Neural signaling contributes to heart formation and growth in the invertebrate chordate, Ciona robusta

Neural signaling contributes to heart formation and growth in the invertebrate chordate, Ciona robusta | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Neural signaling contributes to heart formation and growth in the invertebrate chordate, Ciona robusta Hannah N. Gruner , C. J. Pickett , Jasmine Yimeng Bao , Richard Garcia , Akiko Hozumi , Tal Scully , Shaoyang Ning , Mavis Gao , Gia Bautista , Keren Maze , Karissa Lim , Tomohiro Osugi , Mae Collins-Doijode , Ofubofu Cairns , Gabriel Levis , Shu Yi Chen , TaiXi Gong , Honoo Satake , Allon Moshe-Klein , Eduardo D. Gigante , Yasunori Sasakura , Bradley Davidson doi: https://doi.org/10.1101/2025.04.28.651085 Hannah N. Gruner 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site C. J. Pickett 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jasmine Yimeng Bao 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Richard Garcia 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Akiko Hozumi 2 Shimoda Marine Research Center, University of Tsukuba , 5-10-1, Shimoda, Shizuoka, 415-0025, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tal Scully 3 Harvard Medical School , Warren Alpert Building Room 536, 200 Longwood Ave Boston, MA 2114, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Shaoyang Ning 4 Swarthmore College, Statistics in the Department of Mathematics & Statistics at Swarthmore College , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Mavis Gao 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Gia Bautista 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Keren Maze 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Karissa Lim 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tomohiro Osugi 5 The Bioorganic Research Institute, Suntory Foundation for Life Sciences (SUNBOR) , 8-1-1 Seikadai, Seika-cho, Soraku-gun, Kyoto, 619-0284, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Mae Collins-Doijode 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Ofubofu Cairns 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Gabriel Levis 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Shu Yi Chen 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site TaiXi Gong 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Honoo Satake 5 The Bioorganic Research Institute, Suntory Foundation for Life Sciences (SUNBOR) , 8-1-1 Seikadai, Seika-cho, Soraku-gun, Kyoto, 619-0284, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Allon Moshe-Klein 3 Harvard Medical School , Warren Alpert Building Room 536, 200 Longwood Ave Boston, MA 2114, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Eduardo D. Gigante 6 School of Biological Sciences, College of Sciences, Georgia Institute of Technology , Atlanta, GA 30332, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Yasunori Sasakura 2 Shimoda Marine Research Center, University of Tsukuba , 5-10-1, Shimoda, Shizuoka, 415-0025, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Bradley Davidson 1 Swarthmore College, Department of Biology , 500 College Ave, Swarthmore, PA 19081, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: bdavids1{at}swarthmore.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Neurons contribute to the complex interplay of signals that mediate heart development and homeostasis. Although a limited set of studies suggest that neuronal peptides impact vertebrate heart growth, the specific contributions of these peptides to cardiomyocyte progenitor differentiation or proliferation have not been elucidated. Here we show that the neuropeptide tachykinin along with canonical Wnt signaling regulate cardiomyocyte progenitor proliferation in the chordate model Ciona robusta . In C. robusta, the heart continues to grow throughout adulthood and classic histological studies indicate that a line of undifferentiated cells may serve as a reserve progenitor lineage. We found that this line of cardiomyocyte progenitors consists of distinct distal and midline populations. Distal progenitors divide asymmetrically to produce distal and midline daughters. Midline progenitors divide asymmetrically to produce myocardial precursors. Through single cell RNA sequencing (scRNA-seq) of adult C. robusta hearts, we delineated the cardiomyocyte progenitor expression profile. Based on this data we investigated the role of Wnt signaling in cardiomyocyte progenitor proliferation and found that canonical Wnt signaling is required to suppress excessive progenitor proliferation. The scRNA-seq data also identified a number of presumptive cardiac neural-like cells. Strikingly, we found that a subset of these neuronal cells appears to innervate the distal cardiomyocyte progenitors. Based on the expression of the tachykinin receptor in these neuronal cells, we blocked tachykinin signaling using pharmacological inhibitors and found that this drove reduced proliferation in the distal progenitor pool. Through targeted CRISPR-Cas9 knockdown we then demonstrated that both extrinsic tachykinin and intrinsic, cardiac tachykinin receptors are required for formation of the myocardial heart tube. This work provides valuable insights into how organisms may deploy neural signals to regulate organ growth in response to environmental or homeostatic inputs. Introduction Neural signaling and development Neurons contribute to the complex interplay of signals that dictate cellular behaviours during development. Along with their key role in