Transcriptomic Analysis of Cr(VI)-induced Changes in C2C12 Cells During Myogenic Differentiation

Abstract

Background

Hexavalent chromium [Cr(VI)] is an environmental toxicant extensively used in a variety of industrial processes including chrome plating, leather tanning, textile manufacturing, aircraft production, and stainless-steel production. Our previous study reported that exposure to Cr(VI) inhibited C2C12 myogenic differentiation in a dose-dependent manner, yet the transcriptional mechanisms underlying Cr(VI)-induced disruption of myogenesis remains poorly understood. This study aimed to characterize the global transcriptional alterations during C2C12 myogenic differentiation and identify molecular pathways disrupted upon Cr(VI) exposure.

Methods

C2C12 cells were differentiated in the presence of 0, 2, or 5 μM of Cr(VI) and collected at differentiation days 0, 1, 2, and 4. Whole transcriptome analysis of a total of 30 samples (with 3 biological replicates per condition) was performed using RNA-sequencing followed by differential gene expression analysis, unsupervised fuzzy c-means clustering, GO biological processes functional annotation, and KEGG pathway enrichment analysis.

Results

Cr(VI) exposure resulted in a massive transcriptomic change in differentiating C2C12 cells. Fuzzy c-means clustering identified 12 distinct gene expression patterns, with Clusters 3, 10, and 12 showing significant overlap with Cr(VI)-regulated genes. Functional enrichment analyses revealed Cr(VI) alters genes involved in early-stage cell cycle regulation and DNA repair as well as terminal differentiation processes like sarcomere organization and muscle contraction. Specifically, Cr(VI) suppressed expression of key structural and contractile genes and disrupted pathways essential for myogenic differentiation, cell cycle regulation and DNA damage repair. Furthermore, Cr(VI) disrupted Hippo signaling by downregulating Tead4 and its downstream myogenic targets such as MyoG, Cav3, Mustn1, and Dysf suggesting a mechanism for impaired differentiation.

Conclusion

This study highlights the widespread alterations of Cr(VI) exposure on transcriptional programs – including structural development, genomic stability, and cell cycle withdrawal – governing muscle development and maturation, offering insight into how Cr(VI) exposure affects skeletal muscle health.

1. Introduction

Hexavalent chromium [Cr(VI)] is classified as a Group 1 human carcinogen by the International Agency for Research on Cancer and is associated with a range of adverse health outcomes, including lung cancer, asthma, hepatic toxicity, gastrointestinal ulcers, as well as skin lesions[1-4]. Cr(VI) and its compounds are extensively used in a diversity of industrial processes such as chrome plating, pigments and dyes, wood preservation, leather tanning, textile manufacturing, aircraft production, and stainless-steel production[5]. Cr(VI) is a major environmental pollutant primarily released into our soil, air, food, and water sources through the improper disposal of Cr(VI)-containing industrial waste[5]. In the 1980s to 1990s, Cr(VI) concentrations in drinking water in Hinkley, California reached as high as 580 μg/L, more than ten times the state’s limit of 50 μg/L, after the Pacific Gas and Electric Company (PG&E) released Cr(VI)-contaminated wastewater in nearby unlined water sources[6]. More recently, a nationwide survey conducted by the Environmental Working Group (EWG) from 2013 to 2015 found detectable levels of Cr(VI) in over 75% of US drinking water samples[7,8]. Interestingly, a 3-month study conducted by the National Toxicology Program reported that male and female rats exposed to Cr(VI) in drinking water had a dose-dependent increase in serum creatine kinase (CK), a marker for muscle injury[9]. Consistently, a cohort study revealed increased serum CK activity in chrome plating workers as compared to those in control subjects[10]. Despite clear evidence of muscle injury, little is known about the impact of Cr(VI) exposure on skeletal muscles.

Skeletal muscles account for approximately 40% of human body weight and contain 50-75% of all body proteins, emphasizing its physiological and functional significance[11]. Myogenic differentiation, an essential process for muscle development and regeneration, comprises multiple steps by which muscle precursor cells (or myoblasts) proceed through irreversible cell cycle arrest to differentiate into myocytes and fuse to form multinucleated myotubes[12]. This process is orchestrated by the basic helix-loop-helix family of transcription factors called myogenic regulatory factors (MRFs), which include Myogenic differentiation factor 1 (MyoD), Myogenin (MyoG), Myogenic factor 5 (Myf5), and Myogenic regulatory factor 4 (Mrf4 or Myf6)[12]. Our previous study demonstrated that Cr(VI) inhibits myogenic differentiation in C2C12 myoblasts in a dose-dependent manner[13]. Exposure to 2 μM of Cr(VI) led to delayed myotube formation, whereas exposure to a higher dose of 5 μM Cr(VI) completely abolished myotube formation. Muscle-specific markers, including Muscle creatine kinase (Mck), Myocyte enhancer factor 2 (Mef2c), Myomaker (Mymk), and Myomixer (Mymx), were also significantly reduced in a dosedependent manner. Additionally, the expression of all four MRFs were significantly altered in Cr(VI)-treated cells. Furthermore, the inhibitory effect of Cr(VI) on myogenic differentiation was also confirmed in primary mouse satellite cells, which are essential for skeletal muscle regeneration. However, the underlying molecular mechanisms and transcriptional programs disrupted by Cr(VI) during myogenesis remain poorly understood.

To further elucidate the mechanisms underlying Cr(VI)’s inhibitory effect on myogenic differentiation, we performed RNA-sequencing analysis on differentiating C2C12 cells in the presence or absence of Cr(VI) treatment. Genes that differentially expressed in Cr(VI)-treated groups versus untreated controls at 3 different time points were identified and further analyzed by pathway enrichment analysis.

2. Methods

2.1. C2C12 culture and differentiation

C2C12 cells were obtained from American Type Culture Collection (ATCC; Rockville, USA) and were maintained in high-glucose DMEM medium (Gibco; NY, USA) supplemented with 20% fetal bovine serum (FBS) and 1% penicillin-streptomycin. To induce myogenic differentiation, cells were switched to differentiation medium consisting of high-glucose DMEM medium supplemented with 2% horse serum and 1% penicillin-streptomycin. For Cr(VI) treatment, differentiating C2C12 cells were exposed to 0, 2, or 5 μM of potassium chromate (K2CrO4) in differentiation medium. Cells were cultured in a humidified incubator at 37°C and 5% CO₂. The differentiation medium was changed every two days.

