Active and Passive Mechanical Deficits Precede Spinal Curvature in a Zebrafish Model of Idiopathic Scoliosis

  1. Johnathan R. O’Hara-Smith
  2. Samuel G. Bertrand
  3. Julissa Ortiz-Delatorre
  4. Rachael M. Giersch
  5. Luke A. Rethwill
  6. Damien M. Callahan
  7. Daniel T. Grimes

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Abstract

Idiopathic scoliosis is a common spinal disorder characterized by progressive three-dimensional curvature of unknown cause. Although biomechanical imbalance has long been proposed to contribute to scoliosis, the early physiological states that precede curvature onset remain poorly understood. Here, we investigated this problem using zebrafish uts2r3 mutants, which develop fully penetrant juvenile-onset spinal curvature following disruption of urotensin signaling. Transcriptomic analysis before curvature revealed altered expression of muscle-associated genes, suggesting that Uts2r3 influences axial muscle development or function. However, immunofluorescence, birefringence imaging, and quantitative analysis of myotome morphology showed that mutants lack overt muscle architectural defects or dystrophic pathology. By contrast, direct measurements of isolated larval trunks revealed pre-curvature biomechanical abnormalities: namely, uts2r3 mutants generated reduced active force following electrical stimulation while also exhibiting increased passive resistance to stretch. These findings identify urotensin signaling as a regulator of axial tissue biomechanics during growth and suggest that scoliosis-like curvature can arise from an early imbalance between active force generation and passive tissue stiffness.

Significance

Spinal curvature is common, but the biological events that cause the spine to bend during growth remain poorly understood. Animal models, especially zebrafish, make it possible to study these events before curvature begins. Zebrafish lacking urotensin signaling develop spinal curves that arise during juvenile growth, similar to idiopathic scoliosis in humans. Here, we demonstrate that zebrafish lacking the urotensin pathway receptor Uts2r3 develop an abnormal biomechanical state prior to curve onset. Their axial tissues generate less active force when contracting and, at the same time, show increased passive resistance to stretch—an unexpected combination that reveals a distinct pre-curvature biomechanical state. These findings suggest that spinal curvature can arise from an early imbalance in tissue mechanics during growth and identify urotensin signaling as a pathway that helps preserve spinal morphology through a biomechanical mechanism.

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    Note: This response was posted by the corresponding author to Review Commons. The content has not been altered except for formatting.

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    Reply to the reviewers

    Our central claim is that loss of Uts2r3 creates an abnormal mechanical state in the developing axis before overt spinal curvature, characterized by reduced active force measured from the axial preparation and increased passive resistance to stretch. These findings identify Urotensin signaling as a regulator of axial tissue mechanics during growth, and suggest that a breakdown of axial mechanics underlies spinal curvature phenotypes that are similar to idiopathic scoliosis. The cellular origin of this body-scale mechanical phenotype remain to be resolved and will be the subject of future work.

    Summary of the reviews and revision strategy

    __Overall assessment: __The reviews are highly positive about the importance of the question our manuscript addresses, the quality of the work, and our central observation that altered axial mechanicals precede spinal curvature. Reviewer 1 states that "The paper is well written and easy to follow" and, importantly, that "The conclusions are supported by the presented data." The same reviewer describes this as the first study showing that changes in muscle biomechanical function precede curvature in a model of idiopathic scoliosis. Reviewer 3 likewise states that "The data presented generally support the conclusions" and that the study "has the potential to advance the field". Reviewer 2 is more cautious, but also recognizes that the study addresses an interesting biological question and provides evidence that altered muscle function contributes to the spinal phenotype.

    Main issues raised: The reviews converge on a small number of issues: (1) the incomplete characterization of uts2r3 expression; (2) insufficient context and reporting of the RNA-seq dataset; (3) interpretation of the mechanical phenotype, especially whether reduced force reflects intrinsic contractile dysfunction, reduced contractile cross-sectional area, altered tissue composition, or altered force transmission; (4) the relationship between active and passive mechanical measurements in the intact tail preparation; and (5) methodological details, including the force-velocity analysis, sample numbers, statistical values, and RNA-seq deposition.

    Summary of our revision plan:

    1. uts2r3 expression and fiber type: We will revisit direct expression experiments across relevant stages and analyze existing public single-cell datasets. We will revise the manuscript so that prior evidence is described accurately as strongest for dorsal slow muscle rather than generalized pan-muscle expression. If direct localization remains inconclusive, we will state that limitation explicitly and be sure not to over-interpret negative data. These experiments are underway.
    2. RNA-seq context: We will add library, sequencing and quality control details, numbers of significantly upregulated and downregulated genes, fold changes for highlighted genes, a complete differential expression table, and a less redundant enrichment analysis. Our dataset has now been deposited public database.
    3. Contractile CSA and tissue composition: We will use transverse sections to quantify muscle area and connective tissue composition. We will incorporate MyHC/protein data if interpretable. Regardless of the experimental outcome, we will revise the manuscript to acknowledge the possibility of a 'less contractile tissue' explanation for the physiological phenotypes we observe. These histology and protein experiments are underway.
    4. Interpretation of active mechanics: We will soften the claims that the assay isolates intrinsic sarcomeric force generation, and reframe the measurements as integrated axial mechanics to distinguish force generation from force transmission. Additionally, we will remove the claim that active and passive abnormalities are necessarily independent (we did not mean to imply that). In our discussion, we will add molecular mechanisms through which these mechanical changes may manifest, including myotome shortening, notochord/myoseptal mechanics, and attachment site compliance. These changes to the writing of our manuscript will be performed prior to re-submission.
    5. Methods, statistics and figure reporting: We will expand the mechanics methods, clarify CSA measurements and mutant genotype, reconcile all n and p values, and add the requested kinetic data. The revision will make a clear and deliberate distinction between what the current experiments establish and what remains unresolved. The central supported conclusion is that loss of Uts2r3 produces a pre-curvature biomechanical state characterized by reduced active force measured from the axial preparation and increased passive resistance, accompanied by muscle-associated transcriptional changes and subtle changes in birefringence. The revised manuscript will not claim that these effects are proven to be muscle cell-autonomous.

