Porcine aortic VICs (paVICs) were isolated from pig hearts commercially acquired via a local abattoir (Animal Technologies, Tyler, TX) as previously described (9). Valves from 6 hearts were used per isolation. Briefly, aortic valves from fresh porcine hearts on ice were dissected and washed in cold phosphate buffered saline (PBS) containing 5% antibiotic-antimycotic (ABAM; Thermo Fisher Scientific 15240062) to remove blood and debris. After dissection, the valves were incubated in a solution of 500 U/ml collagenase type II (Worthington Biochemical LS004177) in DMEM (Corning 10-014-CV) with 1% ABAM for 30 min and wiped with a sterile swab to remove the endothelial cell layer. The denuded valves were minced and incubated in a mixture of 300 U/ml collagenase type III (Worthington Biochemical LS004182), 50 U/ml hyaluronidase, and 0.5 U/ml neutral protease (Stemcell Technologies 07913) in DMEM with ABAM and 25 mM HEPES (Millipore Sigma H4034) for 4 h at 37°C in an orbital shaker. The digested suspension was filtered through a 40 μm cell strainer to remove large debris, and the filtrate was centrifuged at 1500 g for 5 min to pellet the cells. The pellet was resuspended in growth medium consisting of 1:1 DMEM:F12 (Cytiva Life Sciences SH30026) with 10 mM HEPES, 1% ABAM, and 10% bovine growth serum (BGS) and plated on a growth flask. Cultures were closely monitored for contamination for the first few days after isolation. Cells were allowed up to 7 days to attach. When cells began to grow, they were passaged once and then frozen in medium consisting of 90% BGS and 10% hybridoma-grade DMSO (Sigma Aldrich D2650) until experimentation.

Reagent preparation was done according to manufacturer directions, and culture conditions were modified from previous studies (9). Dantrolene (EMD Millipore 251680) was dissolved in hybridoma-grade DMSO in stock aliquots of 10 mM and stored at −80°C. Working aliquots were made from thawed vials and stored at −20°C to reduce freeze-thaw. LPC (Sigma-Aldrich L4129) was stored in desiccating conditions at −20°C and dissolved in absolute ethanol (EtOH) at 10 mM just prior to use. For some experiments, osteogenic medium (OM) consisting of 10 nM dexamethasone (Sigma-Aldrich D4902) and 10 mM β-glycerophosphate (Sigma-Aldrich G9422) was used as a comparison to LPC. Reagents were diluted to final concentrations in low-serum medium (1:1 DMEM:F12 with 10 mM HEPES, 1% ABAM, and 1% BGS) unless otherwise specified. During experimentation, cells were exposed to one of 4 conditions, unless otherwise specified: vehicle-only (containing a volume of DMSO and ethanol each equal to the highest volumes of dantrolene and LPC, respectively, used in that experiment), LPC (with DMSO vehicle), dantrolene (with ethanol vehicle), or LPC and dantrolene.

When ready for experimentation, paVICs were thawed and passaged once using growth medium. Once confluent, the paVICs were passaged and seeded onto the appropriate culture plates at the density desired for the experiment, described below. Cells were then exposed to low-serum medium for 1–2 days before experimentation. Experiments were broadly divided into short- and long-term categories, as illustrated in Figure 1. Short-term experiments examined rapid events due to LPC occurring less than 24 h after exposure. For short-term experiments, paVICs were pretreated with 60 μM dantrolene or DMSO vehicle overnight prior to LPC or EtOH vehicle exposure. For long-term experiments, examining events greater than 24 h after initial exposure to LPC, paVICs were simultaneously exposed to dantrolene or DMSO vehicle and LPC or EtOH vehicle, and conditions were maintained throughout the course of experimentation, changing medium every 3 days, as needed.

To determine the appropriate treatment concentration of dantrolene, we first used an alamarBlue (Bio-Rad BUF012) cell viability assay (19). Cells were treated with serial dilutions of dantrolene overnight in an incubator. A live control was treated with only low-serum medium, and a dead control was treated with 10% ethanol. The next day, the culture medium was removed, and the cells were incubated with a 10% alamarBlue solution in HEPES-buffered Hank's Balanced Salt Solution (HHBSS) in an incubator. After one hour, fluorescence was read at an excitation and emission of 530 and 590 nm, respectively, using a SpectraMax M2 plate reader (Molecular Devices). Signal was normalized to that of the live control. Cell viability curves were fitted to a four-parameter logistic curve, and the interpolated concentration that corresponded to 50% signal of the live control was used to calculate the cellular median lethal dose (LD50). The average LD50 of two independent experiments is reported.