2.2. RNA extraction and RNA-sequencing

Total RNA was extracted from control or Cr(VI)-treated C2C12 cells using TRIzol reagent (Invitrogen; MA, USA). A total of 30 samples including triplicates per condition were submitted to the NYU School of Medicine Genome Technology Center (RRID: SCR_017929) for library preparation and sequencing using the Illumina TruSeq RNA Library Prep Kit and the Illumina NovaSeq 6000 by multiplexed paired-end runs with 50 cycles, respectively, according to the manufacturer’s instructions. Our sequencing data can be accessed on the Gene Expression Omnibus (GEO) database (GSE294800).

2.3. Differential gene expression analysis

Raw FASTQ sequence data was imported into the CLC Genomics Workbench Version 21.0.4 (Qiagen; MD, USA) for data analysis. Quality control was performed by trimming low-quality reads and reads containing adapters then the trimmed reads were aligned to the mouse reference genome (Mus musculus ensemble V80) allowing two mismatches. Only uniquely mapped reads were retained for downstream analysis. Read counts were normalized and reported as counts per million (CPM) for downstream differential expression analysis. Differential gene expression analysis was performed using the Advanced RNA-Seq plugin in CLC by comparing each treated group with the corresponding control group. TMM (trimmed mean of M values) normalization was performed to adjust library sizes before differential expression analysis. Genes with a false discovery rate (FDR)<0.05 between control and treated group and a fold-change ∣FC∣>2 were considered significant differentially expressed genes. Volcano plots, Venn diagrams, UpSet plots, bubble plots, and heatmaps with hierarchical clustering were generated using OmicStudio (https://www.omicstudio.cn/tool), a web-based bioinformatics platform for data visualization and analysis. Heatmaps were generated using Hierarchical clustering with Euclidean distance to visualize the z-score normalized gene expression. The z-scores were calculated based on the mean and standard deviation of gene expression across all samples for each gene. The expression fold changes were visualized as paired dot plots using GraphPad Prism (v10.4.0).

2.4. ‘Mfuzz’ cluster analysis

To identify gene expression patterns in untreated C2C12 myogenic differentiation, a fuzzy c-means clustering analysis was performed using the ‘mfuzz’ R package (version 2.50.0) with the minimum fuzzifier parameter set to 2 and the standard fuzzification algorithm was used. The mean CPM values of each gene for untreated differentiation day 0, 1, 2, and 4 were used to generate 12 distinct clusters with a minimum threshold value of 0.25.

2.5. Functional annotation and pathway enrichment analysis

To identify biological processes and pathways associated with Cr(VI)-regulated genes in each cluster, Gene Ontology (GO) functional annotation for biological processes (BP) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses were performed using Database for Annotation, Visualization, and Integrated Discovery (DAVID) v2022q3 (https://david-d.ncifcrf.gov/)[14]. Significantly enriched BPs and KEGG pathways were identified using a p-value threshold of <0.05 and corrected via the Benjamini-Hochberg method. BP and KEGG visualizations were generated using GraphPad (v10.4.0).

2.6. RT-qPCR

Total RNA was extracted using TRIzol reagent (Invitrogen; MA, USA) according to the manufacturer’s instructions. RNA concentrations were measured using a NanoDrop-2000 spectrophotometer (Thermo Scientific; MA, USA). Then, cDNA was synthesized using the LunaScript RT SuperMix (NEB; MA, USA) following the manufacturer’s protocol. RT-qPCR was performed on a QuantStudio 6 PCR machine (Applied Biosystems; MA, USA). Relative mRNA expression levels were calculated using 2^-ΔΔCt and Gapdh was used as a reference gene.

2.7. Western blot analysis

Cells were briefly washed with cold phosphate buffer solution (PBS; Gibco; NY, USA) then homogenized in boiling lysis buffer consisting of 1% sodium dodecyl sulfate (SDS), 1mM sodium ortho-vanadate (Na3VO4), 10mM Tris-HCl (pH 7.4). Protein concentrations were quantified using a NanoDrop-2000 spectrophometer (Thermo Scientific; MA, USA). Whole cell lysates were separated on a 12% SDS-PAGE gel then transferred to a nitrocellulose membrane (Bio-Rad; CA, USA). Membranes were blocked in 5% milk in tris-buffered saline (TBS) at room temperature for 1 hour then incubated with primary antibodies overnight in 4°C. The primary antibodies used included: p21 (Santa Cruz; #sc6246), PCNA (Santa Cruz; #sc56), and H3S10ph (Cell Signaling; #cs9701). The next day, membranes were washed with TBS containing 0.1% Tween-20 (TBST) three times for 5 minutes each then incubated in secondary antibody at room temperature for 1h. The secondary antibodies used were goat-anti-rabbit IgG-h+l HRP-conjugated antibody (Bethyl; #A120-101P) and goat-anti-mouse IgG-h+l HRP-conjugated antibody (Bethyl; #A90-116P). Membranes were imaged using SuperSignal^™ West Pico PLUS Chemiluminescent Substrate (Thermo Fisher; MA, USA, #34577) with the Odyssey^® XF Imaging System (LI-COR Biosciences; NE, USA). GAPDH (Proteintech; #pt60004–1-AP) was used as a loading control.

2.8. Statistical Analysis

Statistical analysis was conducted using Graphpad Prism 10 (GraphPad Software Inc., CA, USA). Data were generated from two independent experiments (each with three replicates) and presented as mean ± SD. Significance was calculated by either two-way ANOVA or unpaired t-test with Welch’s correction, and marked as follows: ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05. Results with a p value greater than 0.05 are not statistically significant (ns).