    Detailed response to reviewer comments

    Reviewer #1

    Evidence, reproducibility and clarity

    The work by O'Hara-Smith et al. explores if alterations in muscle development and function underly the formation of spinal curvature in zebrafish uts2r3 mutants. Using transcriptomic analyses, immunofluorescence and birefringence, the authors reveal a reduction in the expression of muscle-associated genes as early as 28hpf with only mild reduction in birefringence and otherwise no overt changes in muscle morphology or symmetry, even at stages where spinal curvature becomes apparent (14dpf). Biomechanical analyses further show that loss of uts2r3 leads to a reduction in active force and an increased passive resistance to stretch, indicating that urotensin signaling plays a role in the regulation of muscle function.

    Response: We thank the reviewer for this clear summary of the study and for recognizing the combination of transcriptional, structural, and biomechanical evidence. We will preserve this central framing in the revision, while sharpening the distinction between the directly measured phenotype and the unresolved cellular mechanism.

    Major comment 1: The paper is well written and easy to follow. The majority of experiments are described in sufficient detail, and all of the used reagents are listed. The conclusions are supported by the presented data.

    Response: We appreciate this positive assessment.

    Major comment 2: Optional additional experiments that could significantly improve the paper would be a detailed characterization of the expression of uts2r3 as previous expression analyses were limited to one early time point and mostly used analyses of mosaic transgene expression. However, performing detailed expression analyses would be both costly and time consuming.

    Response: We agree that a clearer account of uts2r3 expression would improve interpretation of the mechanical phenotype. We are therefore revisiting expression using complementary approaches: in situ hybridizations, immunofluorescence using custom-made antibodies, and analysis of single-cell transcriptomic datasets from a range of pre-curvature stages. We will also revise the Introduction and Discussion so that prior expression evidence is described more precisely, including evidence for dorsal slow muscle expression and the current uncertainty about the expression in other muscle subtypes. If direct localization remains below reliable detection, we will minimally report single cell transcriptomic data and note the limitation explicitly in the manuscript.

    Minor comment 1: Additional details on the RNA-seq experiment should be added to the methods section to allow interpretation of the results. Was mRNA selection or ribosome depletion used? What was the sequencing mode (paired or single-end)? What read length was used? How many reads were obtained per sample and how many filtered reads could be aligned to the reference transcriptome?

    Response: We agree, and apologize for missing these details in our initial submission. We will be sure to include these details in our expanded Methods section.

    Minor comment 2: To understand the extend of the changes detected by the transcriptomic analyses it would be helpful to include the total number of significantly up- and down-regulated genes as well as the magnitude of change (fold-change) for the highlighted muscle related genes.

    Response: We will report the total numbers of significantly up- and down-regulated genes at the stated adjusted P value threshold and provide the fold changes of all genes available as a supplementary table. Additionally, our data has already been made available on a public repository, and we will provide the accession number in the revised manuscript.

    Minor comment 3: Similarly, for the GO terms a list of differentially expressed genes for each term should be provided and largely overlapping GO terms should be removed. The fact that all terms show almost the exact same fold enrichment could point to a large overlap between the differentially expressed genes detected for each term.

    Response: We appreciate this comment. We will re-analyze the enrichment, collapse highly redundant terms, and provide the gene membership underlying each term.

    Minor comment 4: As the ratio of twitch and tetanic force was quantified and used as a measure for myosin dynamics and force transmission, it would be helpful to show that data in fig. 3 in addition or instead of panel D. Similarly, the half-relaxation time data should be added.

    Response: We will add the twitch-to-tetanus ratio and half-relaxation time measurements to the revised figures. We will also revise the text to allow for more nuanced interpretation that includes other potential explanations for our results in light of Reviewer 3's comments about altered tissue composition in the tail cross section.

    Minor comment 5: In the methods section, it would be helpful for non-experts to indicate that the MANDRA-1 antibody detects dystrophin.

    Response: We agree, and will be more explicit about this in the Methods.

    Minor comment 6: Additional details on how the cross-sectional area was determined should be added to the methods section.

    Response: We will add: "Because zebrafish larvae are semi-translucent at this stage, sarcomere length was measured using an inverted microscope positioned beneath the force-length apparatus. Once secured, specimens were adjusted to a sarcomere length of 1.9 µm, confirmed by visualization of the thick and thin filament striations. Total specimen length was then measured at low magnification (4x), allowing the entire specimen to be viewed within a single field of view. Next, three transverse dimensions were measured at higher magnification (40x) at equally spaced positions along the specimen, centered on the anal vent (Fig. 3A′). Using a pair of prisms integrated into the mounting chamber, the orthogonal transverse dimension was measured at the same three positions from the dorsoventral perspective. Assuming an elliptical cross-section, the paired orthogonal measurements at each position were used to calculate cross-sectional area. The mean of the three cross-sectional area measurements was used as the specimen CSA."

    Minor comment 7: While this is the first report linking urotensin signaling to muscle function in idiopathic scoliosis, previous research has shown a role for urotensin signaling in skeletal muscle metabolism (PMID: 35605093, PMID: 24116164, PMID: 36167277). The findings presented in these studies and how they relate or could help interpret the findings of this study should be discussed.

    Response: We thank the reviewer for pointing out these studies. In the revised Discussion, we will mention these studies with the broader point that Urotensin signaling can influence skeletal muscle metabolism.

    __Minor comment 8: __Previous studies imply that uts2r3 is expressed in dorsal skeletal muscle, which would imply that dorsal and ventral muscles are differently affected in uts2r3 mutants. The authors should discuss this asymmetric expression of uts2r3 in skeletal muscle and its relevance for the observed phenotype. Furthermore, it would be helpful to include future directions on how to dissect differences between dorsal and ventral muscle activity.

    Response: In our revised Discussion, we will bring up the dorsal-enrichment of uts2r3 and its possible relevance to the curvature in uts2r3 mutants. We will also mention the caveats associated with that expression result, namely, it was seen only in a small number of transient transgenic zebrafish. We will note that our intact tail mechanical measurements integrate dorsal and ventral, superficial and deep, and can be influenced by non-contractile tissues. This is advantageous as it provides a tissue-level view of mechanical properties. We will emphasize that future work could involve region- and fiber type-specific mechanical measurements and tissue-specific perturbations, experiments which are currently beyond our scope.

