Cardiomyocyte hypertrophy co-localizes with accumulation of hyaluronan in cMyBPC+/- mice hearts

Anna Strnadová; Madelene Ericsson; Rakel Nyrén; Ilona Dudka; Stellan Mörner; Urban Hellman

doi:10.1371/journal.pone.0342635

Abstract

Hypertrophic cardiomyopathy (HCM) is a genetic heart disease characterized by left ventricular hypertrophy. Mutations in genes for sarcomere proteins, e.g., in cardiac myosin-binding protein C (cMyBPC), may cause symptoms like heart failure and sudden cardiac death. In HCM, the heart utilizes high amounts of glucose. Increased glucose metabolism creates intermediates for hyaluronan (HA) synthesis, resulting in HA accumulation within the extracellular space, which could contribute to cardiomyocyte hypertrophy. We aimed to describe the connection between HA and cardiomyocyte hypertrophy using cMyBPC^+/- mice treated with two different β-blockers – either metoprolol or propranolol – for 11 months. The results showed that HA metabolism is altered in HCM, and that HA is significantly more abundant in hearts of cMyBPC^+/- mice compared to wild-type mice. Areas of high HA abundance were spread around the tissue in a ‘patchy’ manner. This focally increased HA concentration correlated with increased cardiomyocyte size in the same areas of the heart. Gene expression of HA synthase 2 was elevated in cMyBPC^+/- mice. Treatment with propranolol maintained HA synthase 2 at wild-type level, while significantly increasing HA synthase 3 expression. However, treatment with metoprolol or propranolol did not prevent HA accumulation nor decreased cardiomyocyte hypertrophy. These novel findings visualise a spatial connection between high HA abundance and cardiomyocyte hypertrophy in the hearts of cMyBPC^+/- mice.

Citation: Strnadová A, Ericsson M, Nyrén R, Dudka I, Mörner S, Hellman U (2026) Cardiomyocyte hypertrophy co-localizes with accumulation of hyaluronan in cMyBPC^+/- mice hearts. PLoS One 21(9): e0342635. https://doi.org/10.1371/journal.pone.0342635

Editor: Andre van Wijnen, University of Vermont College of Medicine, UNITED STATES OF AMERICA

Received: January 26, 2026; Accepted: August 24, 2026; Published: September 9, 2026

Copyright: © 2026 Strnadová et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the manuscript and its Supporting Information files.

Funding: Region Västerbotten (Bas ALF - RV-1035313 Centrala ALF - RV-1010683) Norrlandska hjärtfonden.

Competing interests: The authors have declared that no competing interests exist.

Abbreviations: HCM, hypertrophic cardiomyopathy; cMyBPC, cardiac myosin binding protein C; HA, hyaluronan; WT, wild type; met, metoprolol; prop, propranolol; PET, positron emission tomography; FDG, fluorodeoxyglucose; ROI, region of interest; c[HA], concentration of hyaluronan; HAS1, hyaluronan synthase 1; HAS2, hyaluronan synthase 2; HAS3, hyaluronan synthase 3; UDP-GlcA, UDP-glucuronic acid; UDP-GlcNAc, UDP-N-acetylglucosamine; TMEM2, transmembrane protein 2; CEMIP, cell migration inducing hyaluronidase 1; GLUT4, glucose transporter 4

Introduction

Hypertrophic cardiomyopathy (HCM) is a genetic disease affecting the cardiac muscle, often caused by heterozygous mutations in genes encoding sarcomere proteins such as myosin heavy chain or cardiac myosin binding protein C (cMyBPC). Affecting primarily the left ventricle, this condition causes hypertrophy, predominantly of the interventricular septum. This may lead to hypercontractility, left ventricular outflow tract obstruction, diastolic dysfunction, and arrhythmias. The prevalence of HCM in the general population is 0.2%, with males being more commonly affected than females, although the inheritance pattern is autosomal dominant [1–3]. Individuals with this condition can begin to experience symptoms at any point in their lives, from newborns to older adults. HCM is a common cause of sudden cardiac death, which may occur as the first manifestation of the disease [4,5]. Treatment consists of either pharmaceutics designed to relieve symptoms, such as β-blockers, or invasive surgical procedures [6,7]. There are no therapies available to prevent the onset of HCM in humans [2,7]. However, in children with HCM, hypertrophy can be attenuated by high doses of β‑blockers, such as propranolol [8,9]. Recently, mavacamten, a cardiac-specific myosin inhibitor, was approved for use in obstructive HCM. In a mouse model expressing a human myosin mutation, early treatment with mavacamten attenuated the development of ventricular hypertrophy [10].

On a microscopic level, HCM hearts are described to exhibit hypertrophied cardiomyocytes, cellular disarray and fibrosis in varying degrees. A previous study [11] reported that hearts from HCM patients showed high variability – not only between individuals, but also within sections from the same heart. Varnava et. al have demonstrated that severely re-modelled and hypertrophied cardiomyocytes neighbour normally sized and aligned cardiomyocytes, creating focal patches of pathological and healthy tissue throughout the heart.