Other cells, treated with a similar amount of dantrolene or DMSO, were treated with a Live/Dead staining kit (Invitrogen L3224) consisting of calcein-AM and ethidium homodimer. Cells were treated with a 1:1,000 dilution of calcein-AM and ethidium homodimer in HHBSS for 30 min in an incubator. After washing, the cells were imaged on an ECLIPSE Ti2E microscope (Nikon Instruments) using FITC for detecting living cells and Texas Red for detecting dead or compromised cells. Acquired images were analyzed with a custom CellProfiler (Broad Institute) pipeline to count the number of objects in each channel and determine percent viability as described elsewhere (20). Six replicates were used per group, and data from two independent experiments are displayed.

To quantify changes in calcium flux, we used a Fluo-8 calcium assay (Abcam ab112128) as previously described. Briefly, cells were confluently seeded onto a clear-bottomed, black-walled 96-well plate. The next day, they were exposed to either 60 μM dantrolene in DMSO or an equivalent amount of DMSO (0.6% v/v) only overnight. The following day, the medium was removed and replaced with assay buffer consisting of 1:1,000 Fluo-8 dye from stock in 1× Pluronic F127 Plus in HHBSS, provided by the kit. The cells were incubated at 37° for 30 min then 30 more minutes at room temperature. After dye loading, calcium flux was immediately measured in a TECAN Spark multimode plate reader at an excitation/emission of 490/525 nm. The appropriate signal gain was calibrated using carbachol (Abcam 141354), a calcium agonist. The plate reader's automated pipettes were then used to deliver LPC at a final concentration of 10 μM in HHBSS or an equivalent volume of 0.1% EtOH in HHBSS. Fluorescence was read every second for 5 s before adding reagents and for 25 s after adding reagents. Data are reported as the maximum fluorescence achieved after the addition of reagents minus the average of the signal before the addition of reagents, which was considered background. Six replicates were used per control group with 18 replicates per dantrolene-treated group.

The formation of paVIC calcific nodules was assayed as previously described (9). Briefly, paVICs were seeded onto a 48-well plate in low-serum medium at 50,000 cells/cm² to yield a confluent monolayer. After two days, the medium was changed to conditioned media, consisting of DMSO only, 10 μM LPC with DMSO vehicle, or a 10 μM, 30 μM, or 60 μM dose of dantrolene with 10 μM LPC. Separate experiments were conducted using OM as a reference. Calcific nodules were allowed to form for 6 days, and calcific media were changed on day 3. On day six, after media removal, the cells were washed with PBS, fixed with 4% paraformaldehyde for 30 min, washed twice with deionized water, and left to dry overnight. The next day, samples were stained for 30 min with 40 mM alizarin red-S (Sigma-Aldrich A5533) in deionized water adjusted to a pH of 4.2–4.4. After staining, the samples were washed with deionized water 3 times until the wash was clear. Samples were imaged on the ECLIPSE Ti2E, and nodule size and number were quantified using ImageJ. Four replicates were included for each experimental group, and data from one representative experiment are reported.

The activity of caspases 3 and 7 was measured as an analog for apoptosis using the Caspase-Glo 3/7 Assay Kit (Promega G8091). Cells were seeded confluently onto a 384-well plate and allowed to attach overnight. The next day, cells were treated with 10, 20, 40, or 80 μM LPC or ethanol control and either 60 μM dantrolene or DMSO vehicle overnight. The following day, the reagents were applied per the manufacturer's directions, and then the luminosity was detected on the SpectraMax M2 plate reader. Four replicates were included per control group, and 6 replicates were included per test group. Data from two independent experiments are reported.