    Significance: Despite the relatively high prevalence of scoliosis in the human population (2-3%) our understanding of the underlying pathomechanisms are limited. To my knowledge this is the first study that shows that changes in muscle biomechanical function proceed the formation of spinal curvature in a model of idiopathic scoliosis. The study advances our understanding of the mechanisms underlying the formation of spinal curvature. If early changes in biomechanical properties of skeletal muscles are found to be a general feature of idiopathic scoliosis, this study could be impactful for the early detection and potentially treatment of idiopathic scoliosis.

    Therefore, these findings will be of interest to researchers focused on basic science but also for translational and clinical research.

    A major strength of the study is that the observed transcriptional changes could be linked to functional consequences through biomechanical testing of muscle function. However, the pathways that link urotensin signaling to the observed alterations in gene expression and muscle mechanical properties were not explored. Previous reports show uts2r3 expression in dorsal skeletal muscle but are limited to one early time point and analyses of mosaic transgene expression. As the presented study explores uts2r3 function in skeletal muscle, detailed analyses of uts2r3 expression at different stages and in different muscle subpopulations would help interpret the results and significantly improve the study.

    Response: We appreciate the reviewer's assessment of the novelty and potential significance of the pre-curvature biomehanical phenotype.

    Expertise: zebrafish genetics, skeletal development, scoliosis, CRISPR/Cas9, transcriptomics.

    Reviewer #2

    Evidence, reproducibility and clarity

    In this manuscript, the authors build upon previous findings showing that disruption of Urp1/Urp2 signaling, or loss of their receptor Uts2r3, results in abnormalities in spine morphology. The authors note that Urp1 and Urp2 are expressed by cerebrospinal fluid-contacting neurons surrounding the central canal, whereas Uts2r3 is expressed in skeletal muscle. To gain mechanistic insight into how this pathway regulates spine development, they performed bulk RNA-sequencing of 28 hpf Uts2r3 mutant embryos, prior to the onset of spinal deformities. Their analysis identified changes in the expression of several muscle-related genes. Although they did not detect major structural abnormalities in skeletal muscle, they found altered muscle force production, suggesting that impaired muscle function may contribute to the spinal phenotype. While the study addresses an interesting biological question, I have several concerns regarding the strength of the mechanistic conclusions.

    Response: We thank the reviewer for identifying the central mechanistic issue. We agree that some statements in the manuscript should be modified to distinguish the observed pre-curvature biomechanical phenotype from stronger claims of tissue autonomy or a particular molecular mechanism. This distinction will be made explicit in our revision.

    Major comment 1: Expression pattern of Uts2r3. It remains unclear where Uts2r3 is expressed. Apart from the statement that it is present in skeletal muscle, I could not find a detailed characterization of its expression pattern. This information is essential for interpreting the specificity of the phenotype and for understanding whether the observed defects arise directly from muscle or from additional tissues expressing the receptor.

    Response: We agree that the current wording is too broad. We will expand the expression analysis as described in our response to Reviewer 1, including in situ hybridization, immunofluorescence, and single cell transcriptomic data. We will modify statements that currently describe Uts2r3 as 'muscle-expressed' to distinguish supported dorsal slow muscle expression from other tissues, where expression is less certain. We will also make it clear that the current experiments do not establish that our measured biomechanical phenotypes are direct muscle cell-autonomous consequences.

    Major comment 2: Transcriptomic analysis. The RNA-seq analysis lacks important context. I could not determine how many genes were significantly differentially expressed in the mutant embryos or what the overall transcriptional changes were. The manuscript focuses primarily on muscle-related genes, but it would be important to know whether these represent a small subset of widespread transcriptional changes or whether the effects are relatively specific to muscle. Presenting the complete dataset, together with pathway enrichment analyses, would help place the findings into a broader biological context.

    Response: We apologize for the lack of context we provided. In our revised manuscript, we will provide the numbers of differentially expressed genes (downregulated and upregulated), a more broad enrichment analysis with less redundancy between categories, and a complete supplementary table of all gene expression changes with fold changes.

    Major comment 3: Mechanism of Urp1/Urp2 signaling. The manuscript does not sufficiently explain how Urp1/Urp2, which are produced by CSF-contacting neurons around the central canal, signal to receptors expressed in skeletal muscle. Is there evidence that these peptides are secreted into the cerebrospinal fluid and reach the musculature? Alternatively, are there intermediate signaling mechanisms? This represents an important mechanistic gap in the proposed model.

    Response: We agree that the physical route by which Urp1/Urp2 peptides reach Uts2r3-positive target cells remain unresolved. In our revised manuscript, we will discuss this issue, and note plausible routes. We will remove any language which implies this route is already known. However, experimentally discovering how Urp1/Urp2 peptides travel to Uts2r3 receptors is far beyond the scope of our study.

    Major comment 4: Relationship between muscle defects and spinal deformity. Although the authors report altered muscle force production, they do not detect obvious structural defects in the muscle. It therefore remains unclear whether the altered muscle function is the primary cause of the spinal phenotype or simply a secondary consequence of broader developmental abnormalities. Response: We agree that we have not established muscle as the cell autonomous cause of this phenomenon. Indeed, tissue-scale biomechanical changes may and often do, derive from multiple factors within composite tissue such as skeletal muscle. In our revised manuscript, we will be clear that we have discovered a pre-curvature axial mechanical state, some aspects of which strongly suggest myofilament antecedents, rather than implying that muscle dysfunction is necessarily a primary cause.O

    Major comment 5: Tissue specificity. A major limitation of the study is that neither the genetic model nor the transcriptomic analysis is tissue-specific. The Uts2r3 mutation affects the whole embryo, and the RNA-seq was performed on whole embryos rather than isolated muscle. Consequently, it is difficult to determine whether the observed transcriptional changes reflect direct effects within skeletal muscle or indirect systemic effects of the mutation. Tissue-specific genetic manipulation and muscle-specific transcriptomic analyses would considerably strengthen the mechanistic conclusions.