Recent research suggests that myocardial energetic pathways are impaired in HCM; sarcomere gene mutations lead to inefficient ATP utilization and mitochondrial dysfunction [12]. It has previously been shown that the expression of genes connected to energetics is decreased in HCM [13]. This results in unmet high energy demands in the heart, causing energetic stress and adverse remodelling [13,14]. HCM forces the heart to re-activate the so-called fetal gene program, where adult cardiomyocytes revert to fetal-like gene expression to endure stress [15,16]. The heart withdraws from using fatty oxidation as its main energy source and starts depending on glucose metabolism to keep up with the elevated energy demands [17,18]. During stress, the proportion of ATP derived from glucose oxidation is increased more than two‑fold compared to normal conditions [19]. In an acute setting, this adaptation is beneficial, however, the sustained pressure overload present in HCM makes this metabolic change permanent. Lasting reliance on glucose metabolism is associated with impaired metabolic flexibility and mitochondrial dysfunction, which may be responsible for reduction in cardiac efficiency [20,21]. In human HCM cardiac tissue, we observed differential expression of immediate early genes which are known to regulate both the re‑expression of fetal genes and synthesis of hyaluronan (HA), a glycosaminoglycan present in the extracellular matrix of the heart [22]. HA is a non‑sulphated glycosaminoglycan consisting of repeating disaccharide units of glucuronic acid (GlcA) and N-acetyl-glucosamine (GlcNAc). It is one of the most abundant components of the extracellular matrix, where it plays a crucial role in tissue hydration, cell survival and cell migration [23].

The exact involvement of glucose metabolism in HCM pathogenesis is not yet properly understood. Nonetheless, an example of high blood glucose-caused cardiac hypertrophy can be seen in infants born to diabetic mothers, where the children suffer from severe cardiac hypertrophy at birth; however, this condition resolves spontaneously as normoglycemia is established over the following months [24].

In a previous study from our lab, we described a novel link between elevated glucose usage and the development of cardiac hypertrophy; our results also suggested an increased risk of arrhythmia and sudden cardiac death in HCM. The connection between glucose metabolism and cardiac hypertrophy was detected in cardiac tissue from human HCM patients, in cultured rat cardiomyocytes, and in rats with induced cardiac hypertrophy [22,25–28]. We observed that increased glucose metabolism leads to increased synthesis of HA [22,25,26,28].

Specifically, analysis of metabolites in cardiac tissue of HCM patients [22] has shown a decrease in fatty acid levels, while lactate levels increased, demonstrating that the HCM heart relies on the less efficient glucose metabolism. UDP-sugars, such as GlcA and GlcNAc, the two precursors for HA synthesis, were also substantially increased [22,28]. It is well-known that elevated levels of HA precursors lead to increase of HA synthesis [29,30]. We found larger amounts of HA, especially low molecular weight fragmented HA, in cardiac tissue of HCM patients than in healthy controls [28]. Similarly, in aorta-ligated rats with cardiac hypertrophy, HA concentration was increased compared to sham-operated controls [25]. Excessive deposition of HA in the heart might be associated with hypertrophic cardiomyocyte remodelling. HA of different molecular weights can stimulate opposite biological effects – overabundant HA in form of fragmented low molecular weight molecules around individual cardiomyocytes could contribute to inflammation [31]. On the contrary, high molecular weight HA has been described to be anti-inflammatory [32]. A way to affect glucose metabolism, and possibly by extension HA metabolism and hypertrophy, is by administering β-blockers; this commonly used therapy for cardiac dysfunction is known to decrease cellular glucose uptake, insulin sensitivity and hepatic glucose production [33,34].

Here, we focused on exploring the connection between HA, glucose and hypertrophy in depth using cMyBPC^-/+ mice, which were described to develop left ventricular hypertrophy akin to human HCM at 11–12 months of age [35]. To investigate the effect of β-blocker treatment on HA and cardiac hypertrophy, the mice were treated from weaning until 1 year of age with either high‑dose propranolol, commonly used for treating children with HCM [8,36], or the standard dose of metoprolol, predominantly used in the clinics to treat adults with HCM [2,37].

Materials and methods

Animal model

Male mice heterozygotic for cardiac myosin binding protein-C (cMyBPC^+/-) [35] and male wildtype (WT) C57BI/6J mice were included in the study. The animals were generated in‑house by breeding heterozygous males with WT female mice, to ensure healthy gestation and maternity. The pups were genotyped and assigned to groups according to their genotype. The first study with metoprolol was performed three years prior to the propranolol study. An overview of included mice and treatments is summarized in Fig 1.

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Fig 1. Overview of mice included in this study.

Male wildtype (WT) C57BI/6J mice and male heterozygous cardiac myosin binding protein-C (cMyBPC^+/-) mice, given either pure drinking water, or β-blocker dissolved in their drinking water; metoprolol (30 mg/kg/day) in Met-group (cMyBPC^+/- + met mice) or propranolol (37 mg/kg/day) in Prop-group (cMyBPC^+/- + prop mice). Reprinted from BioRender under a CC BY license, with permission from BioRender, original copyright 2026.

https://doi.org/10.1371/journal.pone.0342635.g001

The mice were housed in groups of 2–4 mice/cage under standard housing conditions (12:12h light-dark cycle, 22°C temperature and 45–50% humidity), with free access to standard rodent chow and drinking water. Treatment started after weaning. cMyBPC^+/- mice were given either pure drinking water or a β-blocker dissolved in their drinking water (cMyBPC^+/- + met mice given metoprolol (30 mg/kg/day) in Met‑group or cMyBPC^+/- + prop mice given propranolol (37 mg/kg/day) in Prop-group) (Fig 1). Blood glucose was measured at endpoint from tail vein blood using a hand-held ACCU-Chek AVIVA device (Roche Diagnostics, Solna, Sweden). When twelve months old, the mice were sacrificed via cervical dislocation. Hearts were excised immediately after euthanasia, dissected into specific regions (left ventricle, right ventricle, apex and atria), snap-frozen in liquid nitrogen and stored at −80°C until use.

The mice were cared for and used in accordance with Swedish Animal Welfare Act and approved by the Animal Review Board at the Court of Appeal for Northern Norrland in Umeå, Sweden (A62/14, approved on 2014-08-26).