To quantify the production of ROS due to LPC, we used a CM-H₂DCFDA general oxidative stress indicator (Thermo Fisher C6827). To measure ROS, paVICs were plated on 6-well plates and allowed to grow to 90% confluency in growth medium. When ready, the cells were treated with 60 μM dantrolene or DMSO vehicle overnight in low-serum medium. The next day treated media were removed, the cells washed with PBS, and then exposed to 10 μM LPC for 10–60 min in HHBSS or ethanol vehicle for 60 min in an incubator. Other dantrolene treated and untreated control groups were exposed to only HHBSS or 200 μM tert-butyl hydroperoxide (Thermo Fisher 180340050), a potent inducer of ROS, for 30 min. After exposure, cells were washed with PBS and loaded with 10 μM CM-H₂DCFDA in HHBSS for 30 min in an incubator. After dye loading, the cells were lifted from their plates using 0.25% trypsin, collected in HHBSS without calcium or magnesium, and fluorescent intensity was examined using flow cytometry in an MA900 Cell Sorter (Sony). Median fluorescence intensity (MFI) was used to determine changes in ROS production. Flow data were analyzed as MFI using Flowing Software (version 2.5.1). Three replicates were included per experimental group, and data from two independent experiments are reported.

Quantitative real-time polymerase chain reaction (qRT-PCR) was carried out as previously described (9). Briefly, paVIC cultures were exposed to LPC, dantrolene, or their respective vehicles for 1 or 3 days. After experimentation, RNA was extracted using TRIzol reagent (Invitrogen 15596018) and purified using a Direct-zol RNA Microprep spin kit (Zymo Research R2062) per the manufacturer's directions, with a DNAse treatment. RNA was eluted into RNAse-free water. RNA quantity and quality were measured using a Nanodrop 2000 (Thermo Fisher). A value greater than 1.8 was considered acceptable for 260/280 and 260/230 ratios. Reverse transcription was conducted using a High-Capacity cDNA Kit (Applied Biosystems 4368813). qRT-PCR was performed on a CFX96 Real-Time PCR System (Bio-Rad) using probes listed in Supplementary Table S1. Three to 6 replicates were used per experiment.

Total RNA was extracted from paVICs using the same methods as for qRT-PCR. Three experiments, each using cells from separate isolations, were conducted on different days, and each experiment had one replicate per group. Purified RNA samples were sent to Novogene Corporation United States (Sacramento, CA), which performed quality control, library construction, sequencing, and data cleaning. RNA sequencing was conducted using Illumina platforms. After the data were received, annotation was carried out for Sus scrofa using STAR (v2.7.9), and batch effect adjustment was carried out using the ComBat function of the SVA package in R (21, 22). Differential gene expression was determined using DESeq2 (v1.32.0) in R (23). Data visualization was performed in R using the following: heatmaps were generated using ComplexHeatmap (v2.8.0), volcano plots were generated using EnhancedVolcano (v1.11.5), and finally GO, WikiPathways and KEGG pathway analysis was carried out with enrichplot (v1.12.3) (24, 25). The data discussed in this publication have been deposited in NCBI's Gene Expression Omnibus (26) and are accessible through GEO Series accession number GSE227229.

Statistical analysis was performed using R version 4.1 (27). Quantitative data are expressed in the text as mean ± standard deviation and graphically as box-and-whiskers indicating median (crossbar), interquartile range (hinges), and 1.5 times interquartile range (whiskers). Experiments were repeated at least twice with the number of replicates specified unless otherwise noted. Statistical analysis was performed using n-way ANOVA with post hoc Tukey's test or Dunnett's test as appropriate. Significance was called at P = 0.05.

The paVICs demonstrated an LD50 of 297.8 ± 1.8 μM to dantrolene using an alamarBlue assay (Figure 2A). The cells demonstrated good viability until about 150 μM dantrolene and showed much lower viability due to high toxicity at greater doses. A dose of 600 μM regularly killed all cells in the sample. Since alamarBlue detects mitochondrial activity, treatments that stress the cells can result in a slightly elevated signal from a live (non-treated) control. Therefore, we confirmed our results using a Live/Dead stain with microscopic examination (Figures 2C–E). While there was very low baseline cell toxicity at low doses (Figure 2C), increased toxicity could be observed beginning at 75 μM (Figure 2D). At 300 μM, nearly all the cells in the sample were dead or had a compromised membrane (Figure 2E). Based on these results, we chose to use a dose of 60 μM dantrolene or less in all further experiments, which allowed for good viability with minimal interference with other assays.

