Immobilization secondary to cell death of muscle precursors with a dual transcriptional signature contributes to the emu wing skeletal pattern

Tsuboi, Eriko; Ono, Satomi F.; Cordeiro, Ingrid Rosenburg; Yu, Reiko; Kawanishi, Toru; Koizumi, Makoto; Shigenobu, Shuji; Sheng, Guojun; Okabe, Masataka; Tanaka, Mikiko

doi:10.1038/s41467-024-52203-x

Immobilization secondary to cell death of muscle precursors with a dual transcriptional signature contributes to the emu wing skeletal pattern

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Published: 19 September 2024

Volume 15, article number 8153, (2024)
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Immobilization secondary to cell death of muscle precursors with a dual transcriptional signature contributes to the emu wing skeletal pattern

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Abstract

Limb reduction has occurred multiple times in tetrapod history. Among ratites, wing reductions range from mild vestigialization to complete loss, with emus (Dromaius novaehollandiae) serving as a model for studying the genetic mechanisms behind limb reduction. Here, we explore the developmental mechanisms underlying wing reduction in emu. Our analyses reveal that immobilization resulting from the absence of distal muscles contributes to skeletal shortening, fusion and left-right intraindividual variation. Expression analysis and single cell-RNA sequencing identify muscle progenitors displaying a dual lateral plate mesodermal and myogenic signature. These cells aggregate at the proximal region of wing buds and undergo cell death. We propose that this cell death, linked to the lack of distal muscle masses, underlines the morphological features and variability in skeletal elements due to reduced mechanical loading. Our results demonstrate that differential mobility during embryonic development may drive morphological diversification in vestigial structures.

Co-option of the cardiac transcription factor Nkx2.5 during development of the emu wing

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Transcriptomic and epigenomic characterization of the developing bat wing

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Limb patterning genes and heterochronic development of the emu wing bud

Article Open access 20 December 2016

Introduction

Limb reduction has occurred multiple times throughout tetrapod evolution. Snakes, legless lizards and caecilians have lost their limbs, cetaceans reduced their hindlimbs, and ungulates evolved a reduced number of digits. The reduction of limbs in tetrapods is associated with the diversification of their locomotion styles and habitats. Therefore, revealing the mechanisms by which limb reduction occurs is one of the main themes of tetrapod limb evolution research¹.

Among birds, the forelimb reductions found in ratites range from the mild vestigialization seen in the ostrich to the complete loss of the forelimb in the extinct moa, and might have occurred multiple times in this clade^2,3. Of the ratites available for developmental approaches, the emu exhibits an extreme forelimb reduction, which is already evident during embryonic as a very reduced forelimb bud⁴. Several groups have addressed mechanisms of forelimb bud reduction during early stages of emu limb development, which has been attributed to decreased proliferation of limb bud progenitors, and also to a heterochrony caused by modulation and/or delayed expression of typical limb patterning genes^{2,5,6,7,8,9,10}, including higher variability in expression pattern, such as Grem1⁸. Nevertheless, a limb bud is formed, which develops into a patterned wing⁴.

However, reduction of the actual limb bud does not explain mechanisms taking place during later developmental stages that further shape the emu wings, especially their autopodial region. While three digits are initially specified in the emu autopod⁷, most of the cartilaginous elements of the digits 2 and 4 are resorbed during late foetal development⁸, which is followed by fusion of skeletal elements and joint contractures that are typical of adult emu wings^8,11. These appendages display an extensive osteological variation, including presence or absence of phalanges, shape, bone pneumaticity, and muscle attachment¹¹. These differences were not restricted to skeletal elements, as it was found that emus lacked several muscle masses in their wings when compared to volant birds as well¹¹.

Interestingly, defects in both bone growth and joint formation can be secondary to a loss of movement during development, which has been extensively studied in both animal models^{12,13,14,15,16} and clinical examples¹⁷. Limb skeletal defects have been reported in several strains of muscle-less and muscle dysgenesis mice, while avian in ovo models have been crucial to pinpoint the effects of early or late immobilization on longitudinal skeletal growth and joint cavitation^{13,14,15,16,18,19,20,21,22,23,24,25,26,27,28,29,30}. In human patients, a decrease in fetal movement can lead to fetal akinesia deformation sequence, a spectrum of skeletal and joint phenotypes¹³. Furthermore, congenital joint contractures often present with a range of severity, including variation in the affected joints and, to a lesser extent, asymmetric presentation—even when the cause of the disease has been mapped to a specific mutation¹⁷. The highly variable range of phenotypes found in emu wings led us to hypothesize whether it was not only the result of direct genetic factors but also secondary to extrinsic epigenetic factors—more specifically, to a reduced mechanical stimulation.

Here, we show that emu forelimbs exhibit a high intraindividual asymmetry in skeletal patterns, in addition to a reduction in bone length and contractures of joints. Such morphological features seen in emu forelimbs recapitulate the skeletal phenotype found in both experimental and clinical observations of reduced mechanical loading during limb development. In addition, we observe a lack of distal limb movement in emu embryos resulting from the lack of muscle masses at the distal ends of their wings, a condition that is apparent from the stages of prenatal development. Single-cell RNA sequencing (scRNA-seq) data and expression analysis revealed that a subpopulation of muscle progenitors exhibits a dual myogenic/LPM (lateral plate mesoderm) transcriptional signature. This subpopulation undergoes massive cell death and thereby fails to form distal skeletal muscles, which contributes to the unique morphology of emu forelimbs.

Results

High intraindividual left-right variation is found in the adult skeletal pattern of the distal emu wings

Previously, we and others have shown that a great range of interindividual variation exists in the digital pattern of adult emu forelimbs^8,11. Understanding whether the variation is present within limbs of the same individual, or is consistent between left and right limbs, would help elucidate the source of variation itself. Here, a comparison between the pair of forelimbs of eight adult emu specimens revealed an intraindividual, left–right asymmetric skeletal pattern (Fig. 1a; Supplementary Fig. 1a). The variation included different status regarding fusion of the ulna and radius, fusion of autopodial skeletal elements, fusion of joints and bone pneumaticity between left and right limbs of the same individual (Fig. 1a). Furthermore, the humeral, ulnar and metacarpal lengths were relatively asymmetric in all examined emu forelimbs compared to chicken forelimbs (Fig. 1b; Supplementary Fig. 1b). Thus, there is a high degree of intra- and interindividual variability intrinsic to emu wing skeletal elements.

