Zebrafish casr modulates cardiac structure and function

Abstract

Calcium is fundamental to biological processes, and the calcium-sensing receptor (CaSR) is pivotal for maintaining calcium homeostasis. Dysregulation of CaSR is implicated in cardiovascular diseases and cancers. However, its specific function in cardiac development remains incompletely characterized. To address this, we generated casr knockout zebrafish, analyzed cardiac physiology, and performed transcriptomics to elucidate its mechanism. We established three independent casr knockout lines on a Tuebingen (TU) background. Homozygous mutants exhibited reduced heart size, decreased heart rate, impaired myocardial contractility, diminished cardiac output, spinal curvature, and swim bladder inflation failure. Lethality began at 11 days post-fertilization (dpf). Transcriptomics revealed that casr loss disrupted the cardiac muscle contraction pathway, which was associated with swim bladder defects and mortality. Crucially, in a Tupfel long-fin (TL) background, casr^−/− mutants displayed normal swim bladder development, restored expression of cardiac contraction genes, and significantly improved survival. Our results demonstrate that casr is essential for cardiac function and swim bladder inflation in zebrafish, acting through the Wnt signaling and cardiac muscle contraction pathways. Furthermore, the TL genetic background rescues these developmental defects, highlighting a strong effect of genetic background on phenotypic outcomes.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-39063-9.

Introduction

Calcium (Ca²⁺) serves as an essential and universal intracellular messenger and is implicated in a multitude of cellular and biological functions¹, ranging from bone formation and neurotransmission to hormone secretion², muscle contraction³, gene expression⁴ and cell proliferation^3,5. Cells regulate extracellular calcium concentration through various pathways, including calcium channels and calcium-binding proteins. The calcium-sensing receptor (CaSR), a G protein-coupled receptor (GPCR), is activated by extracellular calcium ions. It is highly expressed in the parathyroid gland and kidney, where it plays a central role in regulating systemic calcium homeostasis^6–8.

The calcium-sensing receptor is a plasma membrane class C GPCR, first discovered by Brown and colleagues in 1993⁹ as the first membrane protein known to sense ions. The human CASR gene is located on chromosome 3 and is composed of eight exons¹⁰. It is expressed in key tissues regulating calcium homeostasis, such as parathyroid cells, thyrocalcitonin-secreting C cells¹¹, bone¹² and the intestine¹³, underscoring its central role in calcium-related processes. The CaSR protein primarily maintains systemic calcium homeostasis by regulating the biosynthesis and secretion of parathyroid hormone (PTH)¹⁴. Upon detecting a decrease in the extracellular calcium concentration, the calcium-sensing receptor (CaSR) stimulates the parathyroid to secrete PTH. Secreted PTH functions to reduce renal calcium excretion, augment intestinal calcium absorption, encourage bone resumption, and release skeletal calcium. On the other hand, elevated physiological calcium levels result in receptor activation, consequently suppressing the synthesis and secretion of parathyroid hormone¹⁵. In conclusion, CaSR is instrumental for maintaining both extracellular and intracellular calcium homeostasis; influencing cell secretion, proliferation, differentiation, chemotaxis and apoptosis¹⁶. These intracellular signaling pathways are mutually dependent and can interact with each other.

CaSR dysfunction is implicated in a range of disorders. For instance, loss-of-function or gain-of-function mutations directly cause calciotropic disorders such as familial hypocalciuric hypercalcemia and pseudo-Bartter’s syndrome^17,18. Furthermore, aberrant CaSR signaling has been linked to the pathogenesis of several non-calciotropic conditions, including cardiovascular diseases and certain cancers^19–21. Supporting its role in cardiac pathology, Lu et al.²², demonstrated using rat models that CaSR activity exacerbates heart failure progression. Additionally, epidemiological studies associate alterations in serum calcium levels with an increased risk of cardiovascular diseases, including vascular calcification²³, coronary artery disease and myocardial infarction²⁴, these alterations may also exacerbate known risk factors such as hypertension and dyslipidemia. Collectively, these lines of evidence suggest that CaSR may act as a modulator in cardiovascular pathophysiology by regulating local and systemic calcium ion concentrations.

Zebrafish exhibit significant physiological, pharmacological, and genetic similarities with humans, as they possess organs and systems that resemble those of mammals²⁵. Zebrafish exhibit rapid development, with organogenesis occurring just three days postfertilization^26,27. The swim bladder, an organ unique to fish, provides buoyancy and allows fish to maintain balance with minimal energy expenditure within a certain depth range in the water²⁸. Furthermore, the maturation of the swim bladder strongly correlates with cardiac function. Research indicates that cardiac impairment can result in the inability to inflate the bladder²⁹. Zebrafish are suitable for cardiovascular development research because they can obtain sufficient oxygen through passive diffusion before the seventh day of development, even when they exhibit severe cardiovascular defects. Additionally, the unique attributes of zebrafish, such as optical transparency, ex utero fertilization, and a high number of offspring, greatly facilitate forward genetic screening³⁰. These advantages render zebrafish an appealing vertebrate model organism for congenital heart disease and heart development, as they supplement human and other mammalian studies on this disease.