    Response: We agree with the reviewer and we have attempted tissue-specific experiments including muscle-cell specific Cas9 expression. However, this has failed for technical reasons, which we have learned is not uncommon. We agree that the present data do not establish muscle cell autonomy and we will state this explicitly throughout the manuscript. Nevertheless, we consider tissue-specific experiments to constitute a substantial new mechanistic study rather than a necessary control for our existing biomechanical phenotype. In our revised version, we will be clear to note the supported conclusion that whole animal loss of Uts2r3 produces pre-curvature axial mechanical defects.

    Overall assessment: The study extends previous work implicating the Urp1/Urp2-Uts2r3 pathway in regulating spine morphology and provides evidence that altered muscle function may contribute to this phenotype. However, the current data provide only limited mechanistic insight into how this signaling pathway regulates muscle physiology or spine development. In particular, the lack of tissue-specific genetic and transcriptomic analyses makes it difficult to distinguish direct muscle-specific effects from broader developmental consequences of Uts2r3 deficiency. Additional experiments addressing these issues would substantially strengthen the manuscript.

    Response: We appreciate these comments and will revise the manuscript accordingly. Our goal in the revision will be to make the mechanistic level of the study precise: this paper identifies and quantitatively defines a pre-curvature biomechanical state downstream of Uts2r3 loss. It does leave the precise route of Urp1/Urp2 signaling and the cell autonomous response as open molecular questions, which are beyond the scope of our current work, and which future work will aim to address.

    Reviewer #3

    Evidence, reproducibility and clarity

    This manuscript uses the zebrafish model system, integrating genetic, histological, transcriptomic, and functional approaches to address a challenging and relevant biological question. The authors conclude from their data that urotensin signaling defects affect both passive and active mechanical properties of the larval zebrafish's muscular tail independently, creating an imbalance of forces during development that ultimately leads to a scoliosis-like phenotype. Previous studies in zebrafish had linked urotensin signaling to the developing trunk muscles through the receptor Uts2r3. The present study shows that loss of Uts2r3 leads to changes in muscle specific gene expression, loss of active myotomal muscle force, and increase in passive viscoelastic forces, without overt morphological or dystrophic changes to in the muscle. Importantly, these changes are seen prior to the development of the tail-curvature phenotype.

    Response: We thank the reviewer for highlighting the integrated strength of our study and the importance of measuring active and passive responses prior to curvature.

    Major comment 1: The authors predict involvement of myotomal muscles in the progression of the curvature phenotype based on the reported expression of the Uts2r3 receptor in myotomal muscles during development. According to the cited studies, the receptor is expressed specifically in dorsal slow-twitch muscle fibers (Zhang 2018). Supporting this, the upward tail bending effect of urotensin overexpression is abolished in mutant larvae lacking slow twitch fibers (lu 2020, Gillard 2023). The assertion from these studies being that urotensin regulates tail curvature via regulating dorsal slow muscle tone. Too much signaling increases force and bends the tail upwards; two little has the opposite effect. This interpretation is relevant because slow twitch fibers make up only ~7% of the total myotomal muscle CSA in 5dpf larvae and are not thought to contribute meaningfully to the high speed forces measured using the intact tail mechanics approach. The remaining >90% of CSA consists of fast twitch muscle active only during brief escape maneuvers. According to the same studies, fast fibers do not appear to express Uts2r3. If this is the case, the reported loss of force is more challenging to interpret. As written, fiber type is not mentioned. This is an issue that should be discussed and considered when interpreting the data.

    Response: We agree and consider this an important point. The revised manuscript will explicitly address the potential slow-twitch exclusive expression of the Uts2r3 receptor and how such selective, fiber-type specific targeting in this study may alter interpretation of our findings. We do note, that the previous findings showing dorsal slow-twitch enrichment were based on a small number of transient transgenic zebrafish, meaning expression in other locations was likely missed. We are performing a broader expression analysis using stains as well as single cell transcriptomics. We will therefore test whether there is support for additional expression in presomitic mesoderm or fast muscle populations, and interpret our findings accordingly, in line with the reviewer's suggestions. In any case, a defect present in ~10% of the transactional cross section may still have significant impacts on chronic stress applied to the developing spinal column, especially if that expression is limited to more chronically active, slow-twitch fibers. These important caveats to our initial interpretation will be included in the revised manuscript.

    __Major comment 2: __The authors conclude that twitch and tetanus force deficits in Uts2r3 mutant larvae result from changes in contractility at the sarcomere level. Mis regulation of muscle gene expression and mild loss of structural organization are cited as causes. An alternative explanation, overall reduction in contractile CSA, is ruled out based on estimates from whole animal dimensions (see specific comments below). There are a couple of reasons why this may still be the simplest explanation. First, myotomal muscle accounts for ~70% of the total CSA of the tail, with myofibril CSA a large sub-fraction of that. Therefore, significant changes in the CSA relevant to contractile force may be difficult to resolve from whole-body estimates. This is especially true if increases in passive stiffness result from increased CSA of connective tissue, etc. Second, birefringence is a good measure of damage or structural disruptions to myotomal muscles that are heterogeneous within a sample, as in models of DMD. Uniform reduction in intensity (Fig S2) may result from disruption of the regular order of myosin heads within the sarcomere but could also reflect a reduction in myofibrillar volume. Third, dynamic measurements of contractile function (twitch kinetics, F/V) appear to be unchanged. Disruption of sarcomere organization, a shift in contractile protein isoforms (such as myosin heavy chain), or other changes affecting actomyosin function would be more likely to show up in the kinetics, whereas changes in myofibril CSA would not. Finally, to the extent mRNA levels represent protein abundance, the data presented also support a 'less muscle' explanation. There are experiments that could get at these questions directly, such as a histological examination of tail cross sections. But for the purposes of this study, a more thorough discussion of these possibilities would be sufficient.