**Fig. 1: Skeletal and muscular patterns of the forelimb of emu and chicken.**

Developing emu forelimbs receive less mechanical input due to the absence of distal muscle masses

Studies in chicken and mouse embryos revealed that longitudinal bone growth of the limb and normal joint cavity formation require mechanical input as a result of embryonic muscle contraction^{13,19,20,21,22,23,24,25,26,27,28,29}. Importantly, a marked reduction in the number of muscle masses is found in adult emu wings, especially towards its distal part¹¹, although the remaining muscles display a typical skeletal muscle morphology (Supplementary Fig. 1c). A normal gross morphology of the forelimb muscle masses has been reported in stage 30 emu forelimbs⁷; however, it remains unclear when muscle reduction occurs during autopodial development, which takes place at later stages. To address this question, we analyzed the process of muscle formation in developing forelimb buds of emu embryos and compared them to those in chicken embryos (Fig. 1c; Supplementary Fig. 2a). At stage 30, both chicken and emu display the typical avian forelimb structure containing three digits⁷. While no autopodial muscle mass could be detected at this stage (Supplementary Fig. 2a), they became evident at stages 33, 35 and 37 (Fig. 1c; Supplementary Fig. 2a). In contrast, digits 2 and 4 of emu wings displayed various degrees of vestigialization, which takes place up to stage 37⁸. No autopodial muscles were observed, and most zeugopodial muscles were missing from developing emu forelimb buds up to stage 37 (Fig. 1c; Supplementary Fig. 2a). By recording embryonic movement in ovo, we also found that the distal part of forelimb buds showed hardly any movement in emu embryos at stage 39, while embryonic muscle contraction was already evident in the distal forelimb buds in chicken embryos or the proximal portion of emu wings at the same stage (Fig. 1d; Supplementary Fig. 2b). These results demonstrate that the distal part of the emu forelimb receives less mechanical input during its development in connection to the absence of distal muscle formation.

Loss of mobility recapitulates characteristic features of emu forelimbs

Much research has been devoted to understanding the effect of muscle contraction on the growth of limb bones and joint cavity formation in avian limbs^{13,15,16,24,25,26,27,28,29,30,31,32}. Although the decreased mechanical load secondary to the absence of distal muscle could explain the left–right variation seen in the emu forelimb skeleton, to our knowledge, there have been no reports demonstrating the effect of immobilization specifically on intraindividual left–right asymmetric patterning of limbs. Here, we will address these questions by first focusing on the effect of late-stage immobilization (E10-18) on the longitudinal growth of individual skeletal elements, followed by examining how immobilization starting at pre-cavitation stages (E6-E10) could affect both growth and joint cavitation asymmetrically.

Firstly, we induced rigid paralysis in chicken embryos under conditions known to lead to a reduction in the length of hindlimbs after joint cavitation occurs^{13,24,25,26,27,28,29} and examined the length of skeletal elements, joint cavity formation and left–right patterning of forelimbs (Supplementary Fig. 3). The lengths of both the right and left humerus, ulna and metacarpus of digit 3 were significantly reduced in chicken embryos immobilized between E10-18 (Supplementary Fig. 3a, b). Pharmacologically induced immobility also led to drastic intraindividual variation in the length of the left and right humerus, ulna and metacarpus, compared to controls (Fig. 1e). These results suggest that mobility is necessary for symmetric limb patterning and growth.

Intriguingly, even upon immobilization at a post cavitation stage, two out of thirteen immobilized chickens showed an asymmetric pattern at the wrist joint with fusion of cartilaginous fusion of limb elements (Supplementary Fig. 3c). To understand further the effect of mechanical inputs on symmetrical growth and patterning of autopodial skeletal elements, we have immobilized chicken embryos from E6 (Supplementary Figs. 3d, 5 and 6), when muscle masses had developed, and forelimb movements were apparent (Supplementary Fig. 4a, b). The total length of skeletal elements of digits (metacarpus and phalanges) was significantly reduced compared to that in controls for all digits (Supplementary Fig. 5b). In addition, the reduction in digit length was relatively asymmetric in digits 2 and 4, while the reduction in digit 3 was relatively symmetric (Supplementary Fig. 5c). Of note, emu forelimbs exhibit severe reduction of digits 2 and 4 as well, but not of digit 3 (Fig. 1a; Supplementary Fig. 1a).

As previously described¹², immobilization of the chicken embryo at pre-cavitation stages led to a failure of proper joint cavity formation at the wrist region; we have observed that the cartilaginous fusion of skeletal elements was also asymmetric (Fig. 1f; Supplementary Fig. 6). Thinning or absence of the joint cavity at the wrist is one of the characteristic features of the emu forelimb skeletal pattern⁸ (Fig. 1a). Immobilization significantly reduced the distance between the ulna and the distal carpal in chicken embryos (Fig. 1f). Similar to immobilized chickens, the distance between the ulna and the metacarpus of digit 3 was also narrow in emu embryos at the same stage of development (Fig. 1f). These results indicate that loss of mobility from early stages recapitulated characteristic features of emu forelimbs, such as reduction or loss of the joint cavity at the wrist caused by asymmetric fusion of skeletal elements.

Taken together, these results reveal that the lack of distal muscle formation in emu embryos, when compared to chicken, led to a non-experimental immobilization phenotype that closely resembles experimental immobilization phenotypes in chicken. Specifically, we have shown that the reduction in bone length, increased susceptibility of the digits 2 and 4 to growth reduction when compared to digit 3, joint fusion and/or joint cavitation defects had an intraindividual asymmetric presentation in both emu and immobilized chicken forelimbs.