3. Results

3.1. Cr(VI) induces dose- and time-specific changes in gene expression during C2C12 myogenic differentiation

Our previous study indicated that Cr(VI) exposure inhibits myogenic differentiation of C2C12 myoblasts[13]. To determine genes or pathways that mediate Cr(VI)-induced defective myogenic differentiation, RNA-sequencing was performed to profile transcriptomic changes in differentiating C2C12 cells in response to Cr(VI) treatment. C2C12 cells were induced to undergo myogenic differentiation in the presence of 0, 2, or 5 μM Cr(VI) (Cr0, Cr2, and Cr5) and harvested on differentiation days 0, 1, 2, and 4 (D0, D1, D2, and D4). A total of 30 samples were analyzed with 3 biological replicates for each condition (Fig. 1A). Principal component analysis (PCA) was conducted to evaluate the overall transcriptomic variation across the 30 RNA-sequencing samples. As shown in Fig. 1B, biological replicates clustered tightly within each group, while distinct clustering of the 30 samples was observed based on differentiation day and Cr(VI) concentration, reflecting the significant impacts of these factors on transcriptional profiles. These results suggest that Cr(VI) treatment alters the normal patterns of gene expression observed during C2C12 myogenic differentiation in a dose-dependent and time-specific manner.

Fig.1. Transcriptomic profiling of C2C12 myoblast differentiation under Cr(VI) exposure reveals dose- and time-dependent gene expression changes.

(A) Experimental workflow of differentiating C2C12 cells treated with 0, 2, or 5 μM of Cr(VI) (Cr0, Cr2, and Cr5). Total RNA was collected at 0, 1, 2, and 4 days after differentiation (D0, D1, D2, and D4) for library preparation and RNA-sequencing (n=3 replicates per condition). (B) Principal component analysis plot showing replicate samples have tight clustering patterns based on both timepoint and treatment conditions indicating high internal consistency and reproducibility of the RNA-seq data. PC1, the horizontal axis, has the most variation (18.7%), while PC2 (the vertical axis) capture the second most variation (13.9%). (C) Bar plot summarizing the number of differentially expressed genes between Cr(VI)-treated and untreated groups at each differentiation day. Each comparison was made relative to the time-matched untreated control (e.g., Cr2D4 vs. Cr0D4). Red represents upregulated and blue represents downregulated genes. (D) Volcano plots visualizing differentially expressed genes between 5 μM Cr(VI)-treated and untreated C2C12 myoblasts at D1 (left panel), D2 (middle panel), and D4 (right panel). Blue dots represent significantly downregulated genes, red dots represent significantly upregulated genes, and grey dots represent insignificant genes.

Next, we focused on identification of Cr(VI)-induced gene expression changes in differentiating C2C12 using two complementary approaches. First, gene expression in each treatment group was compared to those in untreated and undifferentiated C2C12 cells (Cr0D0). Genes with a false discovery rate (FDR) < 0.05 between each treatment group and untreated control (Cr0D0) and an absolute fold change equal to or greater than 2 were defined as differentially expressed genes. This generated a total of nine gene lists (Cr0D1, Cr0D2, Cr0D4, Cr2D1, Cr2D2, Cr2D4, Cr5D1, Cr5D2, Cr5D4). The number and direction of differentially expressed genes (up- or down-regulated) in each list are summarized in Supplementary Fig. 1A. The number of differentially expressed genes slightly increased over the course of differentiation; however, the general trends were consistent across both the untreated (0 μM) and Cr(VI)-treated groups (2 and μ5 M). Second, gene expression in Cr(VI)-treated groups was compared to those in time-matched untreated controls, and yield six gene lists: Cr2D1 vs. Cr0D1, Cr2D2 vs. Cr0D2, Cr2D4 vs. Cr0D4, Cr5D1 vs. Cr0D1, Cr5D2 vs. Cr0D2, and Cr5D4 vs. Cr0D4 (Fig. 1D and Supplementary Table 2). Interestingly, the number of differentially expressed genes increased in a dose- and time-dependent manner (Fig. 1D). Compared to the relatively small number of genes changed in cells exposed to 2 μM Cr(VI) (Supplementary Fig. 1B), 5 μM of Cr(VI) treatment resulted in a drastic increase in the number of differentially expressed genes across all timepoints (Fig. 1E). At day 1 of differentiation, 469 differentially expressed genes were identified, with 257 significantly upregulated and 212 significantly downregulated. By day 2, the number of differentiallu expressed genes increased substantially to 1035 genes, including 220 upregulated and 815 downregulated. By day 4, the number of differentially expressed genes further increased to 3295 differentially expressed genes, consisting of 1341 upregulated and 1954 downregulated genes (Fig. 1E). The dose-and time-specific changes in gene expression are consistent with our previous study, in which 2 μM Cr(VI) exposure delayed myotube formation whereas 5 μM Cr(VI) treatment nearly abolished myogenic differentiation in C2C12 cells[13]. Analysis of genes commonly altered in both Cr2D4 and Cr5D4 samples, including 109 upregulated and 112 downregulated genes, further supports a dose-dependent effect of Cr(VI) exposure on gene expression (Supplementary Fig. 2). Interestingly, the expression of the 112 commonly downregulated genes exhibited a significant dose-dependent reduction in 5 μM-treated cells compared to that in 2 μM-treated cells (Supplementary Fig. 2B). In contrast, the expression levels of the 109 commonly upregulated genes did not significantly differ between two Cr(VI) concentrations (Supplementary Fig. 2C), suggesting that Cr(VI) may exert a stronger transcriptional suppression in differentiating C2C12 cells in a dose-dependent manner. Thus, we focused on differentially expressed genes from the 5μM Cr(VI) treatment group for subsequent functional annotation and pathway analysis.