In the present study, we hypothesized that casr is essential for normal cardiac function in zebrafish. To test this hypothesis and investigate the deleterious effects of casr deletion in an in vivo model, we generated three independent casr knockout lines. Our findings revealed that at 5 days post fertilization (dpf), the swim bladder of casr^−/− zebrafish failed to inflate normally, and the homozygotes ultimately died between 11 and 14 dpf. Subsequent experiments indicated a decrease in heart rate and cardiac output in homozygotes. Notably, these swim bladder and heart function defects were rescued in homozygotes with TL background, suggesting a significant influence of genetic background on the development of casr mutant zebrafish embryos. Furthermore, we provided evidence that the observed abnormal cardiac physiological function in casr mutants is closely linked to cardiac muscle contraction pathway.

Materials and methods

Ethics

All experimental procedures were approved by the Animal Care and Use Committee of Hunan Normal University (Changsha, Hunan, P.R China, approval number 2021/196) and were carried out in accordance with relevant guidelines and regulations. All the methods are performed in accordance with ARRIVE guidelines.

Zebrafish husbandry and embryo collection

In this study, wild-type zebrafish Tuebingen (TU) and Tupfel long-fin (TL) strains were obtained from the China Zebrafish Resource Center (Wuhan, China). Tg (nkx2.5: ZsYellow) reporter strain was obtained from the School of Basic Medical Sciences at Wuhan University. All adult zebrafish were maintained under standard conditions. Embryos were collected from natural mating, incubated at 28.5 °C and staged according to standard protocols³¹.

CRISPR/Cas9-mediated gene knockout

casr mutant lines were generated with the CRISPR/Cas9 system following previous methods³². Briefly, the casr target site sequences were located in the second exon, and the two target sites are 117 bp apart from each other. The sgRNAs were synthesized in vitro with T7 RNA polymerase. A total of 400 pg of Cas9-mRNA and 50 pg of casr-gRNA were co-injected into zebrafish embryos at the one-cell stage. For genotyping, polymerase chain reaction (PCR) was carried out using the following primer sets: 5′- GGCTGTTGAAACTAGAGGAAA-3’ and 3′- GAGAAGCTGACCATGTTTTGT-5’.

Quantitative real-time PCR (RT–qPCR)

Total RNA was prepared from 50 embryos at 5 dpf using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. First-strand cDNA was generated using a reverse transcription kit (TaKaRa) with random primers. Quantitative real-time PCR (qPCR) was performed using a SYBR green kit (Takara, Dalian, China) on a QuantStudio 3 Real-Time PCR system (Thermo Fisher Scientific). Relative expression levels of the tested mRNAs were determined using β-actin as an internal reference and the comparative Ct (2^−△△Ct) method, with a p-value of < 0.05 considered statistically significant. The stability of the β-actin reference gene was confirmed by comparison with ef1α and no significant change in its expression was detected between experimental groups. The sequences of primers used in this study are listed in S2 Table.

Whole-mount in situ hybridization (WISH)

Zebrafish embryos at the desired stages were fixed in 4% paraformaldehyde (PFA) overnight before processing for WISH analysis as previously described³³. Digoxigenin-UTP-labeled antisense RNA probes for cmlc1 and cmlc2 were generated via an in vitro transcription method using T7 RNA polymerase (Thermo Fisher, Waltham, MA, USA).

RNA sequencing

A total of 6 cDNA libraries were prepared from three biological replicates of casr^−/− and wild type zebrafish at 5 dpf. RNA quality was confirmed by a Nano200 nucleic acid analyzer (allsheng, China), and the libraries were sequenced with an Illumina Nova seq6000. The qPCR validation was performed on the same three biological replicates, with each sample assayed in triplicate. The data presented are the mean values from these biological replicates.

Cardiac function analysis

All surviving zebrafish embryos were anesthetized with 0.03% tricaine (MS-222; Sigma-Aldrich, Germany) and mounted in 1% low-melting agarose with 0.03% tricaine. At the end of the study, zebrafish were euthanized by an overdose of tricaine. All cardiac function recordings were performed at the standard zebrafish incubation temperature of 28.5 °C to ensure consistent physiological heart rates. Movies of beating hearts from embryos at 52 h postfertilization (hpf) were recorded using an IX71-Olympus inverted microscope equipped with a Hamamatsu C9300 digital camera. Heart rate was counted for 15 s and then multiplied by 4 to calculate the heart rate in beats per minute (bpm). The long-diameter (a), short-diameter (b) and M-mode heart segments at diastole and systole were obtained by semi-automated optical analysis^34,35. All images and data were obtained from zebrafish ventricles. Each experiment was repeated at least three times to ensure reliability.