A population with dual myogenic and LPM cell signature is recognized in emu wings

Our findings suggest that the lack of distal muscle masses in comparison with chicken recapitulated, at least partially, the distinctive skeletal pattern of the emu wing. However, the underlying developmental reasons for the reduction of muscle in emu forelimbs are still not clear. To explore this, we first studied in detail the developmental process of forelimb muscles in emu embryos. In amniotes, muscle precursors delaminated from the ventral dermomyotome extensively migrate toward the distal part of the limb bud and differentiate into skeletal muscles. The delamination of muscle precursors in emu takes place in stages equivalent to chicken⁹; however, they migrate in a diffuse pattern and are not divided into dorsal and ventral muscle masses when compared to the emu hindlimb⁶ or to chicken forelimbs³³. The migratory muscle precursors³³ are characterized by the expression of Lbx1, which encodes a ladybird homeobox transcription factor³⁴, thought to be involved in controlling their migration^35,36,37,38. cMet, which encodes a tyrosine kinase receptor, is expressed in the ventral dermomyotome as well as in muscle precursors migrating toward limb buds^39,40,41. In addition, it has been previously reported that the emu forelimb bud co-opted Nkx2.5 expression⁷, and Nkx2.5-positive cells become skeletal muscle in forelimbs^7,9. Thus, we examined the expression pattern of Lbx1, cMet as well as Nkx2.5 in developing emu wing buds (Fig. 2; Supplementary Figs. 7 and 8). In chickens, the invasion of Lbx1-positive migratory muscle precursors into the forelimb bud became apparent at stage 18³³ (Supplementary Fig. 7a). In emu, transcripts of Lbx1 and cMet were enriched in the lateral region of the dermomyotome, and their distribution in the forelimb field became detectable at stage 18 (Fig. 2a). The expression of Nkx2.5 was seen only in the anterior edge of the forelimb field at the same stages (Fig. 2a). By stage 19, Lbx1/cMet-positive migratory muscle precursors invaded into the dorsal one half to one-third of the forelimb bud region, and Nkx2.5 expression was observed to the central region of the anterior three-quarters of limb buds (Fig. 2a, b). At stage 22, expression of Nkx2.5 was seen from the cervical to the forelimb level in the lateral plate mesoderm (Supplementary Fig. 7b). By stage 23, cMet expression was observed in migratory muscle precursors spread throughout the limb buds, and Nkx2.5 expression was seen in the ventral two-thirds of the forelimb bud (Fig. 2b). To observe gene expression in more detail, in situ hybridization using RNAscope probes was also used. It was found that cMet transcripts were sparsely distributed in proximal muscle precursors derived from the dermomyotome and were more densely distributed in the medial part of forelimb buds, where Nkx2.5 transcripts were observed, at stages 19 and 23 (Fig. 2c; Supplementary Fig. 8). At stage 25, the expression of Lbx1 and Nkx2.5 was observed in the proximal region and the subapical region, and the expression of MyoD, a key regulator of myogenic differentiation, overlapped with these cells at the subapical region (Fig. 2b). Co-localization of cMet and Nkx2.5 transcripts were confirmed by using RNAscope probes, and MyoD transcripts also co-localized at the subapical region (Fig. 2d; Supplementary Fig. 8). Subsequently, the expression of Nkx2.5 was detected in the skeletal muscles (Supplementary Fig. 7c), as previously reported^7,9. Our results and the results of others indicate that muscle progenitors of emu wings are positive for both Lbx1 and cMet, marker genes for somite-derived myogenic cells, as well as for Nkx2.5, a gene typically expressed in the vertebrate cardiac mesoderm⁴².

**Fig. 2: Expression of Lbx1, cMet, Nkx2.5 and MyoD in developing emu forelimb buds.**

While skeletal muscle cells and limb bud cells are derived from the paraxial mesoderm and the somatic layer of the lateral plate mesoderm (LPM), respectively, the cardiac mesoderm is derived from the splanchnic layer of the LPM. Thus, these results raised the possibility that the emu muscle progenitors possess atypical markers, including ones typically linked to the LPM identity. To investigate gene expression in an unbiased manner and at single cell resolution, we performed single-cell RNA sequencing (scRNA-seq) using the Chromium platform and paired-end Illumina next-generation sequencing of the dissociated cells from the emu trunk tissue at the forelimb level at stage 20/21 and the forelimb bud at stage 25. Cells obtained from stage 20/21 trunk tissues (n = 2696) and stage 25 forelimb buds (n = 7363) were used for analyses after quality control of datasets using the Seurat package to remove potential doublets and low-quality reads. Multidimensional reduction through tSNE clustering led to the unbiased identification of the muscle clusters, which contained cells enriched for markers of the somite-derived myogenic cells, such as Pax3⁴³, cMet, and Lbx1 as well as markers of skeletal muscle progenitors, such as TnnT3 (Troponin T3, Fast Skeletal Type), MyoD1 and Myog (myogenin) from stages 20/21 trunk tissues at the forelimb level (Fig. 3a, b; Supplementary Figs. 9 and 10; Supplementary Data 1). Interestingly, in this muscle cluster, 6.2% of cells expressing both Pax3 and Hand2, which is a general marker of the lateral plate mesodermal cells^44,45,46,47, were identified (21/339 cells; Fig. 3c). We also identified the muscle clusters expressing the myogenic markers from stage 25 forelimb buds (Fig. 3d, e; Supplementary Fig. 11; Supplementary Data 2). In the muscle cluster of the stage 25 forelimb buds (Fig. 3d, e), 19.2% of muscle progenitors positive for both Pax3 and Hand2 were identified (97/505 cells; Fig. 3f). In this Pax3 + /Hand2+ subpopulation of cells, significantly higher levels of expressions of other somatic LPM cell markers, such as Prrx1 and Tbx5^{48,49,50,51,52,53}, compared to the Pax3 + /Hand2- progenitors were detected (Fig. 3g; Supplementary Data 3). In addition, the expression of MyoD1 and Tnnt3 was detected in this Pax3 + /Hand2+ subpopulation (Fig. 3g). These results suggest that the Pax3 + /Hand2+ cells found during emu limb development have a dual transcriptional signature of the somite-derived myogenic cell and the LPM cell.