3.2. Clusters 12, 10, and 3 encompass the largest number of Cr(VI)-regulated genes

To characterize the transcriptional dynamics during normal C2C12 myogenic differentiation, we performed unsupervised fuzzy c-means clustering of gene expression profiles from untreated samples collected at differentiation days 0, 1, 2, and 4 (Cr0D0, Cr0D1, Cr0D2, Cr0D4) using the ‘Mfuzz’ package, a tool optimized for soft clustering of time-series gene expression data. This analysis identified a total of 12 distinct gene clusters, each representing a unique temporal expression pattern across the differentiation timeline (Fig. 2A). Clusters 1 and 10 exhibited a continuous decrease or increase in gene expression, respectively, throughout the differentiation process. Genes in these clusters were enriched for functions related to chromatin remodeling, intracellular signaling, transcription regulation, and protein transport, reflecting continuous transcriptional reprogramming to accommodate metabolic and functional demands during myogenesis (Supplementary Fig. 3). Clusters 3, 4, and 9 displayed a sharp change in expression at day 1 that remained stable through days 2 and 4. These clusters were enriched for cell cycle regulation, DNA damage response, mitotic checkpoint signaling, and autophagy – biological processes consistent with cell cycle exit and commitment to the myogenic lineage. Clusters 5 and 6 showed transient upregulation or downregulation at day 1, followed by a gradual return to baseline expression levels by day 4. These clusters likely include genes that are transiently required for initiating differentiation but are dispensable during later stages. Functional enrichment revealed biological processes such as vesicle-mediated transport, lysosome organization, translation, and mitochondrial activity. Clusters 7, 8, and 11 exhibited transient activation on day 1, 2, or 4, respectively. The dynamic expression of these genes suggests stage-specific roles in immune signaling, metabolic reprogramming, and extracellular matrix remodeling. Finally, Clusters 2 and 12 showed minimal changes at day 1 but either progressively decreased (Cluster 2) or increased (Cluster 12) expression at days 2 and 4. Cluster 12 was enriched for genes involved in sarcomere organization, mitochondrial ATP synthesis, and muscle contraction – hallmark processes of terminal differentiation and myofiber maturation (Supplementary Fig. 3). Collectively, these 12 expression clusters delineate the tightly coordinated and stage-specific transcriptional changes underlying C2C12 myogenic differentiation.

Fig. 2. Temporal clustering of gene expression during C2C12 differentiation and overlap with differentially expressed genes altered by Cr(VI) treatment.

(A) Profiling gene expression changes of differentiating C2C12 cells into 12 clusters using the fuzzy c-means clustering algorithm. Each plot represents a cluster of genes with similar expression profiles. The x-axis denotes differentiation timepoints (D0, D1, D2, and D4) while the y-axis denotes normalized expression changes. The black line represents the average expression trend and individual gene trajectories are shown in colored lines. (B-D) Three UpSet plots visualizing the interactions between 12 gene clusters and differentially expressed genes induced by 5 μM Cr(VI) at D1 (B), D2 (C), and D4 (D). The left bar plot represents the set size, indicating the number of genes in each cluster as well as differentially expressed genes at each day of myogenic differentiation. The top bar plot represents the intersection size, displaying the number of genes in each specific interaction. The dot matrix below specifies the composition of the interaction, with a single dot representing genes unique to a specific set and connected dots for genes shared between two sets.

To determine how Cr(VI) disrupts distinct transcriptional programs during C2C12 myogenic differentiation, we conducted an intersection analysis between differentially expressed genes identified in the 5 μM Cr(VI)-treated groups and genes within the 12 previously defined gene expression clusters. Three UpSet plots were generated to visualize the overlap between cluster-specific genes and Cr(VI)-regulated genes at each differentiation timepoint (Fig. 2B-D). At day 1 (D1), Cr(VI) exposure predominantly affected genes within Cluster 12, Cluster 10, and Cluster 8 (Fig. 2B). By day 2 (D2), the greatest overlap was observed in Cluster 12, Cluster 3, and Cluster 10 (Fig. 2C). On day 4 (D4), Cr(VI) induced the most extensive transcriptional alterations, with substantial overlap in Cluster 12, Cluster 3, and Cluster 1 (Fig. 2D). Thus, the top 3 clusters that were most affected by Cr(VI) treatment are Clusters 12, 10 and 3. While genes in Clusters 10 and 12 exhibited progressive upregulation over the course of differentiation, Cluster 3 genes demonstrated a marked downregulation beginning at day 1 that persisted through day 4, suggesting that Cr(VI) alters gene expression at both early and late stages of myogenic differentiation.

3.3. Functional annotation of Cr(VI)-regulated genes in Clusters 12, 10, and 3

Next, we performed a functional annotation and pathway analysis on Cr(VI)-regulated genes in top 3 clusters (Cluster 10, 12 and 3). Among these three clusters, Cluster 12 was the most affected by Cr(VI) treatment, as evidenced by the highest number of genes altered by Cr(VI) – 99 at D1, 269 at D2 and 469 at D4 (Fig. 3A). Cluster 12 genes showed minimal change between D0 and D1 followed by a marked increase from day 1 to day 4. Gene Ontology (GO) functional annotation of genes altered by Cr(VI) treatment revealed that the highly enriched biological processes are critical for myogenic differentiation and muscle function, including sarcomere organization, muscle contraction, calcium ion transport, myofibril assembly, and actin filament organization. Consistently, the pathways associated with muscle structure, function and metabolism were highly represented in Cluster 12 Cr(VI)-regulated genes, including cytoskeleton in muscle cells, motor proteins, glucagon signaling, calcium signaling, and oxytocin signaling pathways. Cluster 10 showed a similar upregulation as Cluster 12 but exhibited a progressive increase starting from day 1, with 44 Cr(VI)-regulated genes in D1, 82 in D2, and 318 in D4 (Fig. 3B). The GO biological processes enriched for these differentially expressed genes included skeletal muscle differentiation, cell-cell adhesion, protein transport, and establishment of cell polarity – hallmarks of early myogenic signaling like structural remodeling. KEGG pathway analysis revealed an enrichment for cytoskeleton in muscle cells, like Cluster 12, as well as multiple pathways related to cancer. In contrast to Clusters 10 and 12, Cluster 3 genes displayed a sharp downregulation in D1 and maintained low expression until the end of differentiation, with 27 differentially expressed genes in D1, 248 in D2, and 442 in D4 (Fig. 3C). Cr(VI)-regulated genes in Cluster 3 were significantly enriched for biological processes involved in the cell cycle and genomic stability, such as cell division, chromosome segregation, DNA repair, mitotic spindle assembly checkpoint signaling, and DNA replication. This is consistent with early downregulation of genes required for cell cycle exit and commitment. KEGG pathways enriched for these differentially expressed genes included the cell cycle, DNA replication, and multiple DNA repair pathways such as homologous recombination, base excision repair, and mismatch repair. Overall, these results demonstrate that Cr(VI) disrupts key transcriptional programs required for both the initiation and maturation phases of myogenic differentiation by targeting genes involved in muscle structure development, muscle function and cell cycle regulation.