    Response: We agree that the submitted manuscript likely overinterprets myotome morphology and whole body dimensions to assume similar muscle transverse cross sectional area. A whole preparation CSA estimate cannot distinguish myofibrillar tissue from other axial tissues, and the uniform reduction in birefringence is indeed compatible with more than one structural interpretation. We are therefore examining matched transverse sections to quantify myotomal area and connective tissue composition via histology. We are also evaluating protein/MyHC measurements via electrophoretic separation as an independent readout of muscle composition, normalized to the measured specimen volume. These data will be included if they are sufficiently quantitative and reproducible and our pilot experiments are promising. Irrespective of the outcome of these additional experiments, we will remove the claim that the force deficit necessarily reflects a sarcomere-level contractile defect, and the revised discussion will explicitly consider the other explanations the reviewer suggests — for which we are grateful.

    Major comment 3: The authors conclude that 'active and passive defects are unlikely to be simple consequences of one another', claiming 'increased passive stiffness does not readily explain the reduction in active force generation.' This may be true but the issue is complex and needs more discussion. The mechanical approach used (Mead 2020) is quite different from how the tail functions in vivo and comes with some caveats. One observation in that study is that 'isometric' contractions, in which the preparation is held at a fixed length, result in the shortening of contracting myotomes by about 10%, accompanied by the stretching of tissue near the tail attachment points. Because of this, there are a couple of ways in which changes in passive properties could impact measured forces in this assay. For example, the notochord is a structure that functions to resist tail compression during swimming in vivo. So, any stiffening of the notochord, or of connections between myotomal muscle and notochord could reduce muscle forces transmitted to the force transducer. Also, unless tails were pre-stretched by ~10% before activation, as in Mead 2020, forces reported here may have been generated at muscle lengths on the ascending limb of the length-tension curve. Therefore, changes in the passive mechanical properties of the tissue at the attachment points could have large effects on measured force by changing the SL at which force is measured. This could be checked if SL or myotome length was measured during contractions as was done in that study. The paper would be strengthened if the interdependence of active and passive mechanical changes were considered in this light.

    Response: We thank the reviewer for these helpful comments. We will remove the statement that the active and passive abnormalities are unlikely to be consequences of one another and include the more complete and potential interrelated nature of these measures, considering notochord involvement as suggested by the reviewer. We will therefore treat the preparation as an integrated mechanical system in which force generation, shortening, myoseptal and notochord mechanics, tissue compliance, and force transmission all interact. We will point out that specimens were secured to the force/length apparatus in these experiments using an approach distinct from Mead et al 2020, and this may have reduced compliance to some degree, as we observed but sarcomere tracking at these speeds was not tenable in our hands. Further, during active measurements, specimens were initially adjusted to a sarcomere length of 1.9 µm before stimulation. This was accomplished by stretching the specimen to ~1.95 or 2.0 µm and then reducing the length slightly. Because the specimens resist further compression beyond 1.9 µm, this at least ensures specimens were as close as possible to optimal contractile length. Because the same specimens were stretched prior to activation during the kinetic experiments to measure force deficits at varying velocities, we will review these data to see if greater strain produced further differences in isometric tension prior to shortening. This will help guide interpretation as to whether resistance to notochord compression influenced active contractile deficits. e While we believe these additional considerations and measures may support the notion that muscle contractile function is altered in Uts2r3 mutants, we cannot exclude the possibility that passive properties alter effective muscle operating length or force transmission and will state this limitation directly in the revised manuscript. We will also discuss how it may change the interpretation of reduced measured force.

    Minor comment 1: Are the Uts2r3 mutant fish used in the study heterozygotes or homozygous null? This line was previously described, but it would be helpful if the type of mutation as well as the when and how the phenotype develops are at least briefly described here as well.

    Response: All mutants are homozygous mutants (b1436) which lack Uts2r3. In the results, when the mutant is first mentioned, we will describe the allele, rather than requiring readers to retrieve the previous publication.

    Minor comment 2: Mechanical measurements are reported as being performed as described in Mead 2020. Deviations are mentioned in method of attachment, instrumentation, and SL measurement. However, it would be helpful to have a more thorough description of the approach. Specific parameters used should be included. For example: length of preparations, any anatomical landmarks for suture placement, range of muscle lengths used for isovelocity maneuvers, etc.

    Response: We agree, and will enlarge our Methods descriptions including specimen preparation and length, suture landmarks, bath composition and temperature, starting sarcomere length, stimulation parameters, shortening velocities and imposed length changes, and the number of specimens contributing to each assay.

    Minor comment 3: Force velocity data (Fig. 3E) shows a force depression at 28 l/sec of only ~30% at 18˚C. This result predicts a Vmax over 100 l/sec at 18C, and possibly twice that at physiologically relevant temperatures, assuming a q10 of 2. Mead et al (2020 and 2024) predicted a Vmax of 45 l/sec at 28˚C and reported ~80% drop in force at ~12 l/sec at 23˚C, stating that limitations inherent to the instrumentation and the tail preparation precluded accurate measurement at higher velocities. Can the authors comment on the discrepancy? Were any modifications made to the approach in order to access these higher velocities?

    Response: We will address this discrepancy in our resubmission. We plan to double check the original code, including exact length changes, velocity normalization, and the isometric contractions used to normalize force. We will reanalyze force-velocity data and compare the resulting relationship directly with Mead et al. If the expected Vmax is not obtained, we will highlight this discrepancy with appropriate assumptions and limitations. In this case, we will revise any interpretations that implies physiologically accurate Vmax at the highest imposed velocities.

    __Minor comment 4: __How were CSA measurements made? Methods states tails were measured in two dimensions at three points. Were anatomical landmarks used? These should be included.

    Response: We agree, and as also noted in response to Reviewer 1, we will add the exact measurement procedure and anatomical reference points. Additionally, we are significantly expanding our histology and CSF analysis in our resubmission, and we will be sure to include detailed methods to address these points.

    Minor comment 5: Stated sample numbers do not match the number of individual points in figures 3 B, C, D, E. P value for tetanus tension stated in text (0.008) does not match Fig 3 C (0.014). The next sentence appears redundant and states a third p value of Response: We apologize for these inconsistencies. Due to technical limitations at the time of data collection, tension data were not recorded in all of the same specimens at all velocities. Similarly, passive mechanics were collected in some specimens but not others. Last, CSA data were lost on one sample, contributing non-normalized data, but no CSA normalized data (tension). We will be more explicit about data inclusion/exclusion for these figures and tables in the revised text, and re-run statistical analysis, then correct inconsistent p values in figures and in the text.