**Fig. 3: A subpopulation of muscle progenitors exhibits a dual somite-derived myogenic cell/LPM cell signature in emu forelimb buds.**

To determine whether such subpopulation of muscle cells with a dual transcriptional signature is a unique feature of emu forelimbs, we analyzed a publicly available single-cell RNA-seq dataset obtained from stage 24 and 27 chicken forelimb buds⁵⁴ and compared them with data from stage 25 emu forelimb buds. In stage 25 emu forelimb buds, we identified clusters of muscle cells enriched with LPM cell markers (Supplementary Fig. 12). In contrast, in stage 24 and 27 chicken forelimb buds, Hand2 expression was not observed in the extracted cluster of muscle cells, and the expression of other LPM markers were not enriched (Supplementary Fig. 13; Supplementary Data 4-5). This suggests that muscle progenitors with an LPM transcriptional signature are distinctive to emu forelimb buds.

Next, we used in situ hybridization chain reaction (HCR) technique to examine the distribution of the subpopulation of cells with the dual Pax3 + /Hand2+ identity in developing emu forelimb buds (Fig. 4; Supplementary Fig. 14). At stage 19, transcripts of Pax3 were detected in the dermomyotome and those of Hand2 were detected in the forelimb mesoderm (Fig. 4a; Supplementary Fig. 14a), but co-localization of Pax3 and Hand2 transcripts was not identified in the forelimb buds. By stage 23, although a few cells begin to exhibit a dual transcriptional signature, most Pax3+ somite-derived myogenic cells and Hand2+ forelimb mesenchyme can be distinguished from each other (Fig. 4b; Supplementary Fig. 14b). However, by stage 25, Pax3 transcripts were co-localized with Hand2 transcripts in a population of cells aggregated at the proximal region of forelimb buds (Fig. 4c, d; Supplementary Fig. 14c, d), and continued to be observed at the same region until stage 27 (Supplementary Fig. 15). This cell population contained extensive pyknotic nuclei when compared with the surrounding cells (Fig. 4c’, e; Supplementary Fig. 15). In contrast, in developing chicken forelimb buds, Pax3+ dermomyotome and muscle progenitors were clearly distinguishable from Hand2+ mesenchymal cells, and unlike in emu forelimb buds, cell aggregation was not observed in proximal region (Fig. 4f–h; Supplementary Fig. 16). The cell population with pyknotic nuclear feature in emu limb buds appeared to be entrapped at the proximal region by forming aggregates, and most importantly, some of their nuclei were condensed or fragmented (Fig. 4c, d; Supplementary Fig. 15). At this stage, MF20-positive muscle cells were recognized among Pax3+ migrating muscle progenitors; however, such a strong MF20 staining was either absent or only sparsely recognized in the aggregated cells (Fig. 4d, Supplementary Figs. 14c and 15). Taken together, in emu forelimb buds, a subpopulation of muscle progenitors exhibits a dual Pax3+ somite-derived myogenic cell/Hand2 + LPM cell signature and can be located forming aggregates at the proximal region during emu wing development.

**Fig. 4: A subpopulation of muscle progenitors co-expressing Pax3 and Hand2 aggregates at the proximal region of emu forelimb buds.**

A population of muscle progenitors with dual LPM/myogenic cell underwent cell death during wing development

Fragmentation of nuclei seen in the Pax3 + /Hand2+ cell population is a characteristic feature of apoptotic cells. In accordance with this prediction, scRNA-seq data showed that the expression of BCL2 Antagonist/Killer 1 (BAK1), encoding a pro-apoptotic protein, Caspase-10 (CASP10), encoding a cysteine peptidase responsible for the activation of the apoptotic cascade, and Apoptotic Peptidase Activating Factor 1 (APAF1), encoding a cytoplasmic protein that assembles into the apoptosome upon cytochrome c binding, were recognized in the Pax3 + /Hand2+ cell population, albeit not statistically upregulated at expression level (Fig. 5a; Supplementary Data 3). Of note, most cells actively undergoing apoptosis have been eliminated from our scRNA-seq during cell preparation steps (density gradient centrifugation excludes apoptotic bodies) or bioinformatics quality control (exclusion of cells with mtDNA content > 15%). Nevertheless, it remained unclear whether the cells exhibiting nuclear fragmentation were congruent with those identified through scRNA-seq analysis. Therefore, to investigate this further, apoptosis assays were conducted using TUNEL staining, along with immunostaining for cleaved caspase-3 and the oxidative damage marker 8-oxoguanine, on sections of emu forelimb buds (Fig. 5b–d; Supplementary Fig. 17). In this population, cells positive for TUNEL, caspase-3, and 8-oxoguanine were highly abundant (Fig. 5b–d). Additionally, some TUNEL-positive cells in the proximal region were surrounded by MF20-positive cells (Supplementary Fig. 17), suggesting that these dying cells were among muscle progenitors. These findings indicate that a distinct population of muscle progenitor cells, characterized by a dual LPM/myogenic cell signature, undergoes cell death.

**Fig. 5: A subpopulation of muscle progenitors aggregated at the proximal region of emu forelimb buds undergoes cell death.**

Discussion

Here, we show that emu forelimbs exhibit left–right asymmetric skeletal patterns, in addition to a reduction in longitudinal bone growth in the limbs and contractures of wrist joints. The observed morphological traits in emu forelimbs can be attributed to a reduction of mechanical loading from embryonic movements, which in turn is due to the absence of muscles at the distal parts of their forelimbs. Impaired skeletal muscles of emu forelimbs are, at least partially, attributed to massive cell death of a population of muscle progenitors exhibiting a dual LPM/myogenic cell signature.