Echocardiography

Cardiac function was assessed using the Vevo 2100 (VisualSonics, Fujifilm, Toronto, Canada), and the MS500D transducer. The examination was performed during light isoflurane anaesthesia (1.0–2.0% in 800 mL/min O₂) (Baxter Medical, Kista, Sweden). The level of anaesthesia was adjusted to keep the respiration rate at 90–110 breaths/min [38]. Image acquisition was previously described [38,39]. Image analysis was performed off-line in a blinded manner by AS, using the Vevo LAB 5.10.0 software (VisualSonics, Fujifilm). The mean of three strokes was used for each parameter and mouse.

Positron Emission Tomography (PET) for glucose uptake

To evaluate cardiac glucose uptake in the Prop-group, the radioactive glucose analog [18F]‑Fluorodeoxyglucose (FDG) was administered. In short, during light isoflurane anesthesia, as described under echocardiography, mice were injected with 70–100 µL (12.8 ± 4.8 MBq) of clinical grade FDG in saline, prepared at the Nuclear Department at Norrlands University Hospital, Umeå, Sweden. A tailor made 27G needle and catheter was used to ensure correct administration in the tail vein. After injection, mice were allowed to stay awake and freely moving in their cage. After 120 minutes, mice were again sedated with isoflurane and CT was performed for gross morphological orientation and then scanned for a 10-minute static PET acquisition (nanoScan PET/CT, Mediso, Hungary). Images were reconstructed over a 98 mm axial field‑of‑view to a 0.4 x 0.4 mm slice resolution, employing a 3D iterative reconstruction, four iterations and four subsets (Mediso Tera-Tomo 3D). The reconstruction was performed with delayed-window random correction, spike filter, scatter, and CT-based attenuation corrections. Tracer uptake in myocardial tissue was evaluated by using imlook4d (https://github.com/JanAxelsson/imlook4d), and quantified as standardized uptake values (SUV), using the formula: SUV = C / (i / m); with C being the measured tissue activity concentration (Bq/mL) in the image, i the injected dose, and m the mouse body weight. After PET/CT scan, mice were allowed to wake up and return to their home-cages. FDG-PET was only performed for Prop-group, as it was not available when Met-treatment was performed

RT-qPCR

Collected tissues were snap frozen in liquid nitrogen and stored at −80°C. The apex of the heart was used for gene expression analyses. Tissues were thawed in RNAlater^TM-ICE (Invitrogen, ThermoFischer; AM7030, Waltham, USA) according to manufacturer’s protocol. Total RNA was extracted using RNeasy Mini Fibrous Kit (Qiagen; 74704; Hilden, Germany). All samples were treated with RNase-Free DNase (Qiagen; 79254). RNA concentration and purity were measured on a NanoDrop. cDNA was prepared with RevertAid H minus Reverse Transcriptase (Thermo Scientific) and Random Hexamer primer (Thermo Scientific) in a Thermocycler from Biometra.

MasterMix TaqMan Fast Advanced (Applied Biosystems, ThermoFischer; 4444557), housekeeping gene (GAPDH-VIC or Rpl32-VIC) and one of the following genes (Table 1) were pipetted onto the MicroAmp Optical 96-Well plates (Applied Biosystems, ThermoFischer; 4306737) and cDNA of each sample was added to a final concentration of 40 ng/well. All samples were run in triplicates. For negative control, cDNA was replaced by nuclease-free water (Fisher Bioreagens; BP2484−50). ROX was used as a passive reference.

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Table 1. Gene probes used for RT-qPCR.

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Gene expression was quantified using QuantStudio 6 Flex System instrument. The amplification was run in comparative mode and analysed in QuantStudio 6 software (v1.7.2). Relative quantification was calculated according to the 2^-ΔΔCT method using either GAPDH or Rpl32 as a housekeeping gene and WT mouse sample as the control group.

Histopathological evaluation and scoring of cardiac fibrosis

Histopathological assessments were performed on 4 µM formalin fixed and paraffin embedded sections from the left ventricle of each mouse [40]. Screening for histopathological changes such as disarray was done using standard hematoxylin and eosin staining. Collagen fibres were visualized using Masson’s trichrome stain (HT15, Sigma-Aldrich, St Louis, MO, USA) according to the manufacturer’s protocol. The presence of interstitial fibrosis was evaluated in a blinded manner by RN in twenty randomly selected high-power fields (0.24 mm²) per heart using a Nikon Eclipse Ci light microscope. Each area was assessed using a semi-quantitative scoring system (0 = no fibrosis, 1 = slight, 2 = moderate, 3 = severe) and presented as mean value per individual [41].

Hyaluronan quantification and cell size analysis

For hyaluronan staining, cardiac sections were pretreated with 3% H₂O₂ in methanol and 1% bovine serum albumin solution (10 mg/mL in PBS) before overnight incubation at 4°C with biotinylated hyaluronic acid binding protein probe (100 ug/mL in PBS, 1:40) (HABP, Sigma-Aldrich, St. Louis, USA).

The HABP probe was visualised using VECTASTAIN® Elite® ABC-HRP Kit (Vector Laboratories, Newark, USA) and DAB substrate kit (Vector Laboratories) according to manufacturer’s instructions. Cell nuclei were stained using Mayers HTX PLUS (Histolab Products AB, Askim, Sweden). The samples were imaged using Pannoramic 250 Flash III digital scanner (3DHistech, Budapest, Hungary).

Using QuPath software (version 0.5.1), each sample was annotated as a region of interest (ROI) using the pixel qualification function. Subsequently, every sample was divided into square tiles (200 µM a side) that were trimmed to ROI borders. This tile approach was implemented to mitigate the individual differences among samples.