Ventricular fractional shortening (FS) was calculated from long or short end-diastolic (D_diastole) and end-systolic (D_systole) diameters, as:

End-systolic area (ESA) and end-diastolic area (EDA) were calculated as the area of an ellipse:

Fractional area change was calculated as:

End-systolic volume (ESV) and end-diastolic volume (EDV) were calculated as the volume of an ellipsoid of revolution:

Stroke volume, which represents the volume of blood being pumped within 1 pumping cycle, can also be calculated from known data by subtracting EDV with ESV:

Finally, with the obtained heart rate and stroke volume data, cardiac output, which represents the volume of blood pumped within a time frame, can be calculated by the following formula:

Bioinformatics analysis

Gene set enrichment analysis (GSEA)³⁶ was performed on the list of genes sorted by fold change in the experiment, and the enrichment of the up- or down-regulated gene sets in the KEGG pathway database was calculated. The gene sets with fewer than 15 genes or more than 500 genes were excluded, and the t-statistic mean of the genes was computed for each KEGG pathway using a permutation test with 1000 replications. The up-regulated pathways were defined by a normalized enrichment score (NES) > 0, and the down-regulated pathways were defined by an NES < 0³⁷. R v3.2.2 was used to perform the data preprocessing. The mapping process was performed through packages such as ClusterProfiler and Bioconductor. A chord diagram was drawn on Sangerbox³⁸.

Gene mapping cloning experiment

The genetic background of the zebrafish was distinguished based on simple sequence length polymorphisms (SSLP) on different chromosomes^39,40. First, primers for identifying 25 pairs of sequence tagging sites (STSs) on zebrafish chromosomes were found in the NCBI database, after which the wild-type (WT) zebrafish genome with a TU and TL background was used as a template for in vitro amplification. The sequences of primers used for LG13 (linkage group 13) in this study were as follows: Z6007-F, AGCCCTAAACAAACCGGC; and Z6007-R, GAGCATGATCTGCTCGTTAGG.

Statistical analysis

All quantitative data are expressed as the means ± SD. Prior to statistical comparisons, the normality of all data distributions was confirmed using the Shapiro-Wilk test. For comparisons between two groups, statistical significance was assessed using the two-tailed Student’s t-test. For comparisons among three groups, a one-way analysis of variance (ANOVA) was performed, followed by Tukey’s post-hoc test for pairwise comparisons. A p-value of less than 0.05 was considered statistically significant and was indicated with asterisks (*p < 0.05, **p < 0.01, ***p < 0.001) or with different lowercase letters. All statistical analyses were performed using GraphPad Prism version 9.3.

Results

Casr is evolutionary conserved from zebrafish to human

To assess the evolutionary conservation of zebrafish casr, we performed multiple sequence alignment of coding sequences and amino acid sequences of CASR orthologs from human, mouse and zebrafish. Comparative analysis revealed a high degree of amino acid sequence identity between zebrafish CASR and its human (71.92%) and mouse (71.76%) orthologs (Fig. 1A). Structural domain analysis demonstrated that the seven transmembrane helices characteristic of CASR proteins were conserved, although sequence homology within these domains varied (Fig. 1A and B). At the nucleotide level, the zebrafish casr coding sequence exhibited 70.51% identity with its human ortholog. To further evaluate genomic conservation, we performed synteny analysis across the three species (Fig. 1C). This analysis showed that the genes flanking CASR in human and mouse (B4GALT4, ARHGAP31) occupied syntenic positions relative to the genes flanking casr in zebrafish (b4galt4, arhgap31). These findings indicate that casr displays substantial evolutionary conservation, particularly within functionally critical domains, suggesting preservation of its molecular role across vertebrate lineages.

Fig. 1.

Evolutionary conservation of casr across species. (A) Multiple sequence alignment of CASR amino acids from human (1078 aa), mouse (1079 aa), and zebrafish (946 aa). Identical amino acids are shaded. The seven transmembrane domains (TMs) are marked with red boxes. (B) Sequence identity of the transmembrane domains among human, mouse, and zebrafish CASR proteins, using human CASR as the reference. (C) Synteny analysis of the genomic region surrounding the CASR locus. Gene names are shown, with numbers indicating chromosomal locations in megabase pairs (Mbp). Chromosome segments are represented with black lines. Orthologs are connected with blue lines.

Expression pattern of Casr during early development of zebrafish

To investigate the spatiotemporal expression pattern of casr during embryonic development, whole-mount in situ hybridization (WISH) was performed. casr transcripts were initially detected at the one-cell stage (Fig. 2A). From 14 to 24 h post-fertilization (hpf), expression was ubiquitous throughout the embryo (Fig. 2B and C). By 48 hpf, expression became localized to the brain (Fig. 2D). No discernible signal was detected at 96 hpf (Fig. 2E). At 120 hpf, casr exhibited tissue-specific expression in the swim bladder and otic vesicle (Fig. 2F). RT-qPCR confirmed that casr mRNA was highly expressed at the one-cell stage and gradually decreased thereafter, which was consistent with the WISH results (Fig. 2G). This dynamic expression pattern suggests a potential role for casr in the development of various tissues.