During myogenesis, muscle progenitors/myoblasts take one of three distinct fates⁵⁵. While the majority of myoblasts fuse to form multinucleated muscle fibers/myotubes, there is a second population of cells located between the basal membrane and the sarcolemma of the muscle fibers that becomes myogenic stem cells/satellite cells. Furthermore, a third population of muscle progenitors that fails to differentiate into muscle fibers undergo programmed cell death⁵⁵. In emu forelimb buds, some of Pax3 + /Hand2- migrating muscle progenitors expressed high levels of myosin heavy chain (MF20), indicating differentiation into muscle fibers. On the other hand, MF20 signals were hardly detectable in a subpopulation of cells exhibiting a dual LPM/myogenic cell signature, suggesting that these cells might have failed to differentiate into muscle fibers and undergo other ‘default’ states, such as cell death. It is reasonable to assume that muscle progenitors with a LPM transcriptional signature may not be able to form muscle fibers which usually form by a fusion of cells in a homogenous population of myoblasts. Interestingly, the differentiation of myoblasts into muscle fibers requires activation of caspase-3⁵⁶, as well as cytochrome c and Apaf-1⁵⁷, suggesting that the mitochondrial death pathway can promote cell fusion as well as cell death. Caspase-dependent non-lethal differentiation has been reported not only in myoblasts, but in a variety of cell types^58,59, however, the mechanisms of how some cell types avoid cell death under the presence of caspase is largely unknown⁵⁷. The duration and intensity of caspase-3/7 activity is shown to be critical to determine whether mouse embryonic stem cells differentiate into cardiomyocytes or undergo cell death⁶⁰. In emu forelimb buds, a subpopulation of cells exhibiting a dual LPM/myogenic cell signature displayed a strong activity of caspase-3, along with oxidative damages of DNA and fragmented nuclei, suggesting caspase-dependent apoptosis occurred. Given that caspase-dependent apoptotic signals are required for differentiation of myoblast into muscle fibers, it is conceivable that cells with a dual LPM/myogenic cell signature fail to differentiate into muscle fibers due to excess level of caspase activity for differentiation, and consequentially, underwent cell death before migrating toward the distal region.

Multiple factors have been associated with the morphological features of emu forelimbs. In the early limb bud of emu embryos, the width of Grem1 expression domain, which indicates the amount of digit progenitors regulated by the SHH/GREM1/AER-FGF system^{61,62,63,64,65}, exhibits individual variation⁸. Recent work showed that regulatory changes lead to the lower expression of Fgf10 and a concomitant failure to express genes related to cell proliferation in the early emu forelimb⁹. Such alterations in gene expression, including a reduction in Fgf8 expression in the apical ectodermal ridge, might contribute to the individual variation in Grem1 expression seen in the early limb bud. The reduction in limb proliferation due to the lower expression of Fgf10 can explain the heterochrony observed in emu forelimb outgrowth^6,9. Loss of embryonic mobility also resulted in a late-stage reduction in the forelimb growth (Fig. 2b), as previously shown for hindlimbs of both chicken and crocodiles²⁹. Thus, the lack of movement of the distal part of the forelimb might also be a source of the heterochronic development found at later stages in emu forelimbs⁶ in regard to their skeletal development. Massive cell death of muscle progenitors is correlated with the reduction of muscle masses in the emu forelimb, which became evident at later stages (Fig. 1c) as well as in adult wings¹¹. The resulting decrease in mechanical stimulation, essential for osteoblast differentiation⁶⁶, likely results in a scarcity of specified skeletal progenitors. This scarcity contributes to a range of skeletal alteration, such as the reduction and fusion of skeletal elements, reflecting the skeletal abnormalities observed subsequent to the loss of mobility during embryonic and fetal stages¹². These data suggest an integrated model in which multiple mechanisms contribute to the extreme reduction and unique phenotype of the flightless emu wing.

We propose that the morphological features seen in emu forelimbs are, at least partially, attributed to a reduction in mechanical loading from embryonic movements, which is caused by the absence of distal muscles. Impaired development of forelimb muscles seems to be associated with the massive cell death of muscle progenitors with a dual LPM/myogenic cell signature. These progenitor cells accumulate at the proximal region, thereby losing their motility, and eventually undergoes cell death (Fig. 5e).

Mechanical loading from embryonic movements, which can be influenced by a range of environmental factors, has been suggested to contribute to the establishment of various skeletal design of limbs^{13,15,16,29,31}. The locomotory system displays a remarkable integration between its muscular and skeletal components, allowing for coordinated phenotypic changes throughout evolution. The interdependency of these systems can be identified at several organizational levels. For example, at a molecular level, the mechanotransducer and transcriptional regulator YAP regulates the development of both the skeleton^67,68 and muscle⁶⁹. At a cellular level, cells with dual muscle and LPM identity were identified during mouse limb development and play an important role in integrating the skeletal and muscle compartments at the myotendinous junctions^69,70,71. And finally, at organismal level, the “two-legged goat” example reveals how a change in behavior led to morphological traits that phenocopied several evolutionary novelties found in bipedal mammals, in a remarkable example of plasticity that is only possible when the development of muscular and skeletal compartments is strongly coordinated⁷².

Species-specific traits can arise not only from distinct load bearing on the adult skeleton, but also from differential motility affecting skeletal proportions and joint formation during development³¹. In a contrasting example of morphological evolution driven by a naturally occurring immobilization, the intrinsic muscles of jerboa feet disappear by a muscle atrophy mechanism without evidence of cell death⁷³. Our results demonstrate that differential embryonic muscle contractions, which can be altered both genetically and epigenetically, may underlie the morphological diversification that has occurred during vertebrate evolution, including in vestigial structures such as the emu wing.

Methods

Data reporting

No statistical methods were used to predetermine the sample size. The experiments were not randomized, and the investigators were not blinded to allocation during the experiments and outcome assessment.

Animals

White Leghorn chicken (Gallus gallus) eggs were incubated at 37.5 °C and staged⁷⁴. Fertilized emu (Dromaius novaehollandiae) eggs were purchased from Kakegawa Kachoen and Okhotsk Emu Pasture, incubated at 36 °C and staged as described in ref. ⁴. Wing samples of adult chickens were purchased post-mortem from a commercial supplier. Wing samples of adult emu were provided post-mortem from Tokyo Nodai Bioindustry Corporation. All animal work was performed in accordance with the guidelines for animal experiments of the Tokyo Institute of Technology, Kumamoto University, and The Jikei University School of Medicine, and the experimental protocols were approved by the committees of Tokyo Institute of Technology, and Kumamoto University. The sex of the animals was unknown.