To visualise hyaluronan staining, pixel qualification was performed (settings: resolution 0.24 µM/px, DAB channel, sigma 0.5, threshold 0.1). Next, in each sample, we identified tiles that contained no tissue edges, no excessive tearing and no vessels. Tiles containing vessels were excluded as vessels naturally contain high amounts of hyaluronan. This could distort the results as there is variation between the samples in how many vessels they contain. Furthermore, tiles with tearing or edges were also disqualified, as they contain a lot of empty space which would be incorrectly quantified as ‘no staining’ due to the software setup, thereby introducing errors into the analysis.

From tiles that fulfilled all criteria, 16 tiles containing high amounts of hyaluronan staining (high c[HA] areas; HA staining > 5%) and 16 tiled that contained little-to-no hyaluronan staining (low c[HA] areas; HA staining < 5%) were selected. In each of these tiles, we quantified the percentage of positively stained tissue, which corresponds to the amount of hyaluronan. For each sample, we could obtain an average percentage of positive staining in high c[HA] areas and in low c[HA] areas.

For cell width quantification, we again used QuPath software. Using the same tiles as for HA quantification, the width of 50 cells per sample was measured, approximately 3 per selected tile. The cell width was measured manually using the ‘Line’ function in QuPath. For the cell to be considered appropriate for measurement, it had to meet the following selection criteria: 1) the cell had to be longitudinally cut, 2) the cell nucleus had to be clearly visible and symmetric, 3) the cell’s edges had to be clearly defined.

Statistical analysis

Data analyses pertaining to comparing different experimental groups, such as one-way ANOVA or t-tests, were performed using GraphPad Prism statistical analysis package (version 10.4.1 (627)).

Pearson correlation analyses were performed using SPSS statistical package (version 29.0.2.0, BMI).

Data are expressed as mean ± SD. Statistical significance was set at P < 0.05.

Results

Cardiac function and glucose uptake by FDG-PET

To evaluate cardiac function and pathological cardiac growth in cMyBPC^+/- mice, as well as possible effects of two different kinds of long-term β-blockade treatment, high-frequency echocardiography was performed. In the first experiment, cMyBPC^+/- mice were treated with metoprolol from weaning to 12 months of age (Met-group). The same experiment was repeated

three years later, but a non-selective β-blocker propranolol was used instead (Prop‑group) (Fig 1). As HCM results in cardiac hypertrophy, left ventricular dimensions were assessed at 12 months. However, neither anterior nor posterior wall hypertrophy was found in cMyBPC^+/- mice in either of the two studies (Table 2). Nevertheless, in Met-group, cMyBPC^+/- mice had significantly reduced stroke volume and cardiac output. cMyBPC^+/- mice in the Prop‑group exhibited similar measurements as WT mice. Overall, none of the β-blockers appeared to influence any of the parameters measured by echocardiography.

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Table 2. Echocardiographic data summary at 12 months of age.

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In both experiments heart weight/body weight ratio was similar between WT and cMyBPC^+/- mice with no significant change after β-blocker treatment (Table 3). Blood glucose, measured at 12 months of age, did not differ regardless of genotype or treatment.

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Table 3. Gravimetrical data summary at 12 months of age.

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To evaluate if cardiac tissue had an increased uptake of glucose in vivo, FDG-PET was used. Glucose uptake was only measured in Prop-group, as the technique was not available at the endpoint of Met-group. The cMyBPC^+/- and cMyBPC^+/- + prop mice showed a non-significant increase in glucose uptake compared to the WT mice. Propranolol treatment did not alter glucose uptake. Interestingly, cMyBPC^+/- + prop mice showed two groups of data points – one group with low uptake and one with high uptake, with no data at the mean (Fig 2).

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Fig 2. Maximal cardiac glucose uptake by FDG-PET presented as mean SUV ± SD.

WT, wild type; cMyBPC^+/- + prop, cMyBPC^+/- mice treated with propranolol. SUV = standard uptake value.

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Histology

General histopathological assessment of cardiac sections stained with standard haematoxylin and eosin did not reveal any areas of disarray in the cMyBPC^+/- mice, one of the histopathological hallmarks of HCM. However, other signs of hypertrophy were found, such as enlarged, irregular and box-shaped cardiomyocyte nuclei. In addition, in some areas of cMyBPC^+/- mice hearts, the cardiomyocytes were focally enlarged (Fig 3, upper panel).

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Fig 3. Representative images of cardiac sections from WT, wild type; cMyBPC^+/- + met, cMyBPC^+/- mice treated with metoprolol.

cMyBPC^+/- + prop, cMyBPC^+/- mice treated with propranolol, stained with hematoxylin and eosin (A) (scale bar = 0.1 mm) or Masson’s trichrome (B) (scale bar = 0.05 mm). (C) Graphs show interstitial fibrosis (0 = no fibrosis, 1 = slight, 2 = moderate, 3 = severe) presented as a mean value per individual.

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Another histopathological hallmark of HCM is interstitial fibrosis, where collagen fibres expand and replace the cardiomyocytes. Using Masson’s trichrome staining, the abundance of fibrous tissue was scored using a semi-quantitative analysis. In Met-group, fibrosis seemed to be more abundant in cMyBPC^+/- + met mice. The staining showed a small amount of interstitial collagen in samples from cMyBPC^+/- mice in Prop-group, often extending from vessels (Fig 3, lower panel). While not significant, the amount of fibrosis seemed to be slightly higher in cMyBPC^+/- + prop group than in WT or cMyBPC^+/- mice. No large fibrotic areas were found in neither of the groups.