We further examined the effects of our reagent combinations on paVICs using an alamarBlue assay (Figure 2B). Dantrolene (60 μM) had no effect on viability compared to vehicle-treated cells (95.9 ± 19.9% vs. 100.0 ± 13.6%, P = 0.992). Meanwhile, treatment with 10 μM LPC showed an apparent increase in viability compared to all other groups (135.5 ± 17.0%, P < 0.001, all comparisons). This apparent increase in viability could be due to increased cell proliferation or mitochondrial stress, either of which would indicate that LPC negatively affects paVIC quiescence. Treatment with dantrolene and LPC reversed this increase in alamarBlue signal (78.0 ± 19.3%, P < 0.001 vs. LPC). Treatment with both dantrolene and LPC decreased the viability of paVICs compared to vehicle-only treatment (P = 0.042) but not dantrolene only (P = 0.151), indicating that there may be some slight toxicity in the combination treatment.

We investigated the effects of 60 μM dantrolene on LPC-induced calcium flux using Fluo-8, a calcium-sensitive dye. We found that overnight pretreatment with dantrolene inhibited LPC-induced calcium flux (Figures 2F–G). Treatment with LPC caused a significant increase in calcium flux compared to vehicle-treated cells (476.4 ± 551.2 vs. −103.3 ± 321.2, P = 0.028). Pretreatment with 60 μM dantrolene successfully blocked this increase in calcium flux due to LPC (95.0 ± 301.6, P = 0.011). Representative curves for each group can be seen in Figure 2G.

Dantrolene inhibited calcific nodule formation of paVICs due to 10 μM LPC (Figures 3A–H). Treatment with dantrolene at doses of 60, 30, or 10 μM decreased total calcified area (Figure 3A), nodule number (Figure 3B), and nodule size (Figure 3C) compared to LPC in vehicle-treated cells. To confirm dantrolene's calcific nodule inhibiting ability, we repeated these experiments with osteogenic medium (OM) (Figures 3A–D,I–L). Comparable results were found with OM, but a 10 μM dose of dantrolene was ineffective at preventing OM-induced nodules (Figure 3L).

LPC induces apoptosis in various cell types, and apoptotic cells may contribute to calcific nodule formation. We measured apoptosis in paVICs via caspase 3/7 activity (Figure 4A). Since 10 μM of LPC did not induce measurable levels of apoptosis in paVICs, we performed serial doublings and determined that 80 μM of LPC was sufficient to induce apoptosis without signal interference from the vehicle. At 80 μM, LPC significantly induced caspase 3/7 activity compared to the vehicle-only control as measured by a luminescent assay (942.8 ± 94.4 vs. 172.3 ± 31.6 RLU, P < 0.001). In cells pretreated with dantrolene, significantly less caspase activity was measured (577.4 ± 273.0 RLU, P = 0.017), but cells treated with 80 μM LPC + 60 μM dantrolene treatment still had higher caspase activity than the vehicle-only control (P = 0.018). These results indicate that dantrolene partially inhibits the apoptotic effects of LPC on paVICs.

LPC induces ROS in a number of cell types, and dantrolene has been shown to inhibit ROS formation. Therefore, we hypothesized that dantrolene would inhibit LPC-mediated ROS. We measured ROS formation via flow cytometry using a ROS-sensitive fluorescent dye (Figure 4B). Compared to an untreated reference, LPC-treated paVICs displayed a significant increase in ROS at 50 min of exposure (3.3 ± 1.6-fold change from paVICs treated with medium-only, P = 0.014); compared to dantrolene + LPC-treated cells, which showed significantly increased ROS at 60 min (6.0 ± 2.0-fold change, P < 0.001). No difference was found between LPC- or dantrolene + LPC-treated groups.

Because of this unexpected result, we next confirmed the antioxidant properties of dantrolene using another ROS-inducing agent. paVICs were treated with 60 μM dantrolene or DMSO vehicle and 300 μM tert-butyl hydroperoxide (TBP), a potent inducer of ROS (Figure 4C). TBP induced a strong ROS response compared to the vehicle-only control (9.46 ± 3.8-fold change vs. 1.0 ± 0.3, P = 0.003). Treatment with dantrolene inhibited this ROS production (3.6 ± 0.9-fold change, P = 0.028). ROS production from treatment with dantrolene was not different from the vehicle-only group (0.45 ± 0.05, P = 0.99). Representative histograms for the ROS flow cytometry can be seen in Figure 4D. These results indicate that dantrolene does not inhibit LPC-mediated ROS but does inhibit ROS induced by other sources.