Manipulation of embryo movement

Rigid paralysis²⁷ was induced in chicken embryos with decamethonium bromide (DMB), as previously described in ref. ²⁹ with slight modifications. Briefly, embryos were immobilized between either E10-17 or E6-10. On the first day, 100 μl of sterile filtered 5 mg/ml DMB in 100 μl phosphate-buffered saline (PBS) was injected through the egg window onto the chorioallantoic membrane to induce immobilization. On each subsequent day, 100 μl of 1 mg/ml DMB in PBS was administered to maintain paralysis. Embryos were monitored by viewing through the egg window on each day of treatment and any DMB-treated embryos that were not fully paralyzed were excluded from the study. Control animals were administered only vehcle (PBS). The embryos were euthanized one day after the treatment and fixed in 4% paraformaldehyde. To quantify the rate of distal movements in the wings of embryos (Supplementary Fig. 1d), we recorded the behavior of stage 39 chicken and emu embryos incubated at 37.5 °C and 36 °C, respectively, through the window of the eggshell. Counts were conducted only when the embryos exhibited active movement, defined as any instance where the head, arms or legs were actively moving for more than 10 s. The rate of distal movements per individual was calculated as the distal movement count divided by the proximal movement count in five minutes of total active movement time. Proximal movement corresponded to extension and flexion of the elbow joint; distal movement corresponded to any wrist or phalangeal movement (Supplementary Fig. 2b).

Probe synthesis and in situ hybridization

Total RNA was extracted from stage 25–26 chickens, and stage 25 emu embryos using RNeasy kit (Qiagen). cDNA was synthesized by reverse transcription and used as a template for PCR. The construct used to synthesize RNA probes were based on the pBlueScript II SK(+), using In-Fusion HD Cloning Kits (Clontech). The primers used to amplify most of the gene fragments were designed with the forward (5′-ATCGATAAGCTTGAT…−3′) and reverse (5′-CTGCAGGAATTCGAT…−3′) prefixes complementary to the ends of the linearized vector, which were followed by the specific sequences of the primers that were based on the indicated published sequences: chicken Lbx1 (Ensembl, ENSGALG00000034189), 5′-CTGCGCTTCAACTTTTGCTC-3′ and 5′-GGTTCTGGAACCAGGTGAT-3′; chicken MyoD (GenBank accession number, NM_204214), 5′-CTTCTATGACGACCCGTGC-3′ and 5′-GTCTTGGAGCTTGGCTGAA-3′; emu cMet (Ensembl, ENSDNVG00000012888), 5′-AGGAGCCATGGACAATGCAA-3′ and 5′-CGAATGGACCTCTTCCTC-3′; emu GlobinA (Ensembl, ENSDNVT00000001340.1), 5′-GAGCTGCAACCATG-3′ and 5′-TGGCTGCTCGCTG-3′. For emu Nkx2.5 (Ensembl, ENSDNVG00000013910.1), the open reading frame (ORF) of emu Nkx2.5 was amplified using specific primers (5′-CCCACCGCAATGTTTCCTAGCCCT-3′ and 5′-CTACCAGGCTCGGATCCCGTGCA-3′) and cloned into the EcoRV site of the pBlueScript SK(−) (pBSK-Nkx2.5). For emu Lbx1 (Ensembl, ENSDNVG00000014711), emu genomic DNA was used as a template for PCR. The 5′ and 3′ fragments of emu Lbx1 were amplified using the specific primers with the sequences complementary to the ends of the linearized pBlueScript II SK(+) (5′ fragment, 5′-ATCGATAAGCTTGATGTGGGCTTCAACTTTTGCTC-3′ and 5′-CTGCAGGAATTCGATGAGCCCCTTGAAGGTCTT-3′; 3′ fragment, 5′-ATCGATAAGCTTGATACCTATTCGTGGCCATGTCG-3′ and 5′-CTGCAGGAATTCGATCCGATGGGGCTGAGTGTAA-3′) and cloned into the vector using an In-Fusion HD Cloning Kit. The first exon sequence was obtained by fusing the DNA duplex synthesized using the oligonucleotides (5′-AGACCTTCAAGGGGCTCGAAGTGAGCGTGCTGCAGGCGGAATTCCTGCAGCCC-3′ and 5′-GGGCTGCAGGAATTCCGCCTGCAGCACGCTCACTTCGAGCCCCTTGAAGGTCT-3′) into the HindIII site (3′ side) of the 5′ fragment. The full-length of emu Lbx1 was obtained by cloning of the first exon sequence using specific primers with HindIII sequence (5′-GGTATCGATAAGCTTGATGTGGGCTTCAACTTTTG-3′ and 5′-GGAGACGCTGCGATTCGCCTGCAGCACGCT −3′), into the HindIII site of the vector including the second exon sequence, which was obtained by fusing the fragment amplified using the following primers by using In-Fusion HD Coning Kits (5′-GGTATCGATAAGCTTAATCGCAGCGTCTCCCCTGC-3′ and 5′-CGACATGGCC ACGAATAGGT-3′) to the 5′ side of the 3′ fragment.

For in situ hybridization, embryos were fixed overnight in 4% paraformaldehyde in phosphate-buffered saline, dehydrated in a graded methanol series and stored in 100% methanol at −20 °C. Whole-mount and section in situ hybridization were carried out as described^75,76. For RNAscope in situ hybridization, probes for emu cMet (targeting 827–1801 bp of XM_026097341.1), emu Nkx2.5 (targeting 2-1564 bp of XM_026122666.1), and emu MyoD (targeting 31–1358 bp of XM_026092794.1) were designed commercially by the manufacturer (Advanced Cell Diagnostics, Inc.). RNAscope in situ hybridization was performed using the RNAscope Fluorescent Multiplex Reagent Kit according to the manufacturer’s instructions. HCR in situ hybridization was conducted according to the manufacturer’s instructions. HCR probes targeting emu Pax3 (XM_026109761.1) and Hand2 (XM_026122869.1) were purchased commercially from Molecular Instruments, Inc. Images were obtained using an LSM 780 confocal microscope (Zeiss).