Elevated expression levels of HA synthases in cMyBPC^+/- mice

Hyaluronan is synthesized at the cell membrane by hyaluronan synthases (HAS1, HAS2, HAS3). In Met-group, no changes in expression of HA synthases were detected. In Prop-group, the expression of Has1 was unaltered in both treated and untreated cMyBPC^+/- mice. However, the expression of Has2 was increased in cMyBPC^+/- mice, whereas in cMyBPC^+/- + prop mice, Has2 expression remained at WT levels. For Has3 this trend was reversed – cMyBPC^+/- + prop mice show significant increase in Has3 expression in comparison with WT mice. In cMyBPC^+/- mice, Has3 levels were unchanged (Table 4).

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Table 4. Gene expression by RT-qPCR.

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Effect of propranolol on cell-surface hyaluronidases

Since we observed changes in expression of genes involved in HA synthesis in Prop-group, we continued to investigate genes connected to HA degradation. The expression of the hyaluronidases Hyal1 and Hyal2 was on the same level in WT and cMyBPC^+/- mice in both Met‑group and Prop-group, regardless of treatment.

In Prop-group, the expression of Cemip was decreased in cMyBPC^+/- + prop mice in comparison with cMyBPC^+/- mice (approximately 57% decrease, P = 0.0549), reaching similar expression level as in WT mice (Table 4). Expression of these genes could not be measured in Met-group due to lack of material.

Influence of β-blocker treatment on metabolic pathways in the heart

Next, we investigated genes that could be affected by Mybpc3 mutations or β‑blocker treatment, namely genes connected to glycolysis, fatty acid oxidation and inflammation. However, not many striking differences were detected.

In cMyBPC^+/- mice in Met-group, we observed an increase in the glucose transporter 4 (Glut4, also known as Slc4a2) expression. In contrast, in cMyBPC^+/- + met mice, the expression of Glut4 was significantly lower compared to cMyBPC^+/- mice, approximately on the same level as WT (Table 4). However, no significant changes were detected in Glut4 expression in Prop-group.

Correlation between cardiomyocyte size correlated and hyaluronan abundance in hearts of cMyBPC^+/- mice

To assess the amount of HA in the samples, sections of cardiac tissue were stained using an HA probe and percentage of positive staining was quantified in 16 + 16 tiles in pre-defined high c[HA] areas and low c[HA] areas. Fig 4. shows representative images of HA staining in low and high c[HA] areas, as well as an example of tilling on a cMyBPC^+/- sample.

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Fig 4. Representative images of HA staining showing a high c[HA] area (A) and a low c[HA] area (B) (scale bar = 0.05 mm) on a left ventricle from a cMyBPC^+/- mouse.

(C) Example of tiling for quantification of HA staining on a cMyBPC^+/- + prop heart, and an example of high c[HA] area (up), and a low c[HA] area (down). The cutoff for an area to be classified as low c[HA] was 5% of HA positive staining. For each type of area, the percentage of positive staining was quantified for 16 tiles and then averaged for the whole sample.

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The results show that in high c[HA] areas, the amount of HA present is generally higher in cMyBPC^+/- mice compared to WT mice. In Prop-group, this difference is highly significant. In the Met-group the data trend in the same direction (Fig 5 left). The β‑blocker treatment did not significantly affect HA abundance, regardless of the β‑blocker used.

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Fig 5. Average positive HA staining in either high c[HA] areas or low c[HA] areas.

WT, wild type; cMyBPC^+/- + met, cMyBPC^+/- mice treated with metoprolol; cMyBPC^+/- + prop, cMyBPC^+/- mice treated with propranolol. n = 7-10 mice per group, 16 tiles/sample. Data displayed as mean ± SD, significance was measured using one-way ANOVA with Tukey’s post‑hoc test. ***P < 0.001, ****P < 0.0001.

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In low c[HA] areas, the HA levels were comparable across all groups – less than 5% in all samples in both Met-group and Prop-group (Fig 5, right). This supports our hypothesis that hearts of cMyBPC^+/- mice contain patches of pathological tissue that produce overabundance of HA, as well as areas with healthy non-hypertrophic cardiomyocytes.

Lastly, we focused on cardiomyocyte size in the same pre-selected areas. The cell width was measured manually on longitudinally cut cells. The average cell width in high c[HA] areas was significantly larger in cMyBPC^+/- mice in comparison to WT mice in both Met-group and Prop‑group (Fig 6, left). The β-blocker treatment did not seem to affect the cardiomyocyte hypertrophy, regardless of the β‑blocker used. In low c[HA] areas, cell width was alike in all groups, between 10–13 µM, which corresponds with the levels of HA staining in these regions, which were alike in all groups as well (Fig 6, right). It is worth noting that for the control group, the cell size was nearly identical in both high c[HA] and low c[HA] areas. The data shows that cMyBPC^+/- mice, regardless of β-blocker treatment, have hypertrophied cardiomyocytes exclusively in areas rich in HA. When cell width from both high[HA] and low[HA] areas is averaged, the cell width in cMyBPC^+/- mice, both treated and untreated, remains significantly larger than in their WT counterparts (11.55 µM, 11.55 µM vs 10.44 µM (P = 0.0063, P = 0.0087) respectively in Met-group, and 14.29 µM, 14.36 µM vs 13.02 µM (0.0177, P = 0.0064) respectively in Prop-group).

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Fig 6. Average cardiomyocyte width in either high c[HA] area or low c[HA] areas.

WT, wild type; cMyBPC^+/- + met, cMyBPC^+/- mice treated with metoprolol; cMyBPC^+/- + prop, cMyBPC^+/- mice treated with propranolol. n = 7-10 mice per group, n = 50 cells/sample. Data displayed as mean ± SD, significance was measured using one-way ANOVA with Tukey’s post‑hoc test. ***P < 0.001, ****P < 0.0001; ns, non-significant.