Fig. 2.

Expression pattern of casr during zebrafish development. (A–F) Spatiotemporal expression of casr analyzed by whole-mount in situ hybridization. Lateral views are shown for the one-cell stage (A), 14hpf (B) and 24hpf (C). (D) Lateral view of 48 hpf. The arrowheads indicate the head area. (E) Lateral view of 96 hpf. (F) Lateral view of 120 hpf. Open and closed arrowheads denote otic vesicle and swim bladder, respectively. Scale bars: 500 μm. (G) Relative expression levels of casr at different developmental stages, as determined by RT-qPCR with β-actin as the reference gene.

Loss of Casr leads to defective swim bladder inflation and spinal curvature

To investigate the function of casr, we generated casr mutant lines using CRISPR/Cas9⁴¹. Three independent lines, each derived from a different F0 founder (carrying 80 bp, 76 bp, or 83 bp deletions), were established. Each mutation disrupted the open reading frame, introducing premature stop codons predicted to cause severe loss of function (Fig. 3A and B). Sequencing confirmed these lesions (Fig S1A-D). RT–qPCR showed significantly reduced casr expression in mutant lines (Fig. 3C). The stability of the β-actin reference gene was validated by comparison with ef1α, showing no differential expression between wild-type and casr^−/− groups (Fig S2). Consequently, β-actin was used for normalization in all subsequent gene expression analyses.

Fig. 3.

casr deficiency results in developmental defects associated with downregulated Hh and Wnt signaling. (A) Schematic of the wild-type (WT) casr locus and the three mutant alleles generated by CRISPR/Cas9. (B) Schematic of the predicted wild-type and mutant CASR proteins (C) RT-qPCR analysis shows significantly reduced casr expression in the mutant line (line3) compared to the wild-type control. (D-G) Compared to control (D, F), casr^−/− larvae exhibited abnormal swim bladder development at 5 dpf (E), and spinal curvature occurred at 7 dpf (G). The red and blue arrowheads indicate the positions of the swim bladder and spine, respectively. (H, I) The relative expression of mRNAs associated with the Hh signaling pathway (H) and the Wnt signaling pathway (I) in the control and casr^−/−. (J, K) Line chart of GSEA results for the Hh signaling pathway (J) and Wnt signaling pathway (K). The ordinate represents the enrichment score (ES), and the ES at the highest peak is the ES value of the gene set. All the genes were tested with 3 replicates (N = 3), each containing 50 embryos (n = 50). The data are presented as the means ± SD. *p < 0.05; **p < 0.01; ***p < 0.001.

Embryos from heterozygous crosses were observed for developmental defects. While no obvious abnormalities were detected before 4 dpf, 24% (52/214) of larvae at 5 dpf exhibited failure of swim bladder inflation (Fig. 3D and E). These larvae rested at the bottom of the dish and showed impaired swimming. Genotyping of a subset (n = 15) of larvae with this defect confirmed they were all homozygous casr^−/−(Fig S2A). Conversely, larvae with normal swim bladders were either heterozygous or wild type (Fig S2B). By 7 dpf, larvae with uninflated swim bladders developed spinal curvature (Fig. 3F and G), a phenotype that worsened with time. Genotyping confirmed these larvae were also homozygous (Fig S2C). Notably, few homozygous mutants died at 11 days, and all the homozygous mutants died at 14 days. These results established the essential role of casr in zebrafish development. As all three mutant lines exhibited identical phenotypes, casr^line3 (casr^+/− and casr^−/− mutants) was selected for all subsequent experiments.

Given the established roles of Hedgehog (Hh) and Wnt signaling in endodermal organ and swim bladder development^42–47. To investigate whether, these pathways were altered in casr mutants, we performed qPCR analysis, which revealed downregulation of key Hh pathway genes (ihhb, shha, shhb, ptc1, ptc2) (Fig. 3H) and Wnt pathway genes (fzd1, fzd5, fzd7b, tcf3b, wnt1, wnt9b) (Fig. 3I) in casr^−/− embryos. Global transcriptomic analysis (RNA-seq) and GSEA further supported the coordinated downregulation of both the Hh (Fig. 3J) and Wnt (Fig. 3K) signaling pathways. Thus, casr deficiency is associated with the suppression of Hh and Wnt signaling, which correlates with the observed swim bladder defects.