neural development and patterning, neuronal signals regulate non-neuronal cell behaviours. The best studied instances of such heterotypic interactions focus on cell behaviours associated with homeostasis, cancer, or regeneration ( 1 – 3 ). In comparison, relatively few studies have documented heterotypic impacts of neural signals on cell fate patterning, morphogenesis or early growth. The first such studies revealed a role for innervation in skeletal muscle differentiation ( 4 ). Critically, these studies indicated that the contribution of neural signals was not entirely dependent on muscle activity ( 5 – 10 ). More recent studies have buttressed these findings, suggesting that signals from both the peripheral and central nervous system inform skeletal muscle patterning ( 4 , 11 ). Additional research has extended the range of morphogenetic processes impacted by neuronal signals including melanocyte differentiation ( 12 ), blood vessel patterning ( 13 ), intestinal progenitor differentiation( 14 ), salivary gland development ( 15 , 16 ), hematopoietic stem cell specification ( 17 – 19 ), pancreatic islet formation ( 20 ), and the proliferation of hippocampal cells ( 21 ). Subsequent studies have also found that both neurotransmitters and neuropeptides can regulate these developmental cell behaviours. Additionally, a limited set of studies suggest that neuronal signals may regulate vertebrate heart growth ( 22 ). Vertebrate heart growth Vertebrate heart growth has been most intensively studied in mammals, particularly in mice ( 23 – 25 ). Human and mouse embryonic hearts exhibit extremely high levels of growth during cardiogenic patterning stages ( 26 , 27 ), peaking during the period of chamber expansion when heart volume increases by over 100-fold ( 28 – 30 ). The period of extensive growth also continues after morphogenesis is complete. Throughout murine embryonic and fetal stages there are similar increases in myocardial volume and cardiomyocyte numbers indicating that heart growth is largely driven by proliferation ( 28 ). Recent studies, including extensive scRNA-seq efforts, have revealed unanticipated levels of heterogeneity within early progenitor populations and mapped some of these distinct progenitors within the heart field ( 31 – 37 ). Studies that encompass later stages of cardiogenic patterning have shown that early progenitor pools persist at the venous and arterial poles where they contribute to heart tube elongation ( 26 , 32 , 33 , 36 – 45 ). Signals that regulate vertebrate heart growth Extensive research has documented signal-dependent coordination of initial cardiogenic patterning along with homeostasis and regeneration of the mature organ ( 46 – 54 ). In contrast, relatively few studies have examined signal-dependent regulation of heart growth during the intermediate stage following completion of morphogenesis ( 23 – 25 ). Numerous paracrine signaling pathways coordinate vertebrate heart growth by modulating either cardiomyocyte specification or proliferation. These include the Fibroblast Growth Factor (FGF), and Insulin Growth Factor (IGF), Hedgehog, Wingless-related integration site (Wnt), bone morphogenetic proteins (BMP), Retinoic Acid (RA), Hippo and Notch pathways along with microenvironmental cues associated with the extracellular matrix (ECM) such as hyaluronic acid ( 48 , 52 , 55 – 66 ). Some of these pathways, such as Wnt and FGF, have extensive, cascading roles in myocardial growth that have been difficult to disentangle ( 48 , 60 , 63 , 67 , 68 ). Recent scRNA sequencing studies have delineated precise spatiotemporal expression patterns of receptors and ligands that are potentially involved in myocardial growth ( 34 ). Predicted receptor/ligand interactions from these studies align with previous functional data for a number of signals including IGF, BMP, and Notch while also revealing new candidate pathways ( 31 , 32 , 34 ). However, these studies have not focused on the potential contribution of neural signals to heart growth. The impact of neuronal signals on vertebrate heart development and growth The vertebrate heart is innervated by extrinsic neurons from the central nervous system along with intrinsic cardiac neurons. Cardiac intrinsic plexuses regulate heart rate and rhythm largely through modulation of the cardiac conduction system. The cardiac conduction system is composed of specialized cardiomyocytes that function as slow-conducting pacemaker cells in the sinoatrial and atrioventricular nodes along with fast conducting cells in structures such as the Bundle of His and Purkinje fiber network ( 69 , 70 ). The vagus nerve is the primary source of extrinsic cardiac innervation and plays a well-defined role in modulating intrinsic plexuses and cardiac conduction ( 71 , 72 ). In mouse embryos, extensions of the vagal nerve first innervate the embryonic heart during a period of dramatic proliferative growth following completion of morphogenesis ( 73 – 75 ). A number of studies indicate that innervation promotes cardiomyocyte maturation ( 76 , 77 ). However, studies regarding the potential role for neuronal signals in vertebrate cardiomyocyte proliferation have largely focused on regeneration or homeostasis ( 1 , 22 , 78 , 79 ). A more limited set of studies have investigated the impact of both neurotransmitters and neuropeptides on cardiomyocyte proliferation during earlier developmental stages. Two of these studies suggest that β-adrenergic signaling regulates the transition from proliferative to