Histology

Whole-mount immunostaining was performed as previously described⁷⁷. Supernatant containing the monoclonal anti-MYH1E antibody was used to detect against myosin heavy chain (MF-20 hybridoma clone, DSHB). Goat anti-mouse IgG, HRP-conjugated antibodies (HAF007, R&D Systems) were used at a concentration of 1:200. Section immunostaining was performed as described in ref. ⁷⁸. Briefly, cryosections were incubated overnight with 1:1000 anti-cleaved caspase-8 antibody (9664, Cell signaling technology) or 1:100 anti-8-oxoguanine antibody (ab 64548, Abcam) at 4 °C. Sections were washed and incubated overnight with 1:1000 goat anti-rabbit IgG Alexa Fluor-488 conjugated antibody (ab150077, Abcam) or goat anti-mouse IgG Alexa Fluor-488 conjugated antibody (Invitrogen) at 4 °C and then washed. For TUNEL staining, cryosections were stained using TUNEL Mix (In situ Cell Death Kit, Roche) according to the manufacturer’s protocol. The sections were then incubated overnight with 1:400 sheep anti-fluorescein-AP antibody (Roche) at 4 °C and then washed. For Safranin-O staining, sections of embryos were prepared as described⁷⁹. Briefly, embryos were fixed with 4% paraformaldehyde, dehydrated in ethanol and acetone, and embedded in Technovit 8100 resin (Heraues-Kulzer, Wehrheim, Germany). Sections were cut at a thickness of 8 μm and stained with Weigert’s iron hematoxylin, Safranin-O and Fast Green as described in ref. ¹⁸. The narrowest parts of the interspace distance between the ulna and the distal carpus (chicken) or metacarpus (emu) were measured using cellSens software (Olympus).

Skeleton staining

Skeletons were stained according to standard protocols using Alcian blue for cartilage and Alizarin red for bone⁸⁰. The length of individual bones was measured using Fiji⁸¹.

Computed tomography imaging

Computed tomography (CT) imaging of adult emu forelimb skeletons was performed by a micro-CT system (Latheta LCT-200, Hitachi Aloka Medical Ltd., Tokyo, Japan). Acquired slice data were rendered as three-dimensional images using VGStudio MAX2.0 software (Volume Graphics GmbH., Heidelberg, Germany). Fusion of skeletal elements was evaluated by RadiAnt DICOM viewer (www.radiantviewer.com/).

Single-cell RNA sequencing and analysis

Trunk tissue at the forelimb level from four stage 20/21 emu embryos, and from fifteen forelimb buds from stage 25 emu embryos, were collected, pooled and then dissociated into single cells by enzymatic digestion as previously described⁸², with modifications. Briefly, limb buds were treated with 2000 U dispase II (Wako) in Ca²⁺- and Mg²⁺-free Tyrode’s solution (CMF-Tyrode) at 4 °C for 50 min and transferred to Dulbecco’s modified Eagle’s medium (DMEM) containing 1% fetal bovine serum (FBS), when their ectodermal sheets were removed by peeling off with a pair of forceps. Then, the remaining limb tissues or trunk tissues were transferred to Cellbanker 1 plus (Nippon Zenyaku Kogyo) and stored at −80 °C. Thawed tissues were washed in DMEM containing 0.04% bovine serum albumin (BSA), and incubated in CMF-Tyrode at 37 °C for 40 min. Softened tissues were dissociated into a single cell suspension by pipetting in DMEM containing 0.04% BSA. The suspension was first filtered using pluriStrainer-Mini 100 μm (pluriSelect), and the debris was removed using Debris Removal Solution (Miltenyi Biotec) according to the manufacturer’s instructions. Then, cells were filtered again using pluriStrainer-Mini 40 μm (pluriSelect), suspended in DMEM containing 2% BSA and proceeded immediately to cell encapsulation, indexing and transcriptome library prepration, as described below.

The single-cell transcriptome data of emu embryos were generated at the Functional Genomics Facility of the National Institute for Basic Biology (NIBB) in Okazaki, Japan. For scRNAseq library construction, barcoded single-cell cDNA libraries were synthesized using 10x Genomics Chromium Single Cell 3’ Reagent Kits v3.1 (Dual Index) according to the manufacturer’s instructions. Libraries were sequenced on an Illumina Hiseq platform at a depth of 117,418 and 48,543 mean reads per cell for stage 20/21 and stage 25 samples, respectively. The single-cell transcriptome data of chicken forelimbs were published previously⁵⁴. Chicken forelimb single cell RNA sequence data of stage 24 (SRA accession numbers: SRR14570167 to SRR14570174) and stage 27 (SRA accession numbers: SRR14570175 to SRR14570178) were downloaded from the Sequence Read Archive (SRA).

Raw sequencing data were processedand aligned with a emu reference genome assembly droNov1 (GCF_003342905.1) or a chicken reference genome assembly GRCg6a (Ensembl release 105) using Cell Ranger software version 7.1.0 (10x Genomics). Cell Ranger filtered outputs files for each dataset were processed using R package Seurat v4.3.0⁸³. Following the standard pre-processing workflow of Seurat, low-quality cells and potential cell doublets were removed from the dataset. Gene expression was log normalized with NormalizedData and scaled with ScaleData using all genes. PCA-reduction was performed with RunPCA and nearest neighbor graph was constructed using the 50 PCs with FindNeighbors, and then clusters were identified using FindClusters. Cluster were assigned specific identities based on differentially expressed genes. To visualize these datasets, tSNE (t-distributed Stochastic Neighbor Embedding) was performed with RunTSNE using 1:50 dims as input. Differentially expressed genes for each cluster of emu and chicken data were provided by using the FindAllMarkers, which performs Wilcoxon rank-sum tests. Differential expression testing between Pax3 + /Hand2+ cells and Pax3 + /Hand2- cells from the muscle cluster of stage 25 emu was conducted using the FindMakers function, which applies Wilcoxon rank-sum tests.