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Furthermore, cell width and HA abundance were positively correlated (Pearson corr., r = 0.489, P = 0.02) for all mice, demonstrating a connection between these two features. The increase of cell width in cMyBPC^+/- + prop mice was also positively correlated with the expression of Has3 (Pearson corr., r = 0.776, P = 0.04).

Discussion

In this study, we investigated the connection between HA, glucose uptake and cardiomyocyte size in cMyBPC^+/- mice, as well as the effect of two β-blockers, metoprolol and propranolol, on HA production and hypertrophy when administered from early age (Fig 7). Metoprolol and propranolol are two of the most frequently used β-blockers. Both drugs are classed as non‑vasodilating β‑blockers, with different targets – propranolol is a non‑selective β‑1,2 antagonist, while metoprolol selectively affects β‑1 receptors. Non‑vasodilating β‑blockers lower heart rate and contractility. Moreover, they have known detrimental effects on insulin sensitivity and glucose uptake and are associated with an increased risk of developing diabetes [33]. These effects on glucose usage originate from unopposed α-1‑adrenergic activity as well as decreased blood flow to the peripheries [33,42].

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Fig 7. Summary of the main hypothesis and aims in the present study.

In HCM, the heart experiences increased energy demands due to a mutation in cardiac sarcomere genes, such as cMyBPC haploinsufficiency. To fulfil these energy requirements, the heart starts primarily using glycolysis to make energy. Glycolytic intermediates can be used in side‑pathways that give rise to HA precursors, UDP-GlcA and UDP-GlcNAc, resulting in increased HA production. Excessive HA accumulation in the extracellular space can disrupt correct cardiac function and contribute to hypertrophy. In this study, we investigated the connection between HA metabolism, glycolysis and cardiomyocyte hypertrophy using β‑blockers known to disrupt glucose uptake. Reprinted from BioRender under a CC BY license, with permission from BioRender, original copyright 2026.

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We demonstrated that hypertrophied cardiomyocytes are solely located in areas with high HA abundance, while in areas with low amount of HA, the cardiomyocyte size remains normal. Indeed, cell width and HA abundance are positively correlated. In cMyBPC^+/- + prop mice, we found a connection between increase of HA concentration and the expression of Has3, showing that propranolol modulates HA synthesis, presumably via increasing Has3 expression, while normalizing the expression of Has2. However, cardiomyocyte size itself was unaffected by the β‑blocker treatment. Although cMyBPC^+/- mice were used for our research to better mimic human disease, complete knockout of the Mybpc3 gene might have been able to show a stronger hypertrophic phenotype and subsequently a stronger effect of the treatment [43].

In the present study, we developed a technique for quantifying the amount of HA in cardiac sections. Dividing each sample into tiles helped attenuate individual differences between samples. Furthermore, we identified areas with high and low c[HA] levels and measured the percentage of positive HA staining in each. Quantifying HA in random tiles around the whole tissue section would diminish the difference between the groups, as areas with very low staining and very high staining would mix. We found that in high c[HA] areas, the amount of HA is significantly elevated in both untreated and treated cMyBPC^+/- mice in comparison to WT. In low c[HA] areas, the amount of HA remained at similar level in all groups. These results support the hypothesis that HCM is a ’patchy’ disease – meaning that the HCM heart comprises of mostly healthy normally-sized cardiomyocytes, which are interspaced with patches of abnormal hypertrophied cells [11].

RT-qPCR assays were performed to identify genes in the HA metabolism that were either affected by the Mybpc3 mutation or by β-blocker treatment. RNA was extracted from whole tissue samples, meaning that hypertrophied cardiomyocytes were mixed with healthy cardiomyocytes and analysed in bulk. It is plausible that altered pathological expression of some genes is diluted by normal gene expression from healthy cells; meaning that transcriptional changes in the hypertrophic cardiomyocytes were masked by the more abundant healthy cardiomyocytes, and only severe changes in expression were detectable. This effect might be responsible for some of the differences that were detected in gene expression between the metoprolol and propranolol treated mice. To be able to distinguish between healthy and pathological tissue within the same heart, spatial transcriptomics would therefore be helpful in future experiments.

In Prop-group, the expression of Has2 was significantly upregulated in untreated cMyBPC^+/- mice, suggesting increased HA production. In contrast, cMyBPC^+/- + prop mice had a significantly increased expression of Has3, and expression of Has2 was maintained at WT level. In HCM, hearts rely primarily on glycolysis to produce energy; it is less oxygen demanding than creating ATP via β-oxidation, like healthy hearts would [17]. The increased amount of glucose is then utilized in glycolysis, creating high numbers of glycolytic intermediates like glucose-6‑phosphate or fructose-6‑phosphate [30]. Instead of continuing via glycolysis into the TCA cycle, these intermediates can turn into HA precursors, UDP‑glucuronic acid (UDP‑GlcA) and UDP‑N‑acetyl‑glucosamine (UDP‑GlcNAc). Specifically, glucose-6‑phosphate is converted into glucose‑1‑phosphate and UDP-glucose. UDP-glucose dehydrogenase then turns UDP-glucose into UDP-GlcA. On the other hand, UDP-GlcNAc is synthesized via the hexosamine pathway, starting with fructose‑6‑phosphate. Hexosamine pathway combines glutamine, acetyl-CoA and uridine‑5’‑triphosphate, resulting in the end product, UDP‑GlcNAc [44,45]

HA synthesis depends on the availability of these UDP-precursors, and, by extension, on the availability of glucose. Up to 5% of all glucose molecules entering glycolysis are shuttled into synthesis of HA precursors [23]. Interestingly, when glucose uptake was measured in cMyBPC^+/- + prop mice with FDG-PET, the data formed two distinct clusters; one with high uptake, one with low uptake, with no data at the mean. This result does not correlate with the data distribution in results of HA positive staining, cell size nor gene expression. It is possible that glucose uptake was influenced by propranolol treatment, however, sensitivity to the β-blocker’s effect on glucose could vary in each individual [46].