hyperplastic growth ( 80 , 81 ). Another study indicated that vagus nerve neuropeptide secretion promotes cardiomyocyte proliferation in neonatal mice ( 22 ). Currently, it is unclear what specific signals and downstream pathways may mediate neural-dependent regulation of cardiomyocyte proliferation during these developmental periods. Studying neural-dependent heart growth in Ciona robusta The tunicate Ciona robusta represents a powerful model to explore the role of neural inputs in heart morphogenesis or growth. Tunicates are chordates and represent the closest extant invertebrate taxa to the vertebrates. This evolutionary relationship is most evident in the tunicate larval tail which exhibits two key chordate traits, a notochord and dorsal nerve cord. However, both of these features are lost during metamorphosis as the larval tail is resorbed. In the resulting juveniles, the central nervous system is restructured and rudiments contained in the larval head/trunk region undergo morphogenesis to form adult organs, including the heart. Despite this close evolutionary relationship, tunicate embryos, including those of C. robusta , have extremely low cell numbers and highly stereotyped cell lineages facilitating high resolution analysis of cell fate specification and morphogenesis. This cellular simplicity has been leveraged to gain profound insights into early stages of C. robusta heart development, along with comprehensive mapping of the larval nervous system and in-depth analysis of the neural circuits driving larval swimming behaviors ( 82 – 84 ). Efforts to analyze neural and cardiac development during C. robusta embryogenesis have also been facilitated by the small size of the C. robusta genome and an associated lack of genetic redundancy. Short intergenic regions permit rapid characterization of lineage specific regulatory elements and the lack of paralogs for transcription and signaling factors greatly simplifies analysis of developmental gene regulatory networks. Recently, C. robusta ’s extreme cellular and genetic simplicity were exploited to permit comprehensive single-cell RNA sequencing (scRNA-seq) of embryogenesis and reconstruction of the transcriptional trajectories for every cell lineage ( 85 ). However, studies of C. robusta development have largely focused on embryogenesis. Thus, post-larval development, including morphogenesis and subsequent growth of the adult heart or nervous system, remain poorly characterized. Cellular composition of the C. robusta heart The C. robusta heart is primarily constructed from two single-cell layers, the myocardium, and pericardium ( Fig 1A ) ( 86 ). The myocardium is a V-shaped tube consisting of myoepithelial cells capable of generating peristaltic bi-directional contractions. Embedded within one side of the myocardium is the undifferentiated line (UL). The UL is a tightly packed group of small round cells arranged into rows spanning the entirety of the myocardial tube ( 87 ). The classification of these cells as undifferentiated is based solely on appearance and their function has not been previously characterized. The pericardium forms a coelom around the myocardial tube and is connected to the myocardium on the opposite side of the heart from the UL by a structure termed the raphe. Superficial to the pericardium is an epithelial lining termed the “epicardium” which surrounds the stomach, intestine, gonad, and heart ( 88 ). Our current understanding of C. robusta heart anatomy is largely based on classic histological studies that have not been updated in over 50 years. Download figure Open in new tab Figure 1. Microscopic anatomy of the adult C. robusta heart. ( A ) Diagram depicting gross anatomy of the adult C. robusta heart. ( B ) Confocal micrograph of an adult C. robusta heart. ( C- F ) Cardiac cell types as labeled. ( G ) Cross section of the adult midline UL. ( H ) Distal end of an adult heart showing expanded UL along with adjacent pericardium and myocardium. Panel below shows a cross-section of H. ( I ) Diagram of the distal portion of the heart ( H ) highlighting a presumptive transit amplifying zone ( J ) composed of myocardial progenitors on one side of the expanded UL. ( J ) Streams of presumptive myocardial progenitors adjacent to the more densely packed cells in the UL region, to the left of the dotted line. In all images, cyan represents DAPI staining, magenta represents phalloidin staining and yellow represents WGA, which appears to preferentially label pericardial cells along with occasional labeling of UL or myocardial cells (see panels G and H ). Intrinsic innervation of the C. robusta heart As initially characterized by Kriebel et al. ( 89 – 92 ), tunicates have pacemakers at either end of the myocardial tube that regulate bi-directional pumping of the heart. Classic studies indicate that electrical coupling across the myocardial tube is facilitated by tight junctions between adjacent myocardial cells without a preferred conduction pathway or direction ( 90 – 92 ). Although some classic studies mention extrinsic neural innervation of the heart, more recent analyses have not detected these extrinsic neurons ( 93 – 95 ). A recent study identified rings of neural-like cells that are integrated within the distal ends of the myocardium ( 96 ). These cells were visualized using a stable transgenic line in which the Prohormone Convertase 2 ( PC2 ) gene regulatory element is used to drive expression of a Kaede fluorescent reporter. PC2 plays a highly conserved role in neuropeptide