Fig. 3. Functional annotation and pathway enrichment analyses of Cr(VI)-regulated genes in Clusters 12, 10 and 3.

Cr(VI)-regulated genes from Cluster 12 (A), Cluster 10 (B) and Cluster 3 (C) were identified at each timepoint (left) and analyzed for biological processes (middle) and KEGG pathways (right). For the bar graphs, the x-axis denotes enrichment score calculated as - log10(p-value). For KEGG dot plots, the dot size represents the gene count, and the color bar represents the -log10(p-value).

3.4. Cr(VI) downregulates genes involved in muscle structure development and contraction

Given that Cluster 12 was the most affected by Cr(VI) treatment and was enriched for genes involved in muscle function, we next examined the expression patterns of genes related to sarcomere organization and muscle contraction. In C2C12 myogenic differentiation, sarcomere organization is critical for assembling the contractile apparatus while muscle contraction-related gene activation reflects the functional maturation of myotubes into contractile myofibers. Proper formation of sarcomeres – a repeating unit composed of actin (thin) and myosin (thick) filaments – is necessary for establishing myofibril alignment, force transmission, and overall muscle integrity[15]. Concurrently, the expression of muscle contraction genes ensures that differentiated myotubes acquire the machinery needed to respond to calcium signaling and execute contractile function.

To assess Cr(VI)’s impact on these processes, heatmaps were generated to visualize the expression dynamics of genes involved in sarcomere organization (Fig. 4A) and muscle contraction (Fig. 4B) across four timepoints (D0, D1, D2, and D4) under control (0 μM) and Cr(VI)-treated conditions (2 μM and 5 μM). In untreated cells, these genes exhibited a robust, time-dependent upregulation from D0 to D4, consistent with progressive differentiation and maturation of C2C12 cells. In contrast, Cr(VI) exposure resulted in a dose-dependent suppression of this transcriptional activation. At 2 μM Cr(VI), many genes failed to reach full induction, while 5 μM Cr(VI) nearly abolished the expected upregulation of structural and contractile genes. Notably, sarcomere-related genes such as Plec, Mybpc1/2, Tnnt2, Myom1–3, and Capn3—which are essential for myofibril alignment, sarcomere stabilization, and Z-line organization—showed markedly reduced expression in Cr(VI)-treated groups. Similarly, muscle contraction genes including Tnnt1–3, Tnni1, Tnnc1, Tpm1/2, and Myh1-4, Myh7, Myh8, which mediate calcium sensitivity and force generation through actin-myosin interactions, were significantly downregulated in response to Cr(VI). This includes the coordinated troponin-tropomyosin (Tnnt1, Tnnt2, Tnnt3, Tnni1, Tnnc1, Tpm1, Tpm2, Tpm3) complex, which plays a key role in regulating myosin binding to actin in a calcium-dependent manner, and myomesins (Myom1, Myom2, Myom3), which anchor thick filaments at the M-line and maintain sarcomeric integrity. RT-qPCR validation revealed that treatment with 5 μM Cr(VI) significantly downregulated Tnnt1, Tnnt2, Myom2, Myom3, Myh1, and Myh3, in a time-dependent manner (Fig. 4C). These findings indicate that Cr(VI) disrupts the coordinated transcriptional upregulation of sarcomere and contractile genes required for functional myotube formation. The dose-dependent inhibition of these gene programs suggests that Cr(VI) exposure impairs both the structural assembly and functional maturation of muscle cells, ultimately compromising skeletal muscle integrity and regeneration capacity. This disruption of sarcomere and contractile gene expression may underlie the broader inhibitory effects of Cr(VI) on myogenic differentiation and muscle tissue function.

Fig. 4. Cr(VI) downregulates genes essential for sarcomere organization and muscle contraction.

(A–B) Heatmap of genes related to sarcomere organization (A) and muscle contraction (B) across all differentiation timepoints and Cr(VI) treatment conditions. The color bar represents the z-scores of normalized expression value. Red represents a positive z-score (upregulation) and blue indicates a negative z-score (downregulation), relative to the mean expression of each gene. (C) Relative mRNA expression levels of Tnnt1, Tnnt2, Myom2, Myom3, Myh1, or Myh3 at each day of differentiation with or without Cr(VI) treatment. All RT-qPCR data are normalized to Gapdh. Data are represented as mean ± SD (n=3). Statistical significance was determined using a two-way ANOVA test compared to the untreated control for each timepoint (ns, no significant, ***p<0.001, and ****p<0.0001).