Activation of paVICs as measured via αSMA (gene symbol: ACTA2) has long been considered a disease hallmark and a pro-CAVD phenotype. Tissue culture polystyrene artifactually induces this expression of αSMA, and factors that reduce its expression are believed to be beneficial in CAVD. We measured ACTA2 expression at 1 and 3 days of exposure to LPC ± dantrolene via qRT-PCR. At 1 day, co-treatment with LPC and dantrolene significantly reduced ACTA2 expression relative to both 10 μM LPC alone (0.8 ± 0.1 vs. 0.6 ± 0.1 fold change, P = 0.006) and vehicle control (1.0 ± 0.1 fold change, P < 0.001) (Figure 4E). Even in the absence of LPC, treatment with 60 μM dantrolene significantly reduced ACTA2 expression compared to vehicle control (0.5 ± 0.1-fold change, P < 0.001).

By 3 days, ACTA2 expression was consistent between the vehicle-only and 60 μM dantrolene groups (1.0 ± 0.0 vs. 1.02 ± 0.2 fold change, P > 0.99). The 10 μM LPC group had significantly higher ACTA2 expression (1.35 ± 0.1 fold change, P = 0.031) than the vehicle control, the 60 μM dantrolene group (P = 0.032), and the dantrolene + LPC group (1.03 ± 0.1 fold change, P = 0.048) (Figure 4H). These results demonstrate that 60 μM dantrolene effectively reduces ACTA2 expression in paVICs.

Osteoblastic differentiation of paVICs drives valve calcification, so we examined the expression of the key regulatory factors Runt-Related Transcription Factor 2 (gene symbol: RUNX2) and SRY-Box Transcription Factor 9 (SOX9) using qRT-PCR at 1 day (Figures 4F–G) and 3 days (Figures 4I–J). We found that both RUNX2 and SOX9 are significantly upregulated at both time points due to 60 μM dantrolene.

RUNX2 expression was significantly upregulated with 60 μM dantrolene treatment compared to vehicle control at 1 day (1.5 ± 0.2 vs. 1.0 ± 0.1-fold change, P < 0.001) and 3 days (1.74 ± 0.1 vs. 1.0 ± 0.1, P < 0.001). Treatment with 10 μM LPC did not significantly affect RUNX2 expression at 1 day (0.9 ± 0.1, P = 0.702) but resulted in decreased RUNX2 expression by 3 days (0.73 ± 0.0, P = 0.047). Co-treatment with dantrolene and LPC significantly increased RUNX2 expression over LPC treatment alone at 1 day (1.4 ± 0.1, P < 0.001) and 3 days (1.6 ± 0.2, P < 0.001).

Regarding SOX9, dantrolene similarly increased its expression compared to vehicle control at both 1 day (2.5 ± 0.5, P < 0.001) and 3 days (2.7 ± 0.7, P = 0.006). LPC did not affect SOX9 expression compared to vehicle control at 1 or 3 days. Co-treatment with dantrolene + LPC increased SOX9 expression compared to LPC alone at 1 day (1.4 ± 0.1 vs. 0.85 ± 0.1-fold change, P < 0.001), but this effect was not significant at 3 days. Overall, these data show that dantrolene increases the expression of the gene regulation factors RUNX2 and SOX9.

Given the broad signaling and regulatory implications of calcium blockade and upregulation of RUNX2 and SOX9, we used RNAseq to survey the global regulatory changes caused by treatment with LPC and dantrolene. Principal component analysis of the ComBat-adjusted treatment groups (Figure 5A) showed close clustering of vehicle-only treated samples, separate from those treated with 10 μM LPC, which also clustered together. Meanwhile, those treated with dantrolene or LPC + dantrolene clustered together, suggesting that these samples shared more similar gene expression profiles to one another than to the LPC-treated group. The sample distance heat map (Figure 5B) affirmed this relationship between samples.

Venn diagrams of differentially expressed genes with P_adj < 0.05 demonstrated that 690–980 genes were upregulated, and 10–85 genes were downregulated in all treatment groups compared to vehicle controls. The majority (approximately 62%–87%) of the significantly upregulated genes were shared between the three treatment groups (Figure 5C), but each group also had uniquely upregulated genes. In contrast, a Venn diagram of downregulated genes (Figure 5D) showed that the LPC group had no commonly downregulated genes compared to the dantrolene group. Further, the LPC + dantrolene group had only one downregulated gene in common with the LPC group, suggesting that dantrolene-mediated downregulation of genes dominated gene downregulation. All significantly different genes can be found in Supplementary Data Sets A–F.