processing and thus expression of PC2 reporter suggests that these distal neural rings are peptidergic. Higher magnification imaging revealed that these distal rings consist of a plexus of interconnected bi-polar cells. This study also indicated that the distal rings are connected by a strand of peptidergic neurons running along the raphe. They also pointed out that the position and morphology of these distal plexuses are reminiscent of the intrinsic sinoatrial and atrioventricular plexuses present in vertebrate hearts. The authors of this study proposed that these distal plexuses function as pacemakers. However, the function and development of these distal peptidergic rings has not been characterized and no connections to extrinsic neurons originating outside of the heart were recorded. In this paper we delineate the microanatomical composition of the C. robusta heart and characterize the cardiac progenitors that drive proliferative heart growth. We have also begun to explore the signaling pathways that regulate heart growth, including a key role for canonical Wnt (cWnt) signaling in suppressing progenitor proliferation. Strikingly, we identified a set of neural-like cells that are closely associated with cardiac progenitors. These presumptive intrinsic cardiac neurons or pacemakers appear to be innervated by extensions from the CNS during early juvenile stages. Single cell-RNA sequencing of the adult heart indicated that these cardiac neural–like cells express a receptor for tachykinin-family neuropeptides (TACR). To begin assessing the role of tachykinin signaling in these cells, we applied a pharmacological inhibitor to the adult heart. Surprisingly this inhibitor had no impact on heart rate but rather suppressed heart cell proliferation. Transient inhibition of tachykinin signaling in juveniles specifically decreased proliferation in the distal progenitor lineages adjacent to intrinsic neurons. Targeted CRISPR knockdown of either TACR in cardiac lineage cells or tachykinin in extrinsic neurons led to nearly complete abrogation of the myocardium. Taken together, these results indicate that tachykinin -dependent stimulation of cardiac neural-like cells is required for cardiomyocyte progenitor proliferation. Results Cellular composition and growth of the adult C. robusta heart To lay the groundwork for exploring tunicate heart growth, we generated a high-resolution microanatomical whole-mount confocal map of the adult C. robusta heart ( Fig 1B ). Each of the four cardiac cell types (undifferentiated line, myocardium, pericardium, and epicardium) were readily distinguished based on their morphology and position. ( Fig 1C - 1F , S1A and S1B ). The undifferentiated line consists of small, tightly packed cells ( Fig 1F ). The myocardium is composed of an organized collection of myofibers radiating out from the UL ( Fig 1E , S1A and 1B ). The pericardium is immediately superficial to the myocardium and consists of flattened, roughly pentagonal cells with peripheral nuclei ( Fig 1D ). The epicardial cells, distinguishable by their more irregular shape and central nuclei, form a thin layer superficial to the pericardium ( Fig 1C ). Similar cell morphologies were observed in the hearts of a related species, Ciona savignyi ( S1C-S1F Fig ). We were particularly interested in the microanatomy of the undifferentiated line (UL). In the mid-section of the heart, the UL consisted of 3-5 irregular rows of compact cells. In cross-section, these rows form a semi-cylinder that is attached to myocardial cells on either side ( Fig 1G , S1A and S1B ). As observed previously by Millar, the UL expands at both the dorsal “visceral” and ventral “hypobranchial” ends of the heart to form larger clusters ( Fig 1H ) ( 87 ). Extensive folding made it extremely difficult to visualize the morphology of these distal UL clusters in intact hearts. To overcome this obstacle, distal regions of the heart were removed and dissected to form a flat sheet ( Fig 1H - 1J ). The observed distal UL clusters were highly variable in size and morphology. As seen in the mid-section of the heart ( Fig 1G ), the central cells of the distal UL clusters form a raised semi-cylinder ( Fig 1H , cross-section). In some cases, the distal UL appeared to display a density gradient, with an extremely dense region at the distal-tip and a wider, less dense proximal region (see Fig 1H ). In this wider proximal region, the UL cluster appeared to be flanked by different cell types on each side ( Fig 1H and 1I ). One side of the UL cluster appears to merge into a field of pericardial cells, as distinguished by their flat, hexagonal shape and enriched WGA staining. On the opposite side, streams of distal UL cells appear to penetrate into regions containing mature myocardial cells, as distinguished by their spindle-like morphology and dense bands of actin. There appears to be a morphological gradient between the distal UL cluster and mature pericardium as tightly-packed UL cells with highly-condensed nuclei transition into relatively sparsely distributed pericardial cells with larger-nuclei ( Fig 1H and 1I ). There also appears to be a morphological gradient associated with myocardial differentiation as penetrating streams of UL cells give way to a zone of less densely packed cells with larger nuclei and lower levels of actin enrichment ( Fig 1H - 1J ). These transitional cells may constitute a population of myocardial precursors. We next investigated the pattern of growth in adult C. robusta hearts. As