Loss of Casr results in reduced ventricular size and cardiac output

To determine whether cardiac function is affected in casr^−/− mutants, we analyzed cardiac physiology at 52 hpf. We recorded high-speed movies of beating hearts and performed semi-automated optical analysis³⁵ to quantify cardiac structure and function. From these videos, we generated M-modes tracings to derive heart period (HP), diastolic diameter (DD) and systolic diameter (SD) (Fig. 4A and B). Representative still images at end-diastole illustrated ventricular morphology in control and mutant hearts (Fig. 4C and D), and a diagram showed the axes used for diameter measurements (Fig. 4E).

Fig. 4.

Analysis of cardiac function in casr mutants. (A, B) Representative M-mode traces from wild-type (A) and casr^−/− (B) ventricles. HP: heart period; DD: diastolic diameter; SD: systolic diameter. (C, D) Ventricular morphology at end-diastole in wild-type (C) and casr^−/− (D) larvae. (E) Schematic illustrating the axes (long and short diameter) used for ventricular measurements. (F-M) Quantification of cardiac parameters in wild-type and casr^−/− mutants: long diameter (F), short diameter (G), fractional shortening(H), fractional area change(I), heart rate (J), ventricular volume(K), stroke volume(L) and cardiac output (M). The data are presented as the means ± SD (n > 10). ^ns p > 0.05; *p < 0.05; **p < 0.01; ***p < 0.001.

Analysis of one-dimensional cardiac parameters revealed that both the long and short diameter were reduced in casr^−/− during diastole compared to controls, while no significant differences were observed during systole (Fig. 4F and G). To further evaluate myocardial contractility, we calculated fractional shortening (FS) and fractional area change (FAC). Both indices were reduced in casr^−/− (Fig. 4H and I), indicating substantially impaired systolic function. Heart rate was also decreased in the casr^−/− (Fig. 4J). Estimation of ventricular volumes revealed a significant reduction in end-diastolic volume, but not end-systolic volume, in casr^−/− (Fig. 4K). Consequently, both stroke volume (Fig. 4L) and cardiac output (Fig. 4M) were reduced in casr^−/−. Together, these data demonstrated that loss of casr resulted in globally impaired cardiac performance.

Casr is required for heart development in zebrafish

To investigate the transcriptional changes caused by casr knockout, we performed RNA-seq. This analysis identified 702 differentially expressed genes (DEGs) between control and casr^−/− zebrafish, with 337 upregulated and 365 downregulated (Fig. 5A). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis of the DEGs indicated a significant downregulation of the cardiac muscle contraction pathway (Fig. 5B). A heatmap of the DEGs was generated for visualization (Fig. 5C). Gene Ontology (GO) analysis was performed using the DAVID bioinformatics resource⁴⁸. The analysis results revealed that the DEGs were predominantly enriched in terms related to molecular function (Fig. 5D). Gene Set Enrichment Analysis (GSEA) further highlighted the coordinated downregulation of four key pathways: calcium signaling, cardiac muscle contraction, adrenergic signaling in cardiomyocytes, and regulation of actin cytoskeleton (Fig. 5E).

Fig. 5.

Transcriptomic analysis reveals dysregulation of calcium and cardiac muscle contraction pathways in casr mutants (A) Volcano plot of differentially expressed genes. Red and blue dots denote up- and down-regulated genes, respectively. (B) KEGG^49–51 pathway enrichment analysis of downregulated DEGs. Bubble size indicates the number of genes; color indicates the p-correction value. The y-axis shows the pathway name, and the x-axis displays the enrichment score. (C) Heatmap of DEGs. Rows represent genes, columns represent samples. Red and blue indicate higher and lower expression relative to the mean, respectively. (D) GO enrichment analysis. y-axis representing the number of genes and x-axis showing the name of the subclass. (E) A line chart of the GSEA results is shown, where the right ordinate represents the enrichment score. (F, G) RT-qPCR validation of key dysregulated pathways. Expression levels of calcium signaling pathway genes (F) and cardiac muscle contraction pathway genes (G) in wild type (TU) and casr^−/− zebrafish at 5 dpf. All the genes were tested with 3 replicates (N = 3), each containing 50 embryos (n = 50). The data are presented as the means ± SD. *p < 0.05; **p < 0.01; ***p < 0.001.

Based on the transcriptomic findings and the known role of CaSR in calcium homeostasis, we used RT‒qPCR to examine the expression of key genes in the calcium signaling pathway. The mRNA expression levels of ncx1b, pmca2, pth2, actc1, vdra, vdrb and runx2a were significantly lower in casr^−/− embryos compared to controls (p < 0.05), whereas pth1 expression was significantly elevated (Fig. 5F). To validate the RNA-seq results for the cardiac muscle contraction pathway, we selected six downregulated genes (fosab, tnnt2a, tnni2b.1, cnn1a, myhc4 and myh7l) for RT‒qPCR confirmation. The mRNA expression of these genes was downregulated in casr^−/− mutants (Fig. 5G), supporting the transcriptomic data. The functions of the abovementioned genes are listed in Supplementary Table S1.