3.5. Cr(VI) modulates genes involved in DNA repair and cell cycle progression

In contrast to Clusters 12 and 10, Cluster 3 exhibited an early and sustained downregulation pattern with genes significantly enriched for DNA repair and cell cycle processes, both of which are critical pathways that preserve genomic integrity during the switch from cell proliferation to differentiation. During early stages of C2C12 myogenesis, effective DNA repair is essential to resolve DNA damage and ensure genomic stability as cells exit the cell cycle and commit to terminal differentiation. Heatmaps were generated to visualize the temporal expression of DNA repair genes under control and Cr(VI)-treated conditions (Fig. 5). In the control (0 μM) group, genes involved in homologous recombination, mismatch repair, and base excision repair – including Brca1, Rad51, Exo1, Lig1, Pole, Pold1, Blm, Chaf1b, and Ercc6l – were highly expressed in proliferating myoblasts (D0) but sharply downregulated upon differentiation (D1-D4), consistent with cell cycle exit as cells transition from proliferation to differentiation. However, exposure to Cr(VI) resulted in a dose-dependent repression of these genes, with the 5 μM treatment group exhibiting further downregulation across all timepoints (Fig. 5A). This suggests that Cr(VI) compromises the cellular response to DNA damage, increasing genomic instability during the differentiation process. Given the integral role of DNA repair in coordinating cell cycle checkpoints, we next assessed the impact of Cr(VI) on cell cycle progression. Heatmap analysis revealed that cell cycle-related genes regulating cell cycle progression, DNA replication, and mitosis – including Cdc6, Mcm2-7, Ccnb1, Ccne1, Cdk1, Cdc20, and E2f1 – were robustly downregulated during normal differentiation, consistent with cell cycle withdrawal. However, Cr(VI) treatment further decreased these genes (Fig. 5B). Interestingly, several genes involved in regulating cell cycle checkpoints – such as Wee1, Chek1, Trp53, Ywhah, Ywhab, Ywhaq, and Ywhaz – were upregulated by Cr(VI), indicating checkpoint activation as a stress response to increased DNA damage. To confirm the impact of Cr(VI) on cell cycle regulation at the protein level, we conducted western blot analysis of p21(Cdkn1a), PCNA, H3S10ph in C2C12 cells differentiated for 4 days in the presence or absence of 5 μM of Cr(VI) (Fig. 5C). p21, a cyclin-dependent kinase (CDK) inhibitor that promotes cell cycle withdrawal and differentiation, was markedly decreased at all timepoints in Cr(VI)-treated cells. Proliferating cell nuclear antigen (PCNA), a replication and proliferation marker, remained elevated with Cr(VI) treatment after four days of differentiation, indicating a failure for cell cycle exit. Additionally, phosphorylation of histone H3 at serine 10 (H3S10ph), a mitotic marker for chromatin condensation – was further decreased by Cr(VI) in day 1 and completely abolished in day 2 and 4, suggesting impaired mitotic progression. In addition, expression of genes selected from both categories in D4 differentiated cells with or without 5 μM Cr(VI) was further validated by RT-qPCR. Consistent with RNA-seq results, genes involved in DNA repair, such as Rad51, Tdp1, Fanci, Fancd2, Ddx11, and Rfc2, were significantly downregulated in Cr(VI) treated cells, while genes related to the cell cycle, including Chek1, Chek2, and Wee1, were upregulated by Cr(VI) (Fig. 5D). Taken together, these findings demonstrate that Cr(VI) contributes to genomic instability and impaired myogenic progression as indicated by downregulated DNA repair pathways, checkpoint activation, and deficient cell cycle withdrawal.

Fig. 5. Cr(VI) dysregulates genes involved in DNA repair and cell cycle control.

(A-B) Heatmap of genes involved in DNA repair (A) and Cell cycle (B) across all differentiation timepoints and Cr(VI) treatment conditions. The color bar represents the z-scores of normalized expression value. Red represents a positive z-score (upregulation) and blue indicates a negative z-score (downregulation), relative to the mean expression of each gene. (C) Western blot analysis of p21, PCNA, and H3S10ph protein levels in C2C12 cells differentiated for 4 days in the absence or presence of 5 μM of Cr(VI) treatment. GAPDH was used as a control. (D) Expression of genes involved in DNA repair and cell cycle (Rad51, Tdp1, Fanci, Fancd2, Ddx11, Rfc2, Trp53, Chek1, Chek2, E2f1, Wee1) at D4 of C2C12 differentiation with or without Cr(VI) treatment. All RT-qPCR data are normalized to Gapdh. Relative mRNA expression shown as fold change relative to untreated and undifferentiated control (Cr0D0). Data are represented as mean ± SD (n=3). Statistical significance between Cr5D4 and Cr0D4 was determined using an unpaired t-test with Welch’s correction. (ns, no significant, *p<0.05, **p<0.01, ***p<0.001).

3.6. Cr(VI) Disrupts Hippo Signaling by Suppressing Tead Transcriptional Activity and Downregulating Myogenic Targets

To determine which signaling pathway was affected by Cr(VI), KEGG pathway enrichment analysis was performed with the genes identified in Cluster 3 (n=2069), Cluster 10 (n=1362), and Cluster 12 (n=1603), the top three clusters altered by Cr(VI) exposure. As shown in Fig. 6A, while Cluster 3 exhibited significant enrichment in the p53 pathway and the TNF pathway, both of which are crucial for regulating cell growth and death, Cluster 10 and Cluster 12 displayed the common enrichment in major pathways involved in cell metabolism, including mTOR pathway, AMPK pathway, FoxO pathway, and insulin signaling pathway. Interestingly, Hippo signaling, a pathway important for cell proliferation, differentiation, and regeneration, was enriched in all three clusters, suggesting its potential role in Cr(VI)-induced defective myogenic differentiation.

Fig. 6. Cr(VI) exposure disrupts Hippo signaling and impairs muscle-related genes in C2C12 cells.

(A) Grouped bubble plot visualizing significant KEGG signaling pathways enriched in the total genes identified in Cluster 3, 10, and 12. The dot size represents the gene count enriched in the indicated KEGG pathway, and the color bar represents the -log10(p-value). (B) Heatmap of key components in the Hippo signaling pathway in C2C12 cells at all differentiation timepoints. The color bar represents the z-scores of normalized expression value. Red represents a positive z-score (upregulation) and blue indicates a negative z-score (downregulation), relative to the mean expression of each gene. (C) RT-qPCR validation of Lats1, Tead1, Tead3, and Tead4 in untreated and treated C2C12 cells at differentiation days 0, 1, 2, and 4. (D) RT-qPCR validation of muscle-related Tead4 downstream targets MyoG, Mustn1, Dysf, and Cav3 in Cr(VI)-treated and -untreated C2C12 cells across differentiation days 0, 1, 2, 3, and 4. All RT-qPCR data are normalized to Gapdh. Data are represented as mean ± SD (n=3). Statistical significance was determined using a two-way ANOVA test compared to the untreated control for each timepoint (*p<0.05 and ****p<0.0001).

Hierarchical clustering of key components of the Hippo signaling pathway across all samples revealed distinct expression patterns during differentiation in response to Cr(VI) exposure (Fig. 6B). Based on their expression in untreated differentiating C2C12 cells, genes involved in Hippo signaling were categorized into three major groups. Group 1 are those downregulated upon myogenic differentiation but increased by Cr(VI) exposure, including Yap1, Lats1, and the members of the 14-3-3 protein family (Ywhab, Ywhaq, Ywhaz). Group 2 displayed a timedependent upregulation during myogenic differentiation, including Tead1, Tead3, Tead4 and Mustn1. These gene failed to be upregulated in the presence of Cr(VI). Genes in Group 3, such as Lats2, Sav1, Fbxw11 and Mob1b, were not significantly changed during myogenic differentiation. However, 5 μM of Cr(VI) treatment significantly upregulated these genes in a dose-dependent manner (Fig. 6B). Taken together, Cr(VI) exposure downregulated genes that were upregulated during myogenic differentiation but upregulated genes that were downregulated or not changed in myogenic differentiation.