posited by Millar, distal UL clusters may serve as proliferative growth zones for heart tube elongation ( 87 ). To test this hypothesis, we assayed gross proliferation rates in distal vs. middle regions of the myocardial tube (including both myocardial and UL cells) using 24-hour EdU pulses. As predicted by the distal growth zone hypothesis, the distal myocardium displayed a significantly higher proportion of mitotic cells in comparison to medial or apical regions ( S2A Fig ). We used a short 6-hour EdU pulse assay followed by a 24-hour chase to further characterize the location of mitotically active cells ( S2B and S2C Fig ). In distal regions, EdU+ cells were largely observed within or bordering the distal UL cluster ( S2B Fig ). By contrast, EdU+ cells were rarely detected in the midline UL ( S2C Fig ). Additionally, scattered EdU+ cells were detected throughout the myocardium. The proliferative distal UL border cells may represent a transit amplifying zone, in which myocardial or pericardial precursors undergo proliferation prior to differentiation (as illustrated in Fig 1I ). Further analysis of adult heart growth was limited by the convoluted morphology of the distal cluster and difficulties in obtaining intact, dissected samples. To overcome these challenges, we began to examine heart growth during early juvenile stages. Cellular composition of the juvenile heart Due to extremely low cell numbers, distinct cellular components of the juvenile heart can be readily distinguished ( Fig 2A and 2B ). The heart first starts beating three days after fertilization (D3), approximately two days after larval settlement. Although early-stage juvenile hearts (D3-D6) are composed of less than 100 cells, they are fundamentally similar in structure to the adult organ ( Fig 2A and 2B ). An inner, cylindrical myocardial tube is surrounded by two roughly spherical pericardium/epicardial layers composed of ∼70 flattened epithelial cells. The myocardial tube is composed of ∼10-24 cardiomyocytes attached along the posterior side to a UL composed of only ∼8-14 cells. As in the adult, the UL consists of a narrow medial line abutted by expanded distal clusters. However, in the juvenile the medial line is a single row and the distal clusters are composed of only two or three cells. Each medial UL cell appears to be associated with a ring-like pair of spindle-shaped myocardial cells ( Fig 2A and 2B ). As juveniles age, the UL gradually lengthens ( Fig 2C ). Download figure Open in new tab Figure 2. Juvenile heart anatomy and cell division dynamics. (A) D5 juvenile heart. (B) Schematic of ( A ). (C) Representative images of D8 (i), D11 (ii), and D15 (iii) juvenile hearts. Yellow dots highlight midline UL nuclei (C’, 13 midline UL; C’’, 15 midline UL; C’’’, 18 midline UL). (D-G) D5 juvenile hearts showing distinct EdU clones containing at least one UL as described in the text along with schematics coloured according to the model in panel B . (H) Representative micrograph and schematic of a juvenile heart showing two pairs of presumptive myocardial precursors at different distances from the UL. (I) Model for UL and myocardial precursor cell division in juvenile hearts, as tracked in a D5-D12 juvenile through photoconversion of Kaede-labeled ventral UL cells as shown in panel ( J ). At the start of the model (first panel, D5) asymmetric division of a distal UL cell in the ventral cluster has produced one distal UL daughter (grey, V0) and one midline UL daughter (dark blue, MUV0). Subsequently (second panel, D6) symmetric division of the newly produced midline UL (MUV0) produces two midline daughters (MUV0 shown in red and MUV1 shown in blue). Over the next few days, both of these midline daughters divide. One (MUV1, blue) divides symmetrically to produce two daughters that remain in the midline (MUV1A and B). The other (MUV0, red) divides asymmetrically to produce one midline UL daughter (MUV0A, red) and a myocardial precursor that leaves the midline (MUV0B). It is not clear from our data whether myocardial precursors are derived from the posterior or anterior daughter. According to this model, each myocardial precursor divides once as it migrates anteriorly, contributing two differentiated cells to the myocardial tube (D10-12 in the model). (J) Kaede conversions of UL cells. Cells were photoconverted on D5 and imaged daily over four days. See text for details of division patterns and cell movements of labeled cells (as illustrated in panel I ). Different collared arrows track three individual photo-converted cells over 96 hours after conversion, as indicated by the time point (T0-T96). Magnified regions shown in the upper left corner of each image. The red and blue arrows correspond to the red and blue cells in the model shown in panel I . Photoconverted cells are yellow due to photoconversion. Green cells are unconverted Kaede-green. For T-96, three different sub-stacks of the same Z-stack are shown to capture cells that reside in three different focal planes, upper UL, lower UL and myocardial as labeled. Juvenile heart growth To begin investigating juvenile heart growth we quantified the UL and myocyte numbers during early stages (D3-D15). UL cells and myocytes were readily identified based on their morphology and position ( Fig 2A - 2C ). Under optimized growth conditions, our analysis indicated that an average of 1-2 midline UL cells and 4-5 myocardial cells are produced each day ( S3 Fig ). Notably, while the number of midline UL cells increased distal cell