Statistical analysis

All statistical analyses, except for Figs. 3g and 5a, were performed with Prism 8.0.2 (GraphPad). n numbers represent the numbers of limbs, and measurements are represented as the mean ± SEM. All measurements were taken from distinct samples. Two-parameter comparisons between measurements of biological samples assumed unequal variances, thus Welch’s unpaired t-test was used. The coefficient of variation was calculated as the ratio of the standard deviation to the mean. Statistical details can be found in the legends. Significance was defined at P ≤ 0.05 (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). For differentially expressed genes in Figs. 3g and 5a, the two-sided Wilcoxon rank-sum tests were used for statistical tests (p-val adjusted < 0.05).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The authors declare that all data supporting the findings of this study are available within the article and its Supplementary Information files. The raw sequencing data generated in this study have been deposited in the DDBJ Sequence Read Archive under accession numbers: DRA017391 and DRA014432; bioproject: PRJDB13845 and PRJDB16987. The publicly available chicken datasets, published before⁵⁴, were retrieved from Sequence Read Archive (SRA) under the accession numbers SRR14570167 to SRR14570174 (https://www.ncbi.nlm.nih.gov/sra/SRX10913414[accn]) and SRR14570175 to SRR14570178 (https://www.ncbi.nlm.nih.gov/sra/SRX10913415[accn]). Source data are provided with this paper.

Code availability

All code used to reproduce the Supplementary Fig. presented in this paper is publicly available through GiHub. The corresponding DOI is https://doi.org/10.5281/zenodo.11127881.

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Acknowledgements

We thank Dr. Q. Zhou for providing wing samples of emu embryos, Drs. J. Miyazaki, T. Ogura and A. P. McMahon for plasmids, Tokyo Nodai Bioindustry Corporation for providing wing samples of adult emu, Kakegawa Kachoen and Okhotsk Emu Pasture for providing emu eggs, Dr. T. Hirasawa for valuable comments, Mr. A. Irifune and Dr. M. Uesaka for technical assistance, and Biotechnology Center of Tokyo Institute of Technology for sequencing services. This work was supported by JSPS KAKENHI Grant Numbers JP20H03301, and JP17KT0106, MEXT KAKENHI Grant Number JP18H04818, NIBB Collaborative Research Program (21-357), Astellas Foundation for Research on Metabolic Disorders, Mitsubishi Foundation and Yamada Science Foundation to M.T.

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Ingrid Rosenburg Cordeiro
Present address: Department of Cell and Molecular Biology, Karolinska Institutet, Stockholm, Sweden
These authors contributed equally: Eriko Tsuboi, Satomi F. Ono, Ingrid Rosenburg Cordeiro.

Authors and Affiliations

School of Life Science and Technology, Tokyo Institute of Technology, B-17, 4259 Nagatsuta-cho, Midori-ku, Yokohama, Kanagawa, Japan
Eriko Tsuboi, Satomi F. Ono, Ingrid Rosenburg Cordeiro, Reiko Yu, Toru Kawanishi & Mikiko Tanaka
Laboratory Animal Facilities, The Jikei University School of Medicine, 3-25-8 Nishi-shimbashi, Minato-ku, Tokyo, Japan
Makoto Koizumi
Trans-Omics Facility, National Institute for Basic Biology, Nishigonaka 38, Myodaiji, Okazaki, Aichi, Japan
Shuji Shigenobu
International Research Center for Medical Sciences, Kumamoto University, 2-2-1 Honjo, Chuo-ku, Kumamoto, Japan
Guojun Sheng
Department of Anatomy, The Jikei University School of Medicine, 3-25-8 Nishi-shimbashi, Minato-ku, Tokyo, Japan
Masataka Okabe

Authors

Eriko Tsuboi
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Satomi F. Ono
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Ingrid Rosenburg Cordeiro
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Reiko Yu
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Toru Kawanishi
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Makoto Koizumi
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Shuji Shigenobu
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Guojun Sheng
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Masataka Okabe
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Mikiko Tanaka
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Contributions

E.T., I.R.C., S.F.O. and M.T. designed the project and wrote the manuscript. E.T. analyzed the length and patterns of skeletal elements and muscles, I.R.C. examined gene expression, S.F.O. examined the movement of embryos, and performed scRNA-seq analyses, S.F.O. and T.K. captured confocal images, M.T. performed histology, immunology and examined gene expression by using DIG-, RNA scope- and HCR-probes, R.Y. constructed plasmids, S.S. provided sequencing research infrastructures, G.S. provided plasmids related to blood differentiation and emu embryos. M.K. took CT images, M.O. assisted with CT scanning.

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Correspondence to Mikiko Tanaka.

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Nature Communications thanks Sevan Hopyan and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Tsuboi, E., Ono, S.F., Cordeiro, I.R. et al. Immobilization secondary to cell death of muscle precursors with a dual transcriptional signature contributes to the emu wing skeletal pattern. Nat Commun 15, 8153 (2024). https://doi.org/10.1038/s41467-024-52203-x

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Received: 13 November 2023
Accepted: 29 August 2024
Published: 19 September 2024
DOI: https://doi.org/10.1038/s41467-024-52203-x
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Developmental insights from non-traditional model organisms

Collection 08 February 2024

Immobilization secondary to cell death of muscle precursors with a dual transcriptional signature contributes to the emu wing skeletal pattern

Abstract

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Introduction

Results

High intraindividual left-right variation is found in the adult skeletal pattern of the distal emu wings

Developing emu forelimbs receive less mechanical input due to the absence of distal muscle masses

Loss of mobility recapitulates characteristic features of emu forelimbs

A population with dual myogenic and LPM cell signature is recognized in emu wings

A population of muscle progenitors with dual LPM/myogenic cell underwent cell death during wing development

Discussion

Methods

Data reporting

Animals

Manipulation of embryo movement

Probe synthesis and in situ hybridization

Histology

Skeleton staining

Computed tomography imaging

Single-cell RNA sequencing and analysis

Statistical analysis

Reporting summary

Data availability

Code availability

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Acknowledgements

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