Cr(VI)-induced upregulation of Lats1 and downregulation of Tead1, Tead3, and Tead4 were further verified by RT-qPCR (Fig. 6C). Lats1 (large tumor suppressor kinase 1) is serine/threonine kinase that phosphorylates and inactivates YAP/TAZ activity. The TEAD (TEA domain) transcription factors (Tead1-4) are key components that coordinate with YAP/TAZ to activate muscle-specific genes. Among these, Tead1 and Tead4 are predominantly expressed in skeletal muscles and are important in C2C12 myogenic differentiation[16,17]. Both Tead1 and Tead4 exhibited a dose-dependent decrease in expression at all differentiation timepoints with 5 μM of Cr(VI) treatment, with Tead4 showing nearly complete suppression of mRNA levels for the entire 4 days of differentiation. Tead4 has been shown to directly regulate myogenic factors such as MyoG, Cav3, Mustn1, and Dysf[18]. MyoG is an important myogenic regulatory factor essential for differentiation of myoblasts into myocytes. Cav3 is a key component of the muscle cell membrane involved in maintaining structural integrity and muscle function. Mustn1 plays a role in muscle regeneration and is upregulated during repair of damaged muscle tissues. Lastly, Dysf is important for the stability and maintenance of the muscle membrane during contraction. Our results showed that all four genes were upregulated throughout the entire course of myogenic differentiation in the absence of Cr(VI) but Cr(VI) treatment at 5 μM abolished the upregulation of these genes at all timepoints (Fig. 6D). Taken together, these results indicate the dysregulation of Hippo signaling by Cr(VI) in differentiating C2C12 cells, possibly through Tead inhibition.

4. Discussion

Increasing evidence suggests that Cr(VI) exposure may possess an adverse impact on muscle development and function[13,19,20]. Our previous study reported that Cr(VI) inhibits myogenic differentiation in C2C12 cells in vitro and primary mouse myoblasts ex vivo [13]. However, the molecules and pathways mediating Cr(VI) inhibitory effect remain less understood. In the present study, we conducted the first whole transcriptomic profiling of C2C12 cells undergoing myogenic differentiation in the presence of 0, 2, or 5 μM of Cr(VI). Our results revealed that Cr(VI) treatment led to massive changes in gene expression across all differentiation timepoints. The genes associated with muscle structure and function as well as DNA repair and cell cycle regulation are significantly altered in Cr(VI) exposed samples in a dose and time-specific manner. The results align with our previous finding that Cr(VI) treatment inhibited C2C12 differentiation and myocyte fusion in a dose-dependent manner[13].

Numerous studies have reported the transcriptomic changes along the course of myogenic differentiation[21-23]. Consistent with these existing reports, untreated C2C12 cells in our study showed a strong upregulation of well-established myogenic factors MyoD, MyoG, Myf6, Mef2a, Mef2c, and Mef2d[24-26] as well as myofiber components Mb, Myl1, Myl4, Myh3, Myh8, Mybph, Ckm, Neb, and Des[27-29] with differentiation (Fig. 4). In addition, cell proliferation genes Myf5, Cdk1, Cdk4, Cdk6, Ccnd1, and Ccne1 [29-32]were progressively downregulated with myogenesis, consistent with previously reported trends (Fig. 5). Thus, our transcriptomic data of untreated C2C12 cells is largely consistent with previously reported RNA-seq analyses.

The most affected genes in Cr(VI) exposed cells are those involved in sarcomere organization and muscle contraction (Fig. 3 and 4). Sarcomere organization is the assembly and alignment of the basic contractile units of striated muscles called sarcomeres – which are composed of actin (thin) and myosin (thick) filaments along with structural proteins such as tropomyosins (Tpm1, Tpm2, Tpm3, Tpm4), troponins (Tnnt1, Tnnt2, Tnnt3, Tnni1, Tnnc1), myomesins (Myom1, Myom2, Myom3), titin (Ttn), and α-actinin (Actn1, Actn2, Actn3, Actn4) [15,33,34]. These genes form the structural framework necessary to enable coordinated muscle contraction, which is a hallmark of terminal myogenic maturation. In untreated C2C12 cells, we observed a robust, timedependent upregulation of these genes consistent with their established roles in sarcomere assembly and myotube function. However, Cr(VI) exposure suppressed this normal transcriptional activation in a dose-dependent manner (Fig. 4). At 2 μM, many of these genes failed to reach full induction while their upregulation was almost completely abolished with 5 μM of Cr(VI) treatment. Disruption of these genes has been shown to impair myogenesis. For instance, treatment of C2C12 cells with 17β-estradiol inhibited myogenesis accompanied by a decrease in myosin heavy chain (MHC), tropomyosin, and MYOG protein levels[34]. Similarly, exposure to 50 μM of eicosapentaenoic acid or docosahexaenoic acid both suppressed C2C12 differentiation and RNA-sequencing analysis further revealed that muscle-specific genes like insulin-like growth factor 2 (Igf2), troponin T3 (Tnnt3), myoglobin (Mb), myosin light chain phosphorylatable fast skeletal muscle (Mylpf) and myosin heavy polypeptide 3 (Myh3) were downregulated[35]. In zebrafish embryos and larvae, copper (Cu²⁺)- stressed zebrafish embryos reduced locomotor speed and decreased myofibrils in skeletal muscles as seen by the downregulation of muscle fiber markers mylpfa (Mylpf) and smyhc1l (Myh7)[36]. These findings collectively underscore the importance of structural and contractile proteins in proper myofiber development. Overall, our results indicate that Cr(VI) impairs both sarcomere assembly and contractile machinery development, compromising the structural integrity and functional capacity of mature myofibers.