numbers remained fairly constant. Distal clusters contained 2 or 3 cells except for occasional instances when there appeared to be only one isolated distal cell. We also observed occasional gaps in the otherwise continuous and tightly packed midline UL nuclei. Additionally, on examining these images we noted the presence of large, round cells between the UL, which is located on the posterior side of the heart, and the myocardial nuclei, which are located in a more anterior region ( Fig 2A and 2B ). Based on their location and lack of actin enriched fibers, we hypothesize that these cells represent myocardial precursors. Clonal analysis of the undifferentiated line To further explore the potential contribution of the UL to myocardial growth we performed a series of EdU pulse/chase experiments. We first established that a 30-minute EdU pulse could be reliably used to label single UL cells for clonal analysis. In particular, we found that when D3 juveniles were exposed to a 30-minute pulse of EdU, the majority (87/110) exhibited no EdU+ UL nuclei and nearly all of the remaining samples exhibited a single EdU+ UL nucleus (22/110). In the one sample with two EdU+ UL cells, labeled nuclei were widely separated, one in the dorsal distal cluster and the other in the ventral distal cluster. Because the pericardium is highly proliferative, we sometimes observed adjacent labeled UL and pericardial cells but we did not observe adjacent labeling of UL and myocardial precursors. Further analysis of these data indicated that distal UL cells have a substantially higher proliferative index (6.2%) than midline UL cells (1.7%). To initiate clonal analysis, we subjected 3D-old juveniles to a 30-minute EdU pulse followed by a 72-hour chase. We then assessed the composition of presumed clones containing at least one EdU+ UL cell. These labeled clusters often consisted of 1-4 EdU+ cells indicating that up to two divisions had occurred during this time period. On the distal end, we observed clones containing paired distal and midline UL cells ( Fig 2D ). In the midline, we often observed pairs of adjacent midline cells ( Fig 2E ). In rare instances we observed what appear to be newly divided midline cells in which one daughter was located posterior to the UL ( Fig 2F ). Additionally, we observed triplet clones consisting of two adjacent midline UL cells associated with a presumptive myocardial precursor ( Fig 2G ). Interestingly, we noted that these presumptive myocardial precursors were present as singlets just anterior to the UL ( Fig 2A and 2B ), while they appeared as doublets when they were located closer to the myocardial nuclei ( Fig 2H ). Based on these observations we generated a model for heart growth ( Fig 2I ). According to this model, distal UL clusters function as self-renewing progenitor populations, dividing asymmetrically to produce midline cells while maintaining a steady population of distal progenitors. Each resulting midline cell either divides symmetrically to further increase UL length or asymmetrically to generate a myocardial precursor. The newly produced precursors then detach from the UL, migrating anteriorly along existing myofibrils as they divide to produce two differentiating myocardial daughters. To more definitely test the hypothetical contribution of the UL to myocardial growth, we traced individual midline UL cell lineages using a photoconvertible Kaede reporter. The Mesp regulatory element ( 97 ) was used to drive expression of a green->red photoconvertible Kaede protein ( 98 ) in heart cell nuclei in transgenic juveniles ( Mesp>Kaede::NLS ( 99 )). In each sample either a single midline nucleus, or a cluster of nuclei near the ventral or distal end of the UL were photoconverted and juveniles were then cultured separately. Photoconverted lineages in cultured juveniles were imaged every 24 hours for up to five days. As predicted by the low proliferative index observed in our EdU pulse experiments (see above), UL cell divisions were rarely observed. Despite this challenge, we were able to document both symmetric ( Fig 2J and S4A ) and asymmetric divisions of midline UL cells ( Fig 2J , S4B and S4C ) as predicted by the EdU clonal data. Critically, we were able to document that asymmetric midline UL divisions gave rise to one daughter that remained associated with the UL and one daughter that migrated anteriorly, presumably serving as a myocardial precursor ( Fig 2J , S4B and S4C ). We also observed a single labeled cell in the midline (a presumed myocardial precursor, S4D Fig ) that migrated anteriorly. Even after four days, we did not observe a subsequent division of this presumed myocardial precursor indicating that myocardial precursor division did not occur or that division was disrupted due to phototoxicity. In another sample ( S4E Fig ) we labeled a single presumed myocardial precursor that appeared to differentiate as indicated by shifts in nuclear morphology, eventually taking on the narrow, ovoid nuclei characteristic of differentiated myocardial cells. In one of our samples, we were able to observe the division of two labeled, adjacent midline UL cells close to the ventral end of the UL ( Fig 2J ). As suggested by previous observations ( Fig 1K , 2F), these cells appear to divide along the anterior/posterior axis producing one anterior daughter that remains in the UL and a posterior daughter that is no longer associated with the UL. Intriguingly, one of these midline UL cells appeared to divide symmetrically (blue arrow