Together, these data indicated that casr deficiency was associated with transcriptional dysregulation of genes involved in calcium signaling and cardiac muscle contraction.

The TL genetic background rescues phenotypic and functional defects in Casr mutants

To further explore the role of casr in heart development, we crossed casr^+/− fish with Tg (nkx2.5: ZsYellow) transgenic fish on a TL background. From these crosses, homozygous casr^−/− mutants were obtained that not only survived to adulthood but also exhibited normal swim bladder inflation (Fig. 6A and C), in stark contrast to the lethality and swim bladder defects observed in the TU background. All surviving homozygotes displayed the long-fin trait characteristic of the TL strain, regardless of the presence of the nkx2.5: ZsYellow fluorescence signal. (Fig S4A and S4B). Genetic background analysis using a polymorphic marker on linkage group 13 (LG13) confirmed that these viable homozygotes carried either a pure TL or a mixed TU/TL background, but never a pure TU background (S4C Fig).

Fig. 6.

The TL genetic background rescues morphological, functional, and molecular defects in casr mutants. (A-I) Diagrams of the phenotype (A-C), M-Mode (D-F) and ventricular morphology (G-I) of wild-type (TU), TU casr^−/− and TL casr^−/− zebrafish. The red arrowhead indicates the swim bladder. HP: heart period; DD: diastolic diameter; SD: systolic diameter. (J-M) Quantification of cardiac function. heart rate (J), FAC(K), stroke volume(L) and cardiac output (M) in the three groups. The plotted data are presented as the means ± SD (n > 10). (N, O) mRNA expression levels of genes from the calcium signaling pathway (N) and cardiac muscle contraction pathway (O). ^ns p > 0.05;*p < 0.05; **p < 0.01; ***p < 0.001.

We then assessed whether the TL background also ameliorated the cardiac functional defects. Cardiac imaging revealed improved ventricular dynamics in TL casr^−/− mutants compared to their TU casr^−/− counterparts (Fig. 6D and I; Movies S1-S3). Quantification showed that while heart rate remained depressed in both mutant groups (Fig. 6J), FAC and stroke volume in TL casr^−/− mutants were restored to levels not significantly different from TU wild-type controls, and were significantly higher than in TU casr^−/− mutants (Fig. 6K and L). Consequently, cardiac output in TL casr^−/− was significantly improved compared to TU casr^−/− and was comparable to wild-type levels (Fig. 6M). At the molecular level, RT-qPCR analysis of genes associated with the calcium signaling and cardiac muscle contraction pathways was performed on TU wild-type (TU), TU genetic casr^−/−, and TL genetic casr^−/− samples. As expected, the expression of these genes was downregulated in TU casr^−/ compared to TU. In TL casr^−/− mutants, while most genes in these pathways were still expressed at lower levels than in TU wild-type, their expression levels were generally intermediate: lower than in TU wild-type but significantly higher than in TU casr^−/−. Specifically, the expression of tnnt2a and cnn1a showed significant upregulation in the TL background compared to the TU casr^−/− mutants (Fig. 6N and O). These findings supported our hypothesis and demonstrated that the TL genetic background partially rescued the cardiac physiological defects caused by casr deficiency.

Casr deficiency is associated with altered expression of cardiomyocyte differentiation markers

To determine whether casr affects cardiomyocyte differentiation, we examined the expression of cardiac differentiation markers cmlc1 and cmlc2 by whole-mount in situ hybridization. WISH analysis revealed smaller hearts in both TU and TL casr^−/− mutants compared to TU wild-type controls (Fig. 7A and F). Ventricular short diameter was quantified from these images, confirming the size reduction (Fig. 7G). To further determine whether the mRNA expression levels of these two genes were changed, we performed RT‒qPCR. The results showed that the mRNA expression levels of cmlc1 and cmlc2 were significantly lower in the casr^−/− of TU background than in the WT (TU) (p < 0.05), and compared to casr^−/− of TU background, the mRNA expression level of cmlc2 was significantly higher in the casr^−/− of TL background (p < 0.05); however, cmlc1 expression was not significantly different (Fig. 7H). These findings indicated that casr deficiency was associated with reduced expression of cardiomyocyte differentiation markers (cmlc1, cmlc2) and a smaller ventricular size in the TU background. Further studies are needed to determine if this size reduction is due to changes in cardiomyocyte number, size, or both.

Fig. 7.

Reduced expression of cardiomyocyte differentiation markers in casr mutants correlates with smaller heart size and is partially rescued by the TL background. (A-F) Whole-mount in situ hybridization for cmlc1 (A-C) and cmlc2 (D-F) in the wild type of wild-type (TU), TU casr^−/− and TL casr^−/− at 48 hpf. n > 30. (G) Quantification of ventricular short diameter. (H) The mRNA expression of the cmlc1 and cmlc2 genes in the TU and casr^−/− lines with different genetic backgrounds. The plotted data are presented as the means ± SD. ^ns p > 0.05; **p < 0.01; ***p < 0.001.