In addition to muscle-related genes and pathways, Cr(VI) exposure significantly disrupted pathways involved in cell cycle regulation and DNA repair – two critical processes that are tightly coordinated with myogenic progression and cellular integrity during differentiation. Proper regulation of the cell cycle is essential for myoblasts to exit proliferation and commit to terminal differentiation. A central mediator of this transition is Cdkn1a (p21), a cyclin-dependent kinase (CDK) inhibitor that facilitates cell cycle withdrawal by inhibiting key regulators of DNA replication, including proliferating cell nuclear antigen (PCNA)[37,38]. In normal C2C12 differentiation, p21 expression is low in undifferentiated myoblasts and increases upon induction of differentiation (peaking in day 1 and 2) then declines as cells reach terminal differentiation by day 4 [37,39-41]. In our study, p21 protein levels in untreated cells followed this expected trend; however, 5 μM of Cr(VI) exposure led to an accelerated and sustained decrease in p21 levels (Fig. 5C). Given that p21 is transcriptionally regulated by MyoD[41] – a factor that we previously showed to be suppressed by Cr(VI) in a dose-dependent manner[13] – these findings suggest that Cr(VI) may impair cell cycle withdrawal by interfering with MyoD-mediated p21 induction. Consistently, PCNA expression remained abnormally high in Cr(VI)-treated cells throughout day 3 and 4 of differentiation, suggesting persistent activation of the DNA replication machinery and failure to properly exit the cell cycle (Fig. 5C). This finding aligns with observations in other studies reporting an inhibitory effect on C2C12 cells with reduced cardiac lineage protein 1 (CLP1+/−)[42] or treated with excretory-secretory products from muscle larvae of Trichinella spiralis (an intracellular parasite of skeletal muscles)[43], both of which maintained high levels of PCNA expression. In addition to cell cycle regulation, multiple pathways involved in DNA damage repair were highly enriched in Cr(VI)-regulated genes, including homologous recombination, base excision repair, mismatch repair and the Fanconi anemia pathway (Fig. 3C and Fig. 5). Cr(VI) is a known inducer of genotoxic stress including DNA base modifications, single- and double-strand breaks, and Cr-DNA adducts [44-48]. Brca1, Rad51, and Rad51ap1, play critical roles in the homologous recombination repair (HRR) pathway that resolves DNA double-strand breaks[48-50]. In untreated cells, these genes exhibited high expression in undifferentiated myoblasts that declined as differentiation progressed. However, Cr(VI) treatment led to an even greater suppression of HRR genes (Fig. 5A). This suggests that Cr(VI) may compromise the ability of differentiating myoblasts to resolve DNA damage, thereby increasing susceptibility to genomic instability. These findings are consistent with a study that showed prolonged exposure to particulate Cr(VI) in immortalized human lung fibroblast cells (WTHBF-6 cells) induced a dose-dependent increase in double-strand breaks and a corresponding decrease in RAD51-mediated repair capacity[51]. Collectively, these results suggest that Cr(VI)-induced suppression of cell cycle inhibitors and DNA repair machinery disrupts the coordination between differentiation and genome stability, thereby contributing to impaired myogenesis.

It is interesting that Hippo signaling is one of top signaling pathways affected by Cr(VI) exposure. Hippo signaling coordinates both muscle-specific gene expression and cell cycle control during skeletal muscle differentiation[52]. Several studies have shown that the Hippo pathway plays an important role in myogenic differentiation and muscle regeneration[16,53-56]. Among the Hippo signaling components, Tead transcription factors (Tead1-4) are key transcriptional regulators to coordinate genes essential for muscle cell fate determination, proliferation, and differentiation[18]. Silencing of Tead1 or Tead4 impaired C2C12 differentiation but not primary myoblast differentiation[16], suggesting the important role of Tead1 and Tead4 in differentiation of C2C12 cells. The Tead1 transcription factor is vital for developmental processes including cardiac development, cardiac hypertrophy, skeletal muscle formation, myoblast differentiation, and skeletal muscle regeneration[57]. It regulates the expression of key muscle-specific genes such as cardiac troponin T (Tnnt2), β-myosin heavy chain (Myh7), smooth muscle α-actin (Acta2), and skeletal α-actin (Acta1)[57]. In C2C12 cells, Tead1 was upregulated during myogenic differentiation but Tead1 knockdown inhibited cell differentiation[57-60]. Similarly, our results showed an increase of Tead1 with differentiation in untreated C2C12 cells, but a significant downregulation at each timepoint with Cr(VI) treatment (Fig. 6). Downregulation of Tead1’s transcriptional activity further supported by the Cr(VI)-induced downregulation of muscle-specific target genes Tnnt2, Acta1, and Acta2 in our results (Fig. 4). TEAD4 is specifically expressed in developing skeletal muscle of mouse embryos[61] and is strongly upregulated during C2C12 differentiation[18]. It has been reported that Tead4 binds to a conserved MCAT motif within the MYOG promoter, and its knockdown or disruption of its DNA-binding domain has been shown to significantly reduce MyoG activation[18]. Our study showed that Tead4 mRNA levels were increased with C2C12 differentiation but almost completely abolished by Cr(VI) treatment at all differentiation timepoints (Fig. 6). Consistently, many Tead4 downstream targets, such as MyoG, Myh1, Myh2, Myh3, Myh7, Myh8, and Cav3, were significantly down-regulated in Cr(VI)-treated cells. Collectively, our findings suggest that Tead1 and Tead4 are important mediators of Cr(VI)’s inhibitory effect on C2C12 myogenic differentiation, likely through its regulatory role in musclespecific genes and cell cycle genes.

In summary, this study provides the first comprehensive transcriptomic analysis of Cr(VI)-exposed C2C12 cells during myogenic differentiation. Our findings demonstrate that Cr(VI) exposure impairs key transcriptional programs responsible for sarcomere assembly and muscle contraction while disrupting essential pathways governing cell cycle withdrawal, DNA damage repair, and Hippo signaling. These alterations correlate with the phenotypic deficits described in our previous study. Together, these data provide new mechanistic insight into how environmental toxicants like Cr(VI) impair skeletal muscle development and regeneration. Future studies are warranted to explore whether interventions targeting these disrupted pathways may restore proper myogenesis.