Volcano plots of genes with a P_adj < 0.05 (Figures 5E,F) showed that LPC strongly promoted gene upregulation, as did dantrolene, to a lesser extent. The volcano plot of differentially regulated genes between the LPC + dantrolene group and the LPC group (Figure 5G) revealed many interesting genes that may provide insight into the regulation of calcification in valve disease. Notably upregulated genes include CYP1B1, a cytochrome P450 family enzyme shown to be atheroprotective (28, 29) and associated with decreased oxidative stress and inflammation (30); BCL3, an NFκB inhibitor associated with atherosclerotic changes (31); IL6R, the interleukin-6 receptor, mutations of which have been implicated in increased vulnerability to CAVD (32, 33); and NQO1, an NAD(P)H oxidoreductase for which upregulation has been linked with decreased calcification (34). Notably downregulated genes of interest include a DNA histone deacetylase, HDAC9, which has been associated with vascular smooth muscle cell calcification (35, 36); the angiotensin II receptor, AGTR1, which in addition to its role in blood pressure and general cardiovascular disease has also been associated with aortic stenosis and calcification (37); an actin organization gene, SORBS2, which has been implicated in exosome regulation (38) and myxomatous mitral valve disease (39); and the collagen synthesis enzyme prolyl-4-hydroxylase, P4HA1, which is induced by hypoxia inducible factor 1 (HIF1) (40), a known contributor to CAVD (41).

Analysis of the top 20 differentially regulated genes between all groups (Figure 5H) yields other interesting gene changes. The top 2 differentially regulated genes, CYP1B1 and CYP1A1, cytochrome p450-family enzymes have been shown to be atheroprotective in endothelial cells (28, 29), were upregulated in all of the dantrolene-treated groups, regardless of treatment with LPC, but not in any of the other groups. The inflammatory regulator CXCL8, which encodes for interleukin-8, was comparatively downregulated in the vehicle control and LPC-treated groups. ANGPTL4, a neovascularization gene with a role in lipid metabolism (42, 43), was upregulated in both LPC treated groups and downregulated in the dantrolene only and vehicle control groups. Other genes that were strongly differential were regulators of DNA replication and repair (TOP2A, STAG2, SMC3, NIPBL) and organelle movement (KTN, EEA1, PCM1, VPS13C), inflammation (SLPI), and oxidative stress (TXNIP). Overall, these results suggest that dantrolene upregulates genes protective of calcification in paVICs while reducing the expression of inflammatory and oxidative genes.

KEGG pathway analysis revealed that the most upregulated genes in all groups were related to viral carcinogenesis and salmonella infection (Figure 6A). These pathways typically involve inflammation, apoptosis regulation, and cell proliferation, which overlap with the phenotypes demonstrated here. Ubiquitin-mediated proteolysis and a number of other signaling pathways appeared in all groups. Uniquely, the LPC-only group had upregulated genes relating to smooth muscle cell contraction, and the LPC + dantrolene group showed upregulation of T_H17 cell signaling and VEGF signaling. When comparing the LPC + dantrolene group to the LPC-only group, few genes were differentially regulated with only 1–3 genes per pathway. Notably enriched pathways involved Jack-STAT signaling, cytochrome p450, and HIF.

Regarding downregulated genes (Figure 6B), the LPC-only group did not show clustering of gene pathways. However, the dantrolene-only group showed HIF1-pathway genes as the most downregulated, a change that persisted in the LPC + dantrolene group. Both dantrolene-treated groups showed a downregulation of genes relating to glycolysis/gluconeogenesis and other sugar metabolism pathways. The dantrolene-only group showed a downregulation of genes pertaining to vascular smooth muscle cell contraction, contrasting with its upregulation in the LPC-only group. The LPC + dantrolene group compared to the LPC-only group again had few pathways that were less enriched (data not shown), but notably genes in pathways relating to HIF, TGF-β, actin contraction, renin-angiotensin signaling, and osteoclast differentiation were downregulated. Overall, these results suggest multifaceted changes in paVICs due to LPC and dantrolene and highlight dantrolene's complex regulation of the HIF1 pathway.