in Fig 2J , blue-shaded cell MUV1 in Fig 2I ) producing two midline UL daughters while the neighbouring cell appeared to divide asymmetrically (red arrow in Fig 2J , red-shaded cell MUV0 in Fig 2I ) producing one midline UL daughter and one daughter that migrated anteriorly and along the Z-axis, ending up on a different confocal plane. Based on this data we have further refined our model of UL division and heart growth ( Fig 2I ). In particular, we propose that once a new midline UL cell is generated following asymmetric division of a distal UL, it divides to produce midline UL daughters that exhibit distinct division patterns. One daughter divides symmetrically while the other daughter divides asymmetrically to produce a myocardial precursor. Although the details of this model may shift as more data is collected, our data strongly supports the following core premise: The UL constitutes a cardiac progenitor population that plays a central role in myocardial growth. In accordance with this premise, we will subsequently refer to UL cells as cardiomyocyte progenitors. Transcriptional profiling of C. robusta heart cell types To delineate the transcription profile of the cardiomyocyte progenitors and other heart cell types we performed single-cell RNA sequencing of two adult heart samples ( Fig 3 ). Protocols were optimized to ensure dissection resulted in intact hearts that were subsequently dissociated into pools of viable cells that represented comprehensive and distinguishable cell types. To ensure high viability during encapsulation we deployed a protocol developed by the Klein Lab. As seen in relation to blood cells, the use of cell encapsulation solutions that better match the osmotic environment of cells in marine organisms produced a substantial increase in viability for dissociated C. robusta heart cells [87]. Bioinformatic analysis was performed as described (see methods) to identify distinct cell states. Because dissected hearts contain blood cells, we aligned our data against existing scRNA-seq data from the Klein Lab and found that many of our clusters closely matched blood cell states (grey in Fig 3A ), leaving us with eight presumptive heart cell clusters ( Fig 3A ). Nearly all of these clusters were distinguished by highly enriched expression for a number of transcription factors ( Fig 3B ) along with other marker genes ( Fig 3C-F ). Interestingly, the central cluster (Cluster 1, Fig 3A ) was not as transcriptionally distinct in regards to both of these criteria. The lack of distinct expression for Cluster 1 was also evident when we ran a Pearson’s correlation coefficient analysis to assess cluster similarity based on global gene expression ( S5 Fig ). In particular, this analysis indicated that Cluster 1 was most similar to Cluster 7. Download figure Open in new tab Figure 3. Transcriptomic analysis of adult heart scRNA-seq data. (A) UMAP of scRNA-seq data. Cardiac clusters colored according to Louvain clustering, presumptive blood cell clusters in grey. (B) Heatmap of transcription factor expression. (C) Heatmap displaying three of the most highly enriched marker genes for each cluster. (D) Heatmap of additional marker genes associated with each cluster. (E) Heatmap of marker genes for cell proliferation. (F) Heatmap of signaling pathway genes in Cluster 7. Z-score was derived from comparing all 8 heart clusters, but only Cluster 7 is shown. (G) Louvain sub-clustering analysis of Cluster 7 (UL) and Cluster 8 (Neural/Pacemaker, N/P). (H-I) Heatmaps comparing expression for genes of interest across each UL sub-cluster (H) or Neural/Pacemaker sub-cluster (I) . We next used marker gene expression to provisionally assign each of these eight clusters to one of the distinct cell types identified through our microanatomical analysis ( Fig 3D ). Through this approach we identified two presumptive pericardial clusters, Pericardial-A (Cluster 2) and Pericardial-B (Cluster 3). Previous studies have indicated that the C. robusta pericardium arises from the second heart precursor (SHP) lineage ( 84 ), and both of these clusters express known SHP markers including Matrilin ( MATN ) 1/3/4 and the transcription factor Dachshund ( DACH ) ( Fig 3D ; see S1 Table for list of analyzed genes) ( 100 ). The strong correlation between these two clusters ( S4 Fig ) supports their co-assignment as pericardial sub-types. Cluster 1 was difficult to assign due to the lack of unique markers. However, we tentatively designated this cluster as representing differentiated myoepithelial cells due to somewhat enriched expression for numerous orthologs to muscle-related genes including Muscarinic Cholinergic Receptor 2 ( CHRM2 ), Titin ( TTN ), troponin-like 1 ( TNNL1 ), Myosin Heavy Chain 6 ( MYH6 ), myosin heavy chain 7 ( MYH7 ), Myomesin-1 ( MYOM1 ) ( Fig 3D ). We designated Cluster 4 as representing epicardial cells based primarily on reporter analysis for the highly enriched marker gene synaptotagmin-15 ( SYT15 ) ( S6 Fig ). Additionally, this cluster displays enriched expression of the ortholog to Sine Oculis homeobox 1/2 ( Six1/2 ), a transcription factor that is required for mammalian epicardial progenitor development ( 101 ). We hypothesize that Cluster 5 represents cells lining the blood sinuses adjacent to the heart, which may be equivalent to the vertebrate outflow tract ( 102 ). This cluster displays enriched expression of orthologs to Islet ( Isl ) and E26 transformation-specific or Erythroblast Transformation Specific ( ETS1) , which are known