When there is little glucose present in the cell, the concentration of UDP-precursors will be very low as well, resulting in changes in HA production. The activity of the individual HAS isoforms depends primarily on the cellular pool of UDP‑GlcNAc and UDP‑GlcA; while HAS2’s and HAS1’s activity increases with concentration of UDP-sugars, HAS3 is capable of HA synthesis at high speed even with minimal substrates available [29].

The elevated expression of Has3 in cMyBPC^+/− mice treated with propranolol suggests that HAS3 may serve as a compensatory mechanism for the reduced Has2 levels or for lowered quantities of HA precursors. This indicates a critical functional redundancy within HA synthesis pathways. If Has2 expression or HAS2 enzymatic activity is decreased, HAS3 could assume a pivotal role in upholding HA production essential for cardiac function. The importance of this mechanism is underscored by previous findings showing that Has2 knockout models are non‑viable; absence of HAS2 leads to severe cardiac malformations and embryonic lethality [37]. Consequently, HAS3 may represent an evolutionary safeguard designed to ensure sufficient HA synthesis, even under conditions where HAS2 activity is compromised. However, HAS3 tends to produce slightly shorter HA molecules than HAS2 [47]; it is not known how this change influences the heart. These results indicate that both the Mybpc3 mutation and propranolol treatment can affect HA synthesis, possibly by influencing glycolysis or glucose levels.

Surprisingly, in Met-group, no changes in expression of Has genes were observed (Table 4). The discrepancies between propranolol and metoprolol treated animals were presumed, due to difference in dosage as well as receptors affected. Propranolol, that affects both β1 and β2 receptor, was administered in a high dose of 37 mg/kg/day, while metoprolol, a specific β1 inhibitor, had a dosage of 30 mg/kg/day. Unexpectedly, no changes in expression of HA metabolism genes were detected in the cMyBPC^+/- mice in Met-group, whereas in untreated cMyBPC^+/- mice in Prop-group, the expression of Has2 was elevated. Since the treatment was administered via drinking water over the course of approximately 8 months it is unlikely that these differences are due to drinking habits of the mice. Interestingly, it has been reported that rats given propranolol in drinking water have a lower water intake than control rats [48]. The discrepancies in Has2 expression could be due to individual differences between the mice in Met-group and Prop-group, due to heterogeneity – which is a key aspect of this disease, or due to a compensatory effect from the remaining intact Mybpc3 gene allele.

Nonetheless, in Prop-group, the increase in Has2 expression was connected with higher abundance of HA around cardiomyocytes. In areas with high[HA], cardiomyocytes were substantially larger than in low[HA] areas. There was no change in Has2 expression in Met-group. However, we still observed an increase in cell width in areas with high abundance of HA. This could imply that there might be several pathways affecting HA production. HA synthesis could be affected directly via elevated Has2 expression. There could also be another, more indirect, pathway, where HA production is affected via an increase of HAS2 activity, but not expression, due to a large spillover of glycolytic intermediates into HA synthesis. Changes in HA metabolism seem to be a part of the ECM remodelling occurring during the development of cardiac hypertrophy. The expansion of hydrated HA network in the ECM could provide the cardiomyocytes with space to grow. Our previous study has reported that synthesis of HA increased after fibroblasts were incubated in media previously incubated with cultured cardiomyocytes [26]. Cardiomyocytes likely secrete intercellular signalling factors that induce HA production in fibroblasts.

In a prior study from our lab, aorta-ligated rats, which develop pressure-originated cardiac hypertrophy, showed a six-fold increase in the expression of Has1 and Has2 at day 1 post operation. Over the course of 42 days, Has2 dropped to basal levels, while Has1 dropped from six- to three-fold increase above normal level [25]. On the contrary, in human myectomy samples, we detected a decrease in expression of HYAL2 and HAS2, and upregulation of HAS3; a similar pattern to what we observed in this study in cMyBPC^+/- + prop mice. This similarity between gene expression in mice in this study and human patients from the previous study could be explained by the fact that several patients were being treated with β-blockers [49]. To our knowledge, the exact mechanism in which β‑blockers affect cardiac muscle cells and HA synthesis is unknown; however, in asthmatic airway smooth muscle cells, β2 antagonists have been observed to reverse pathological changes in HA metabolism when used together with steroids [50].

Depolymerisation of HA begins at the cell membrane, where the large molecules are cleaved into shorter fragments and then internalized. Recently, new cell surface hyaluronidases have been described – TMEM2 and CEMIP. While their function in mice has not yet been completely elucidated, they both can degrade HA into shorter fragments, which can then contribute to inflammation [51–54]. Even though we saw no changes in expression of traditional hyaluronidases, Hyal2 and Hyal1, Cemip was influenced by propranolol treatment. In cMyBPC^+/- + prop mice, the expression of Cemip was around 57% lower than in cMyBPC^+/- mice (P = 0.0549). This trend of lowered expression of HA degrading enzymes could indicate that HA degradation pathway is affected by propranolol. In our previous research, we have shown that hearts of both HCM patients and rats with pressure‑derived cardiac hypertrophy contain high amounts of fragmented LMW HA [25,28]. If LMW HA fragments are involved in the hypertrophic process, changes of expression or activity of HA degrading enzymes, such as CEMIP, could be the source of fragmentation.