Discussion

In this study, we elucidated crucial roles for zebrafish casr during early development. CRISPR/Cas9-generated casr mutants on a TU background exhibited severe morphogenetic defects, including swim bladder inflation failure and spinal curvature. This likely led to an inability to feed, resulting in 100% lethality by 14 dpf. The spinal curvature is consistent with the known role of CaSR in skeletal development. Chang et al.⁵² demonstrated that targeted deletion of Casr in mice results in profound skeletal defects, including reduced bone mineralization and abnormal skeletal morphology, primarily attributed to disrupted systemic calcium homeostasis. While the murine model reveals a direct role for CaSR in osteoblast and chondrocyte development, the spinal curvature in our zebrafish model could result from either disrupted calcium signaling in developing bone or be secondary to severe muscle weakness caused by impaired cardiac function. Nevertheless, its presence underscores the evolutionarily conserved function of CaSR in maintaining skeletal integrity. Furthermore, studies have reported a correlation between swim bladder inflation failure and cardiac dysfunction²⁹. Our analysis revealed that casr deficiency leads to global cardiac dysfunction. This is evidenced not only by a significant reduction in heart rate and cardiac output but also by impaired intrinsic myocardial contractility, as demonstrated by decreased fractional shortening (FS) and fractional area change (FAC). The concomitant reduction in ventricular diastolic dimensions and stroke volume further supports compromised pumping capacity. Mechanistically, this dysfunction may stem from disrupted calcium handling. It has been suggested that increased concentration of extracellular Ca²⁺ or other CaSR agonists might lead to increased sarcoplasmic reticulum Ca²⁺ release through the G protein-phospholipase C-inositol triphosphate signaling pathway, thereby triggering cardiac contraction. Conversely, a decrease in CaSR expression could result in a reduced Ca²⁺ concentration in the sarcoplasmic reticulum, impairing myocardial cell excitation–contraction coupling and subsequently leading to compromised myocardial contractile function we observed^53,54. These findings suggest that casr is essential for maintaining cardiac function, and its absence leads to a condition resembling heart failure.

The concomitant reduction in heart rate and stroke volume in casr^−/− mutants is physiologically intriguing. Under conditions of reduced cardiac output, compensatory tachycardia is a typical homeostatic response to maintain systemic perfusion^55,56. The absence of this compensation in our mutants suggests an impairment in the regulatory mechanisms governing heart rate. Several non-exclusive possibilities could account for this phenotype: (1) intrinsic dysfunction of the cardiac pacemaker or conduction system due to casr deficiency or associated calcium signaling dysregulation; (2) altered autonomic nervous system input to the heart during early development; (3) a broader systemic developmental delay or metabolic insufficiency; or (4) a primary defect in cardiomyocyte electrophysiology, given the central role of calcium. Future studies employing pharmacological challenges (e.g., with β-adrenergic agonists or antagonists) and direct electrophysiological recordings will be required to distinguish between these possibilities and to understand how casr influences the integrated regulation of heart rate and contractility.

We next explored potential molecular mechanisms. Wnt signaling is required for the morphogenesis of zebrafish swim bladder tissues, and its inhibition disrupts growth and differentiation⁵⁷. Hedgehog (Hh) signaling also plays a conserved role in the development of lungs and other endodermal organs^58,59. Given these established roles, we analyzed the impact of casr knockout on these pathways. Our results showed downregulation of genes in both the Wnt and Hh signaling pathways, suggesting that casr loss affected these pathways, which correlates with the swim bladder defect. Similarly, genes involved in the calcium signaling pathway were downregulated, while pth1 was upregulated, suggesting a compensatory effort to maintain calcium balance. The concurrent downregulation of Wnt and calcium signaling genes is intriguing, but whether they function in parallel or within an integrated pathway requires future study. Our analysis is limited to the RNA level; effects on protein expression and modification need investigation.

Transcriptomic analysis of casr mutants revealed significant downregulation of genes involved in the cardiac muscle contraction pathway (Fig S5), providing a molecular correlate for the observed contractility deficits (reduced FS and FAC). This link is particularly compelling when considering the distinct calcium handling mechanisms between zebrafish and mammals. In mammalian cardiomyocytes, contraction relies predominantly on sarcoplasmic reticulum (SR) calcium release, a process modulated by intracellular signaling. In contrast, the zebrafish heart depends more heavily on trans-sarcolemmal calcium influx via channels such as the L-type and T-type calcium channels for excitation-contraction coupling⁶⁰. This species difference is crucial for interpreting our findings. As an extracellular calcium-sensing receptor, CaSR is ideally positioned to modulate this primary source of activator calcium in zebrafish. We therefore propose a model wherein casr loss disrupts the regulation of calcium influx, leading to inadequate activation of the contractile machinery and resultant systolic dysfunction. The precise molecular intermediaries in this pathway await future discovery.