    Minor comment 6: Will RNA seq data be reported in a supplemental file or uploaded to a database? Both are standard practice.

    Response: Yes, we plan to provide a table of all differentially expressed genes and their fold changes and adjusted p values.

    Minor comment 7: The primary curvature defect in the Uts2r3 mutant seems to be dorsal-ventral in nature, though this is not explicitly described in the text. Can any insight into this or any other specific feature of the phenotype be gained from the data? Why down and not up or left vs right? The data presented shows no change in the relative 2D area of hemisomites resulting from this curvature. However, geometry would indicate some remodeling (different-modeling?) must take place in these mutant fish. Is possible to measure either muscle fiber length, sarcomere length, or sarcomere number from these images?

    Response: We will describe the phenotype more explicitly, instead of relying on readers accessing the earlier publication. The current hemisomite data argue against a gross dorsoventral difference in 2D muscle area or myoseptal angle, but they do not explain why the curve is ventrally directed. We will state in our discussion that our present data do not resolve the directional basis of the curve phenotype. We agree with the reivewer that this is worth discussing as it is a particularly interesting aspect of the Urotensin-associated curve phentotype.

    Minor comment 8: Fig S3B is very hard to see. Can 'hemisomite abstraction overlay' be defined in the figure caption?

    Response: Apologies. We will improve the visibility of the figure and be explicit about definitions in the figure legend.

    Significance: The main strength of this study is in exploiting the highly integrative nature of the zebrafish model to explore a challenging and relevant biological question. The methods used here, in particular the mechanical experiments, are extremely challenging to perform an interpret. The data presented generally support the conclusions, but important context is missing. A more nuanced discussion of potential mechanisms and experimental caveats will better inform subsequent experiments and improve the overall impact of the paper. This study has the potential to advance the field by incorporating direct mechanical measurements of active and passive muscle force in a genetically tractable vertebrate model system. It will be of interest to the fields of muscle development and regulation, tissue mechanics, scoliosis research. My main expertise is in comparative physiology, evolution, and biophysics of muscle contraction. I have experience with a wide range of model species, including zebrafish. I have a more recent interest in muscle development and repair. I am not familiar with the fields of scoliosis research and urotensin signaling.

    Response: We appreciate this assessment and agree with the proposed direction of revision. The revised paper will place the direct mechanics at the center of the study while adding the missing physiological context and being more explicit about what an intact axial preparation can and and cannot resolve.

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    Referee #3

    Evidence, reproducibility and clarity

    Summary

    This manuscript uses the zebrafish model system, integrating genetic, histological, transcriptomic, and functional approaches to address a challenging and relevant biological question. The authors conclude from their data that urotensin signaling defects affect both passive and active mechanical properties of the larval zebrafish's muscular tail independently, creating an imbalance of forces during development that ultimately leads to a scoliosis-like phenotype. Previous studies in zebrafish had linked urotensin signaling to the developing trunk muscles through the receptor Uts2r3. The present study shows that loss of Uts2r3 leads to changes in muscle specific gene expression, loss of active myotomal muscle force, and increase in passive viscoelastic forces, without overt morphological or dystrophic changes to in the muscle. Importantly, these changes are seen prior to the development of the tail-curvature phenotype.

    Major comments

    1. The authors predict involvement of myotomal muscles in the progression of the curvature phenotype based on the reported expression of the Uts2r3 receptor in myotomal muscles during development. According to the cited studies, the receptor is expressed specifically in dorsal slow-twitch muscle fibers (Zhang 2018). Supporting this, the upward tail bending effect of urotensin overexpression is abolished in mutant larvae lacking slow twitch fibers (lu 2020, Gillard 2023). The assertion from these studies being that urotensin regulates tail curvature via regulating dorsal slow muscle tone. Too much signaling increases force and bends the tail upwards; two little has the opposite effect. This interpretation is relevant because slow twitch fibers make up only ~7% of the total myotomal muscle CSA in 5dpf larvae and are not thought to contribute meaningfully to the high speed forces measured using the intact tail mechanics approach. The remaining >90% of CSA consists of fast twitch muscle active only during brief escape maneuvers. According to the same studies, fast fibers do not appear to express Uts2r3. If this is the case, the reported loss of force is more challenging to interpret. As written, fiber type is not mentioned. This is an issue that should be discussed and considered when interpreting the data.

    2. The authors conclude that twitch and tetanus force deficits in Uts2r3 mutant larvae result from changes in contractility at the sarcomere level. Mis regulation of muscle gene expression and mild loss of structural organization are cited as causes. An alternative explanation, overall reduction in contractile CSA, is ruled out based on estimates from whole animal dimensions (see specific comments below). There are a couple of reasons why this may still be the simplest explanation. First, myotomal muscle accounts for ~70% of the total CSA of the tail, with myofibril CSA a large sub-fraction of that. Therefore, significant changes in the CSA relevant to contractile force may be difficult to resolve from whole-body estimates. This is especially true if increases in passive stiffness result from increased CSA of connective tissue, etc. Second, birefringence is a good measure of damage or structural disruptions to myotomal muscles that are heterogeneous within a sample, as in models of DMD. Uniform reduction in intensity (Fig S2) may result from disruption of the regular order of myosin heads within the sarcomere but could also reflect a reduction in myofibrillar volume. Third, dynamic measurements of contractile function (twitch kinetics, F/V) appear to be unchanged. Disruption of sarcomere organization, a shift in contractile protein isoforms (such as myosin heavy chain), or other changes affecting actomyosin function would be more likely to show up in the kinetics, whereas changes in myofibril CSA would not. Finally, to the extent mRNA levels represent protein abundance, the data presented also support a 'less muscle' explanation. There are experiments that could get at these questions directly, such as a histological examination of tail cross sections. But for the purposes of this study, a more thorough discussion of these possibilities would be sufficient.

