRPV4. ARS staining and CPC extraction and quantification was performed as mentioned above.

Cells were cultured on 12 mm cover slips in 24 well plates. After differentiation, cells were fixed with 4% Paraformaldehyde (PFA, purchased from Sigma) for 15 min in room temperature (∼25°C). PFA was then aspirated from the wells and cells were washed three times with PBS. Cells were then permeabilized with 0.1% Triton X-100 in PBS (1X) for 4 min and blocked with 5% Bovine Serum Albumin (BSA) for 30 min. The coverslips were then washed thrice with 200 μL 0.1% PBS-T for 5 min each. After washing with PBS-T, cells were probed with the anti-TRPV4 antibodies (Alomone Labs, Israel, Cat#ACC-034) at a dilution of 1:250 in different experimental settings. Alexa Fluor-488 conjugated secondary antibody (Invitrogen) with 1:500 dilution was used to visualize TRPV4 channel and DAPI (5 μM) was used as a counterstain to visualize the nucleus. For confirming the specificity of the antibodies for the TRPV4 channel, epitope-specific peptide procured from Alomone Labs (Cat#BLP-CC034) were used. Firstly, the antibodies were incubated with respective antigenic peptides for 30 min and then cells were probed with the antigen-antibody complex formation.

For live cell imaging of basal Ca²⁺-level, rat calvarial osteoblasts (RCOs) were plated on 25 mm coverslips in 35 mm tissue culture dishes (Corning) and cultured in differentiation media for 3 days. Cells were treated with pharmacological activator of TRPV4 (4αPDD, 10 nM) for 2 days. Subsequently, cells were treated with 0.02% Pluronic^® F-127 (Thermo Fisher Scientific) and 5 μM Fluo-4 AM (Molecular Probe) for 45 min and live-cell imaging was then performed in a Zeiss LSM 780 confocal microscope.

For the measurement of cytosolic Ca²⁺ level the MC3T3-E1 cells were transfected with genetically encoded Ca²⁺-sensor GCaMP. Post 24 h. of transfection, the Ca²⁺-imaging was performed by using 4αPDD (5 µM) and RN1734 (10 µM). The drugs were added on the 20th frame of the time series and the whole live cell imaging was recorded up to 200th frame (∼3.5 min). We further treated the cells with RN1734 (10 µM) for 1 h and subsequently the cells were activated by 4αPDD (5 µM) (without washing RN1734) and the Ca²⁺-influx was recorded up to 200th frame (equivalent to ∼3.5 min). Higher concentration of drugs was used to check the instant activation of TRPV4 channel. The change in calcium intensity was quantified by ImageJ.

For measurement of cytosolic Ca²⁺ in MSCs, the cells were labelled with Fluo-4-AM dye (5 μM) and live cell Ca²⁺ imaging was performed by adding GSK1016790A (1 µM) at the 20th frame. In corresponding control experiments, the cells were pre-incubated with RN1734 (10 μM for 1 h) and instant Ca²⁺ influx was measured using GSK1016790A (without washing RN1734) as the stimulus.

Quantitative real-time polymerase chain reaction (qPCR) was performed to determine the relative expressions of osteoblast specific genes in drug-treated RCOs. RCOs were treated with 4αPDD (10 nM) for 48 h along with α-MEM media (Sigma Aldrich) supplemented with 10% FBS and 100 units/mL Penicillin + 100 μg/mL Streptomycin solution. GAPDH was used as the internal control. Primers were designed by the Universal Probe Library (Roche Applied Science) for the following genes: RunX2, Bmp2, and Col1. The primer list is described previously (Acharya et al., 2021). All cDNAs were synthesized with Revert Aid cDNA synthesis kit (Fermentas, Austin, United States) using 2 µg total RNA. SYBR green chemistry was employed to perform quantitative detection of relative expression of mRNA levels of these genes using a Light Cycler 480 (Roche Molecular Biochemicals, Indianapolis, United States) as described before (Siddiqui et al., 2010).

For the preparation of gel sample, the RCOs grown in differentiation media for 7 days were harvested. The gel sample is prepared by adding Laemmli buffer supplemented with protease inhibitor cocktail (Sigma) and subsequently samples were boiled for 5 min at 95°C. The samples were analyzed by 10% SDS-PAGE and followed by Western blot analysis as described before (Kumar et al., 2016; Acharya et al., 2021).

Glycolysis and mitochondrial respiration were quantified using a previously described procedure (Porwal et al., 2017). Briefly, rat calvarial osteoblast were cultured in α-MEM with 10% FBS. Glycolysis and mitochondrial respiration experiment was carried out without the presence of mineralization media. 48 h prior to assay 40,000 cells/well were plated in a 24-well polystyrene Seahorse V7-PS Flux plate with no additional coating and treated with 4αPDD (10 nM). Before starting the assay, cells were washed twice with assay media (XF Base minimal DMEM, catalog number 98546008) and kept at 37°C for 1 h. Glycolysis was analyzed by stepwise addition of A: glucose (10 mM), B: oligomycin (1 μM) and C: 2-deoxy-D-glucose (50 mM). Mitochondrial respiration was assayed by stepwise addition of A: oligomycin (1 μM), B: carbonyl cyanide-4-(trifluoromethoxy) phenylhydrazone (FCCP) (2 μM) and C: rotenone (0.5 μM). Three measurement cycles of 2 min mix, 1 min wait, and 5 min measure were carried out after each addition.