A striking finding was the complete rescue of lethality and swim bladder defects when the casr mutation was introduced into the Tupfel long-fin (TL) genetic background. Compared to TU casr^−/−, TL casr^−/− mutants survived to adulthood and showed significant improvement in cardiac function (e.g., FAC, stroke volume, cardiac output), alongside a partial restoration of gene expression in calcium and cardiac muscle contraction pathways (Fig. 8). This highlights a powerful modifying effect of genetic background on mutant phenotypes. Prior studies have indicated that diverse zebrafish strains possess unique morphological and behavioral traits, such as AB, Tubingen (TU), Wild India Kolkata (WIK), and Tupfel long fin (TL) strains. These traits, including swimming ability, are influenced by factors such as water temperature, caudal fin length, and genetic background⁶¹. Furthermore, variations in fin size, inherited traits, and physiological and behavioral characteristics in the juvenile and adult stages have been observed^61–63. Studies have shown that in other species, different backgrounds can affect the functions of genes. Kammenga⁶⁴ reviewed the phenomena of phenotypic variations caused by gene interactions, epigenetics and stochasticity in model organisms. In human, the polymorphism of the CYP2C19 gene alters the efficiency of drug metabolism, and there are differences in enzyme activity in different ancestral backgrounds^65,66. In addition, a strong background effect was found for mutation in polymerase-γ (POLG) in humans. Many patients carry a homozygous mutation but the disease phenotypes are highly variable, ranging from childhood-onset Alpers-Huttenlocher syndrome to neuropathy and deterioration of eye muscles⁶⁷. Vu et al.⁶⁸ knocked down ∼1,400 genes in the N2 Bristol and CB4856 Hawaiian isolates of Caenorhabditis elegans with RNAi and found that ∼20% of genes differ in the severity of phenotypes in these two genetic backgrounds. These evidences reveal that the genetic background is as important – if not more important – than the “disease-causing” mutation itself due to gene interactions, epigenetics and stochasticity. However, for this phenomenon, potential mechanisms and steps still need to be taken to provide more general insights. Furthermore, it is worth delving into the question of whether the TL background directly influences heart development or whether it alters the expression levels of related genes, ultimately affecting heart development.

Fig. 8.

Expression changes of cardiac muscle contraction pathway genes after casr gene knockout. The green shading and red indicate down-regulated and up-regulated expression in the casr^−/− with a TU background, while the orange shading and gray indicate up-regulated and insignificant differences expression in the casr^−/− with a TL background.

To explore the cellular basis of the smaller ventricle in casr mutants, we examined the expression of cardiomyocyte differentiation markers cmlc1 and cmlc2, the zebrafish homologs to essential and regulatory myosin light chains. Their expression was significantly downregulated in TU casr^−/− mutants. Intriguingly, the TL genetic background had a differential effect: it partially rescued the expression of cmlc2, but not cmlc1. Given that deletion of cmlc1 or cmlc2 in zebrafish affects cardiomyocyte size and number, leading to altered ventricular volume⁶⁹, this coordinated downregulation in our mutants raises the possibility that the casr deficiency phenotype may involve similar deficits in cardiomyocyte differentiation or growth. However, our study did not directly quantify cardiomyocyte size or number. Therefore, whether the smaller ventricle results primarily from a reduced number of cardiomyocytes, a reduction in cardiomyocyte size, or other structural alterations remains an open question, requiring future investigation with cellular-resolution imaging (e.g., membrane and nuclear staining). The regulatory relationship between casr and the cmlc genes also warrants further exploration.

In summary, our study demonstrates that casr is essential for cardiac function and swim bladder development in zebrafish. Loss of casr led to systolic dysfunction, which was associated with transcriptional downregulation of genes involved in calcium and cardiac muscle contraction. Notably, these defects were profoundly rescued by the TL genetic background, underscoring the powerful influence of genetic context on phenotypic outcomes. Future work will focus on identifying the specific genetic modifiers within the TL background that compensate for casr deficiency.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

LL: Conceptualization, Data curation, Methodology, Software, Validation, Visualization, Writing – original draft, Writing – review and editing. YH: Methodology, Software, Validation, Visualization, Writing – original draft. BX: Methodology, Software, Visualization. TZ: Methodology, Software. GT: Funding acquisition, Resources. HX: Investigation, Data curation. QS: Resources, Visualization. JL: Conceptualization, Writing – review and editing. HX: Conceptualization, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing – review and editing.

Funding

This work was supported by Hunan Normal University (NOs.053112 − 3826) and National Natural Science Funding, China (NOs.82170308).

Data availability

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. The RNA sequencing data from this study were deposited in the NCBI Sequence Read Archive under accession SRA: PRJNA967502([Danio rerio (ID 967502) - BioProject - NCBI](https:/www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA967502))

Declarations

Competing interests

The authors declare no competing interests.