    3. The authors conclude that 'active and passive defects are unlikely to be simple consequences of one another', claiming 'increased passive stiffness does not readily explain the reduction in active force generation.' This may be true but the issue is complex and needs more discussion. The mechanical approach used (Mead 2020) is quite different from how the tail functions in vivo and comes with some caveats. One observation in that study is that 'isometric' contractions, in which the preparation is held at a fixed length, result in the shortening of contracting myotomes by about 10%, accompanied by the stretching of tissue near the tail attachment points. Because of this, there are a couple of ways in which changes in passive properties could impact measured forces in this assay. For example, the notochord is a structure that functions to resist tail compression during swimming in vivo. So, any stiffening of the notochord, or of connections between myotomal muscle and notochord could reduce muscle forces transmitted to the force transducer. Also, unless tails were pre-stretched by ~10% before activation, as in Mead 2020, forces reported here may have been generated at muscle lengths on the ascending limb of the length-tension curve. Therefore, changes in the passive mechanical properties of the tissue at the attachment points could have large effects on measured force by changing the SL at which force is measured. This could be checked if SL or myotome length was measured during contractions as was done in that study. The paper would be strengthened if the interdependence of active and passive mechanical changes were considered in this light.

    Minor comments

    1. Are the Uts2r3 mutant fish used in the study heterozygotes or homozygous null? This line was previously described, but it would be helpful if the type of mutation as well as the when and how the phenotype develops are at least briefly described here as well.

    2. Mechanical measurements are reported as being performed as described in Mead 2020. Deviations are mentioned in method of attachment, instrumentation, and SL measurement. However, it would be helpful to have a more thorough description of the approach. Specific parameters used should be included. For example: length of preparations, any anatomical landmarks for suture placement, range of muscle lengths used for isovelocity maneuvers, etc.

    3. Force velocity data (Fig. 3E) shows a force depression at 28 l/sec of only ~30% at 18˚C. This result predicts a Vmax over 100 l/sec at 18C, and possibly twice that at physiologically relevant temperatures, assuming a q10 of 2. Mead et al (2020 and 2024) predicted a Vmax of 45 l/sec at 28˚C and reported ~80% drop in force at ~12 l/sec at 23˚C, stating that limitations inherent to the instrumentation and the tail preparation precluded accurate measurement at higher velocities. Can the authors comment on the discrepancy? Were any modifications made to the approach in order to access these higher velocities?

    4. How were CSA measurements made? Methods states tails were measured in two dimensions at three points. Were anatomical landmarks used? These should be included.

    5. Stated sample numbers do not match the number of individual points in figures 3 B, C, D, E. P value for tetanus tension stated in text (0.008) does not match Fig 3 C (0.014). The next sentence appears redundant and states a third p value of < 0.05.

    6. Will RNA seq data be reported in a supplemental file or uploaded to a database? Both are standard practice.

    7. The primary curvature defect in the Uts2r3 mutant seems to be dorsal-ventral in nature, though this is not explicitly described in the text. Can any insight into this or any other specific feature of the phenotype be gained from the data? Why down and not up or left vs right? The data presented shows no change in the relative 2D area of hemisomites resulting from this curvature. However, geometry would indicate some remodeling (different-modeling?) must take place in these mutant fish. Is possible to measure either muscle fiber length, sarcomere length, or sarcomere number from these images?

    8. Fig S3B is very hard to see. Can 'hemisomite abstraction overlay' be defined in the figure caption?

    Significance

    The main strength of this study is in exploiting the highly integrative nature of the zebrafish model to explore a challenging and relevant biological question. The methods used here, in particular the mechanical experiments, are extremely challenging to perform an interpret. The data presented generally support the conclusions, but important context is missing. A more nuanced discussion of potential mechanisms and experimental caveats will better inform subsequent experiments and improve the overall impact of the paper.

    This study has the potential to advance the field by incorporating direct mechanical measurements of active and passive muscle force in a genetically tractable vertebrate model system. It will be of interest to the fields of muscle development and regulation, tissue mechanics, scoliosis research.

    My main expertise is in comparative physiology, evolution, and biophysics of muscle contraction. I have experience with a wide range of model species, including zebrafish. I have a more recent interest in muscle development and repair. I am not familiar with the fields of scoliosis research and urotensin signaling.

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    Referee #2

    Evidence, reproducibility and clarity

    In this manuscript, the authors build upon previous findings showing that disruption of Urp1/Urp2 signaling, or loss of their receptor Uts2r3, results in abnormalities in spine morphology. The authors note that Urp1 and Urp2 are expressed by cerebrospinal fluid-contacting neurons surrounding the central canal, whereas Uts2r3 is expressed in skeletal muscle. To gain mechanistic insight into how this pathway regulates spine development, they performed bulk RNA-sequencing of 28 hpf Uts2r3 mutant embryos, prior to the onset of spinal deformities. Their analysis identified changes in the expression of several muscle-related genes. Although they did not detect major structural abnormalities in skeletal muscle, they found altered muscle force production, suggesting that impaired muscle function may contribute to the spinal phenotype. While the study addresses an interesting biological question, I have several concerns regarding the strength of the mechanistic conclusions.

    Major comments

    1. Expression pattern of Uts2r3 It remains unclear where Uts2r3 is expressed. Apart from the statement that it is present in skeletal muscle, I could not find a detailed characterization of its expression pattern. This information is essential for interpreting the specificity of the phenotype and for understanding whether the observed defects arise directly from muscle or from additional tissues expressing the receptor.

    2. Transcriptomic analysis The RNA-seq analysis lacks important context. I could not determine how many genes were significantly differentially expressed in the mutant embryos or what the overall transcriptional changes were. The manuscript focuses primarily on muscle-related genes, but it would be important to know whether these represent a small subset of widespread transcriptional changes or whether the effects are relatively specific to muscle. Presenting the complete dataset, together with pathway enrichment analyses, would help place the findings into a broader biological context.

    3. Mechanism of Urp1/Urp2 signaling The manuscript does not sufficiently explain how Urp1/Urp2, which are produced by CSF-contacting neurons around the central canal, signal to receptors expressed in skeletal muscle. Is there evidence that these peptides are secreted into the cerebrospinal fluid and reach the musculature? Alternatively, are there intermediate signaling mechanisms? This represents an important mechanistic gap in the proposed model.