Mitochondrial Ca²⁺-imaging was performed with ratiometric Mito-Pericam constructs (A gift from Dr. Atsushi Miyawaki, Saitama, Japan). Ratiometric Mito-Pericam is a Ca²⁺-indicator made by fusion of the yellow fluorescent protein (YFP) and Calmodulin which is able to translocate within mitochondria (Nagai et al., 2001). The binding of Ca²⁺ to Mito-Pericam changes its excitation wavelengths from 415 nm to 494 nm, while its emission spectrum is maintained at 515 nm. Mito-Pericam was expressed in MC3T3-E1 cells by transient transfection and 24 h after transfection. Cells were imaged with a confocal microscope. Ca²⁺-imaging was performed by adding 4αPDD (5 µM), RN1734 (10 µM) at the 20th frame during the imaging. In some experiments, cells were pre-incubated with RN1734 (10 µM) for 1 h and subsequently activated by 4αPDD (5 µM). TRPV4 activator (4αPDD) was used at 5 μM (slightly higher than the optimum concentration) to assess the instantaneous influx of Ca²⁺ inside the mitochondria. Using particle-based quantification by ImageJ, we have measured the mitochondrial Ca²⁺- level. For that purpose, time-series imaging was performed (total of 200 frames (∼7.5 min) and 4αPDD was added on the 20th frame in each condition). Images at different time points (Frame-1, 20, 30, 50, 100, 150, and 200) were used to analyse the Ca²⁺-level. More than 18,000 mitochondrial particles from multiple cells were quantified in each frame.

For the quantification of mitochondrial morphological parameters, we used mitochondria morphology plugin (https://imagejdocu.tudor.lu/plugin/morphology/mitochondrial_morphology_macro_plug-in/start) as described before (Dagda et al., 2009). The quantification of ratio of mitoDsRed intensity with GFP intensity (of TRPV4-WT and TRPV4 mutants), fluorescence intensity for GFP as well as DsRed from individual mitochondria were analysed by ImageJ.

Statistical analysis were performed by using GraphPad Prism 7 (Version 7.00). Data represented in the graphs as mean ± SEM. One-way ANOVA test with Brown-Forsythe test, Bartlett’s test and Sidak’s multiple comparisons test, and Student’s t-test were done as and where required.

All the animal experiments were performed at the animal house facility of NISER and CDRI as per the guidelines of IAEC (NISER/SBS/AH/IAEC-56 & NISER/SBS/AH/IAEC-55, CDRI/IAEC/2013/34). All the experimental methods were carried out by following relevant guidelines and regulation of CPCSEA.

We studied the endogenous expression of TRPV4 in rat calvarial osteoblasts (RCOs) by immunofluorescence and western blotting for which the specificity of primary antibodies has been confirmed by respective peptide block (Figures 1A, B). Similar immunofluorescence approach was applied for studying TRPV4 expression in murine pre-osteoblast cell line MC3T3-E1 (Figure 1C). Next, we studied the functionality of TRPV4 and thus performed Ca²⁺-imaging in MC3T3-E1 osteoblasts using a TRPV4 activator, 4αPDD. MC3T3-E1 cells were transiently transfected with GCaMP. 24 h after transfection, cells were treated with 4αPDD and we observed Ca²⁺-rise that sustained in a manner (Figure 1E). In a similar experiment, when cells were treated with a TRPV4 inhibitor, RN1734 (10 µM) the intracellular Ca²⁺-level was reduced (Figure 1F). However, when cells were pre-treated with RN1734 (10 µM) for 1 h and subsequently treated with 4αPDD, the intracellular Ca²⁺-level was further elevated (Figure 1G). In differentiating RCOs, the basal cytosolic Ca²⁺-levels increased in the 4αPDD (10 nM) treated cells (Figure 1D). Instantaneous Ca²⁺-influx in response to GSK1016790A (1 µM, as an another agonist for TRPV4) in MSC was also observed (Supplementary Figure S1). When cells were pre-treated with RN1734 (10 µM, for 1 h), Ca²⁺ influx in response to GSK1016790A was also observed, mainly due to lack of Ca²⁺-mediated desensitization (discussed later). We observed elevated Ca²⁺ level in both the experimental conditions, but the presence of RN1734 dampened the Ca²⁺-influx faster as compared to only GSK1016790A (Supplementary Figure S1).