We have also investigated the expression levels of genes related to glycolysis. We found that not the Mybpc3 mutation itself, but treatment with metoprolol affects glucose uptake. The expression of Glut4 was significantly downregulated in cMyBPC^+/- + met mice. Despite being applied in a higher dose, propranolol showed no effect on Glut4 expression (Table 4). Latest research suggests that β2 receptor activity, which is blocked by propranolol, is required for GLUT4-mediated glucose uptake [55]. The decreased expression of Glut4 observed after metoprolol treatment might be the cause for previously reported increased plasma glucose after metoprolol usage in diabetic patients [56]. However, we did not observe any changes in blood glucose levels in cMyBPC^+/- mice, regardless of treatment.

Echocardiography was used to monitor cardiac function. Data about morphological pathology of heterozygous cMyBPC mice is conflicting [35,57–59]. While Carrier et al. reported significant asymmetric septal hypertrophy of heterozygous cMyBPC mice at around 45 weeks of age [35], we only observed reduced cardiac output in the Met-group after 50 weeks of age and did not detect any changes in the Prop‑group. Similarly, Harris et al. reported that only 32% of heterozygous cMyBPC mice in their study developed cardiac hypertrophy by 50 weeks of age. They also did not detect any histological abnormalities, such as fibrosis or myofibrillar disarray, at this age, which is consistent with our findings [57]. Heterozygous cMyBPC mice older than 125 weeks were shown to exhibit these pathological changes in around 50% of cases, suggesting that cardiac hypertrophy can have very late onset in this mouse model. However, similar histological findings were also observed in age-matched WT mice, probably due to age‑related decline of physical functions [58,60].

Caution should be taken when comparing different studies due to different instruments and anaesthetic regimes or lack thereof used when measuring echocardiographic data. In the present study, mice were under light isoflurane anaesthesia, and wall thickness was measured in M‑mode with the papillary muscle used as an anatomical hallmark. It is possible that hypertrophy would have been detected if more areas were chosen for measurements.

The heterogeneity of this mouse model is likely caused by different levels of the full-length cMyBPC protein being produced. Studies have shown that truncated cMyBPC proteins rising from the mutated allele are rapidly degraded via the ubiquitin-protease system. In mice, no truncated proteins have been found to be incorporated into the sarcomere; therefore the pathomechanism likely involves haploinsufficiency [61,62]. In transgenic mice expressing only 40% of the full‑length protein, the cardiac function is preserved, indicating an effective compensatory mechanism [63]. Although this mouse model often does not develop visible cardiac hypertrophy, at a cellular level we have detected patches with cardiomyocyte hypertrophy in all cMyBPC^+/- mice, both treated and untreated.

A limitation of the present study was that a detailed investigation of HA pathways, such as causal testing at the genetic level or assessment of cardiac protein levels of HAS and other metabolic proteins, could not be carried out due to limited availability of murine cardiac material, and must therefore be addressed in future investigations.

The present study describes a novel connection between HA abundance and cardiomyocyte hypertrophy in cMyBPC^+/- mice. We observed that HA metabolism is affected by MyBPC mutation as well as β‑blocker treatment. Cardiomyocyte size is significantly increased in areas with high HA concentration. This suggests that changes in glucose metabolism could contribute to hypertrophy by modulating extracellular matrix, including HA. Interestingly, we observed several differences between mice in Met-group and Prop-group. In Prop-group, we demonstrated an increase in Has2 expression and higher abundance of HA around cardiomyocytes. In areas with high[HA], cardiomyocytes were substantially larger than in low[HA] areas. There was no change in Has2 expression in Met-group. However, we still observed an increase in cell width in areas with high abundance of HA.

Altogether, the study describes an association between HA and the hypertrophic process. Further studies will be needed to explore the intricacies of HA pathways, as well as to elucidate the exact mechanisms that occur during the development of hypertrophy, e.g., using cultured cardiomyocytes or cardiac organoids, or by using metabolic radiotracers, i.e., fatty acids, for in vivo energetic characterization.

Supporting information

S1. Dataset.

Overview of raw data for all experiments.

https://doi.org/10.1371/journal.pone.0342635.s001

(XLSX)

Acknowledgments

We acknowledge the Small Animal Research Imaging Facility (SARIF) at Umeå University for providing assistance in using the PET/CT and ultrasound systems.

We acknowledge Kristina Eriksson for her laboratory assistance.

“Fig 1. Overview of mice included in this study “ and “Fig 7. Summary of the main hypothesis and aims in the present study.” were created in BioRender under user Hellman, U.. Figures can be accessed at https://BioRender.com/w4gbm0k and https://BioRender.com/bnuzyrp, respectively. Figures are licensed for use under CC BY 4.0.

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Figures

Abstract

Introduction

Materials and methods

Animal model

Echocardiography

Positron Emission Tomography (PET) for glucose uptake

RT-qPCR

Histopathological evaluation and scoring of cardiac fibrosis

Hyaluronan quantification and cell size analysis

Statistical analysis

Results

Cardiac function and glucose uptake by FDG-PET

Histology

Elevated expression levels of HA synthases in cMyBPC+/- mice

Effect of propranolol on cell-surface hyaluronidases

Influence of β-blocker treatment on metabolic pathways in the heart

Correlation between cardiomyocyte size correlated and hyaluronan abundance in hearts of cMyBPC+/- mice

Discussion

Supporting information

S1. Dataset.

Acknowledgments

References

Elevated expression levels of HA synthases in cMyBPC^+/- mice

Correlation between cardiomyocyte size correlated and hyaluronan abundance in hearts of cMyBPC^+/- mice