    4. Relationship between muscle defects and spinal deformity Although the authors report altered muscle force production, they do not detect obvious structural defects in the muscle. It therefore remains unclear whether the altered muscle function is the primary cause of the spinal phenotype or simply a secondary consequence of broader developmental abnormalities.

    5. Tissue specificity A major limitation of the study is that neither the genetic model nor the transcriptomic analysis is tissue-specific. The Uts2r3 mutation affects the whole embryo, and the RNA-seq was performed on whole embryos rather than isolated muscle. Consequently, it is difficult to determine whether the observed transcriptional changes reflect direct effects within skeletal muscle or indirect systemic effects of the mutation. Tissue-specific genetic manipulation and muscle-specific transcriptomic analyses would considerably strengthen the mechanistic conclusions.

    Overall assessment The study extends previous work implicating the Urp1/Urp2-Uts2r3 pathway in regulating spine morphology and provides evidence that altered muscle function may contribute to this phenotype. However, the current data provide only limited mechanistic insight into how this signaling pathway regulates muscle physiology or spine development. In particular, the lack of tissue-specific genetic and transcriptomic analyses makes it difficult to distinguish direct muscle-specific effects from broader developmental consequences of Uts2r3 deficiency. Additional experiments addressing these issues would substantially strengthen the manuscript.

    Significance

    Overall assessment

    The study extends previous work implicating the Urp1/Urp2-Uts2r3 pathway in regulating spine morphology and provides evidence that altered muscle function may contribute to this phenotype. However, the current data provide only limited mechanistic insight into how this signaling pathway regulates muscle physiology or spine development. In particular, the lack of tissue-specific genetic and transcriptomic analyses makes it difficult to distinguish direct muscle-specific effects from broader developmental consequences of Uts2r3 deficiency. Additional experiments addressing these issues would substantially strengthen the manuscript.

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    Referee #1

    Evidence, reproducibility and clarity

    The work by O'Hara-Smith et al. explores if alterations in muscle development and function underly the formation of spinal curvature in zebrafish uts2r3 mutants. Using transcriptomic analyses, immunofluorescence and birefringence, the authors reveal a reduction in the expression of muscle-associated genes as early as 28hpf with only mild reduction in birefringence and otherwise no overt changes in muscle morphology or symmetry, even at stages where spinal curvature becomes apparent (14dpf). Biomechanical analyses further show that loss of uts2r3 leads to a reduction in active force and an increased passive resistance to stretch, indicating that urotensin signaling plays a role in the regulation of muscle function.

    Major Comments:

    The paper is well written and easy to follow. The majority of experiments are described in sufficient detail, and all of the used reagents are listed. The conclusions are supported by the presented data. Optional additional experiments that could significantly improve the paper would be a detailed characterization of the expression of uts2r3 as previous expression analyses were limited to one early time point and mostly used analyses of mosaic transgene expression. However, performing detailed expression analyses would be both costly and time consuming.

    Minor Comments:

    1. Additional details on the RNA-seq experiment should be added to the methods section to allow interpretation of the results. Was mRNA selection or ribosome depletion used? What was the sequencing mode (paired or single-end)? What read length was used? How many reads were obtained per sample and how many filtered reads could be aligned to the reference transcriptome?

    2. To understand the extend of the changes detected by the transcriptomic analyses it would be helpful to include the total number of significantly up- and down-regulated genes as well as the magnitude of change (fold-change) for the highlighted muscle related genes.

    3. Similarly, for the GO terms a list of differentially expressed genes for each term should be provided and largely overlapping GO terms should be removed. The fact that all terms show almost the exact same fold enrichment could point to a large overlap between the differentially expressed genes detected for each term.

    4. As the ratio of twitch and tetanic force was quantified and used as a measure for myosin dynamics and force transmission, it would be helpful to show that data in fig. 3 in addition or instead of panel D. Similarly, the half-relaxation time data should be added.

    5. In the methods section, it would be helpful for non-experts to indicate that the MANDRA-1 antibody detects dystrophin.

    6. Additional details on how the cross-sectional area was determined should be added to the methods section.

    7. While this is the first report linking urotensin signaling to muscle function in idiopathic scoliosis, previous research has shown a role for urotensin signaling in skeletal muscle metabolism (PMID: 35605093, PMID: 24116164, PMID: 36167277). The findings presented in these studies and how they relate or could help interpret the findings of this study should be discussed.

    8. Previous studies imply that uts2r3 is expressed in dorsal skeletal muscle, which would imply that dorsal and ventral muscles are differently affected in uts2r3 mutants. The authors should discuss this asymmetric expression of uts2r3 in skeletal muscle and its relevance for the observed phenotype. Furthermore, it would be helpful to include future directions on how to dissect differences between dorsal and ventral muscle activity.

    Significance

    Despite the relatively high prevalence of scoliosis in the human population (2-3%) our understanding of the underlying pathomechanisms are limited. To my knowledge this is the first study that shows that changes in muscle biomechanical function proceed the formation of spinal curvature in a model of idiopathic scoliosis. The study advances our understanding of the mechanisms underlying the formation of spinal curvature. If early changes in biomechanical properties of skeletal muscles are found to be a general feature of idiopathic scoliosis, this study could be impactful for the early detection and potentially treatment of idiopathic scoliosis.

    Therefore, these findings will be of interest to researchers focused on basic science but also for translational and clinical research.

    A major strength of the study is that the observed transcriptional changes could be linked to functional consequences through biomechanical testing of muscle function. However, the pathways that link urotensin signaling to the observed alterations in gene expression and muscle mechanical properties were not explored. Previous reports show uts2r3 expression in dorsal skeletal muscle but are limited to one early time point and analyses of mosaic transgene expression. As the presented study explores uts2r3 function in skeletal muscle, detailed analyses of uts2r3 expression at different stages and in different muscle subpopulations would help interpret the results and significantly improve the study.

    Expertise: zebrafish genetics, skeletal development, scoliosis, CRISPR/Cas9, transcriptomics

  5. Version published to 10.64898/2026.04.29.721663 on bioRxiv