We investigated the role of TRPV4 in the differentiation in MC3T3-E1 cells. To this aim, we performed an alkaline phosphatase (ALP, 7 days) assay, a marker of osteoblast differentiation. MC3T3-E1 cells were cultured with or without 4αPDD or RN1734 in increasing concentrations (Figures 2A, B). However, the increment of ALP intensity remains unpredictable, especially in certain doses of RN1734 conditions. This might be due to complex biological variation and/or some unexplained reasons. We observed that 4αPDD increased the differentiation of MC3T3-E1 cells by ∼1.7 folds compared with DMSO control (Figure 2A). We next studied if TRPV4 modulated the differentiation of RCOs. To this aim, RCOs were cultured in the presence of increasing concentration of 4αPDD or RN1734 (10 pM to 1 μM) for 2 days and then cells were fixed in 4% PFA (Figures 2C, D). TRPV4 activation significantly promoted differentiation of RCOs (at 1 nM, 4αPDD increased differentiation by ∼1.4 folds) compared with DMSO control (Figure 2C). Moreover, 4αPDD (10 nM) increased the differentiation of RCOs at day 7 by ∼2.7 folds compared with DMSO control (Figure 2E).

We next studied the ability of 4αPDD (10 nM) in stimulating the formation of ex vivo mineralized nodule formation by bone marrow stromal cells obtained from rats. As shown in (Figure 2F), 4αPDD increased mineralized nodule formation by ∼2.5-fold compared with control cells. In the same condition, cells were activated with 4αPDD alone (at 1 μM) or in presence of RN1734 (1 μM). We noted that 4αPDD alone causes more mineralization as compared to the control. However, presence of RN1734 reduces the efficacy of 4αPDD-mediated mineralization significantly (∼42%). Therefore, RN1734 can reverse the activity of 4αPDD (Supplementary Figure S2). Using a similar protocol, we have performed induction of in vitro mineralization in MSC using another TRPV4 agonist GSK1016790A (10 nM, 100 nM, 1 µM). GSK1016790A treatment increased mineralized nodules as compared to control condition (Supplementary Figures S3A, B).

Moreover, in RCOs, 4αPDD (10 nM, 48 h) significantly increased the mRNA levels of osteogenic genes including runt-related transcription factor 2 (Runx2), bone Morphogenic protein-2 (Bmp2), and collagen type-1 (Col1) (Figures 2G–I). Taken together, data suggests that TRPV4 activation induces the expression of osteogenic genes required for osteogenic differentiation.

Osteoblast differentiation is a highly energy-intensive process and it uses both glycolysis and mitochondrial respiration to fuel the process (Guntur et al., 2014). Therefore, we studied the effect of TRPV4 activation on the regulation of energetics of RCO. Using extracellular flux analyzer, we observed that TRPV4 activation increased basal glycolysis (extracellular acidification), glycolytic reserve and capacity, under normal glucose concentration (1 mg/mL) (Figure 2J). Next, we studied the effects of TRPV4 activation in mitochondrial respiration. TRPV4 activation increased both basal and maximum mitochondrial respiration (oxygen consumption), spare respiratory capacity and ATP production in osteoblasts (Figure 2J).

Several point mutations in TRPV4 are known to induce musculoskeletal defects (Verma et al., 2010). In order to assess for the impact of TRPV4 mutations on the mitochondrial function, we co-expressed full-length wild-type (WT) or a series of TRPV4 point mutants (R616Q, F617L, L618P, V620I, L596P) along with a marker for mitochondria (i.e., mitoDsRed) in MC3T3-E1 cells by transient transfection (Figure 3A). In all these cases, colocalization of TRPV4 with mitoDsRed was observed albeit with variable extent. We then quantified the parameters including mitochondrial number, size, distribution, roundness and protein import that determine mitochondrial health. Mitochondrial number and distribution in osteoblasts did not alter with the altered TRPV4 function (Figure 3B). However, both area and perimeter of the mitochondria were reduced in the presence of most of the TRPV4 mutants (Figures 3D, F). The mito content and the overall area-perimeter ratio were also calculated (Figures 3G,H). Both solidity and circularity of mitochondria, indicative of their morphology, were affected in the presence of specific mutants. L596P, L618P, and V620I mutations increased the solidity of mitochondria (Figures 3C, E), whereas L596P and L618P mutations increased mitochondrial circularity (Figure 3C). To understand the mitochondrial translocation/import, we measured the intensity of TRPV4 as well as mitoDsRed for cells expressing TRPV4-WT or different mutants. Both mitoDsRed and TRPV4 fluorescence intensities were increased in the presence of mutations (Figures 3I–K). The ratio of TRPV4-GFP intensity with mitoDsRed intensity, however, was increased only in the presence of F617L, while it was decreased in L596P and V620I mutations, suggesting that different mutants affect the mitochondrial import in different extent (Figures 3I–K).

Mitochondrial Ca²⁺-level is a critical parameter for the function of osteoblasts. Thus, we characterized the level of mitochondrial Ca²⁺ upon pharmacological activation of TRPV4. Our data show that mitochondrial Ca²⁺ was significantly increased upon the activation of TRPV4 by 4αPDD (5 µM) in MC3T3-E1 cells (Figures 4A, D). By contrast, when cells were treated with RN1734 (10 µM), marginal Ca²⁺-influx was observed (Figures 4B, D). Further, we treated the cells with RN1734 (10 µM) for 1 h and subsequently treated with 4αPDD (5 µM). In this condition also, increase in the mitochondrial Ca²⁺ was observed suggesting that TRPV4 as a regulator of mitochondrial Ca⁺²-levels in osteoblasts (Figures 4C, D).