transcriptional markers for C. robusta blood sinus cells ( 102 ). In addition, both Isl and ETS1 play conserved roles in vertebrate outflow tract specification ( 103 , 104 ). We also noted that Cluster 5 displays specific, highly enriched expression of the orthologs to transgelin ( TAGLN ), an established cardiac outflow tract smooth muscle marker ( 105 ) and the smooth muscle marker myosin light chain ( MYL )- 12A/12B/L9 ( 106 ). Our data indicate that Cluster 6 may represent a population of endothelial-like cells. Although endothelial cells are considered to be a vertebrate-specific innovation ( 107 ), it is possible that a precursor cell-type exists in C. robusta . In line with this hypothesis, Cluster 6 displays enriched expression of Gata.b , the ortholog to GATA-2 which is necessary for proper endothelial development in vertebrate models ( 108 , 109 ). Cluster 6 also displays expression of orthologs to the endothelial and smooth muscle marker sphingosine-1-phosphate receptor 1 ( S1PR1 ) along with hairy and enhancer of split B ( HES-B ) a transcription factor that is required for endothelial blood vessel-fate specification in mice ( 110 ). We were particularly interested in identifying which of the presumptive heart cell clusters represented the cardiomyocyte progenitors (UL). Cluster 7 displays highly enriched expression of transcription factors associated with early embryonic cardiac lineage specification in C. robusta and other organisms, including the sole C. robusta orthologs to GATA4/5/6 and Nkx2.5 ( Fig 3B ). These expression patterns may reflect re-initiation of the embryonic cardiogenic specification program during post-larval cardiogenesis. Cluster 7 also displays expression of the ortholog to SET and MYND domain containing 1 ( SMYD1 ) which is necessary for embryonic heart proliferation and morphogenesis in mice ( 111 – 113 ). To further explore whether Cluster 7 represents the heart progenitor population, we examined the relative expression of five genes associated with proliferation. This analysis revealed strong, and relatively specific, enrichment for four of these proliferative marker genes in Cluster 7 ( Fig 3E ). To more definitely test the hypothesis that Cluster 7 represents that cardiomyocyte progenitor lineage (UL), we examined the in-situ expression pattern for the most uniquely enriched gene in this cluster (KH.C1.1154, also represented by the NCBI gene entry XM_026835482). This gene encodes a tunicate-specific, uncharacterized protein that we have named CrUL1 . In situ expression analysis in juvenile samples demonstrates that CrUL1 is only expressed in the heart. Critically, as predicted by our clustering analysis, CrUL1 expression is strongly enriched in cardiomyocyte progenitors, including both midline and distal portions of the UL ( S7A Fig ). CrUL1 is also weakly expressed in a bulging region on the opposite side of the heart, potentially associated with the raphe. Intriguingly, CrUL1 mRNA is strictly localized to the anterior side of each progenitor cell. The UL1 protein is predicted to have a signal peptide ( 114 ) and the CrUL1 mRNA appears to be localized to the anterior side of the UL, potentially facilitating unidirectional secretion into the extracellular matrix (ECM) on this side. We also conducted in situ expression analysis for Gata.a ( S7B Fig ). As predicted by our scRNA-seq data, Gata.a is highly expressed in the UL along with myocardial cells in juvenile samples ( S7B Fig ). We noted that the expression of some TFs and markers appeared to be more enriched in specific sets of cells within the putative cardiomyocyte progenitor cluster (Cluster 7). To further explore these differential expression patterns, we performed an additional Louvain clustering analysis on Cluster 7 alone leading to the identification of three sub-clusters (UL-A, B, and C, Fig 3G and 3H ). UL-A displayed enriched expression of the ortholog to transcription factor nuclear receptor subfamily 4 group A ( NR4A ), which regulates the cardiac response to neurohormonal signaling in mice ( 115 ). UL-B is distinguished by high levels of expression of the sole C. robusta ortholog to Tbx2 and Tbx3 , transcription factors associated with specification of the atrio-ventricular canal and associated cardiac conduction cells in mice and zebrafish ( 70 , 116 ). We also noted that the UL marker gene CrUL1 is primarily enriched in UL-B. UL-C is distinguished by high levels of expression for orthologs to SRY-Box Transcription Factor 5/6/13 ( Sox5/6/13 ), basic helix-loop-helix tunicate 1 ( Bhlhtun1 ), and dynein axonemal heavy chain 12 ( DNAH12 ). Intriguingly, the latter two genes are both established markers for the C. robusta inner atrial siphon muscle precursors, a sister lineage to the heart precursors and Bhlhtun1 has been shown to promote self-renewal in these siphon muscle precursors ( 99 , 117 ). Expression of signaling pathway genes in cardiomyocyte progenitors We next assessed the expression of orthologs to receptors or downstream target genes from a range of developmental signaling pathways in the presumed cardiomyocyte progenitor scRNA-seq cluster (Cluster 7, Fig 3F ). This cluster displayed enriched expression for a number of orthologs to signaling receptor genes including Epidermal growth factor receptor-a ( EGFRa ), Ephrin-a ( Eph.a ), Notch , insulin receptor ( InsR ), Transforming growth factor ( TGF-β receptor-Ia, -Ib, -Ic, -IIb ) and Retinoic Acid Receptor alpha ( RARA ). Orthologs to two genes from the cytochrome P450 superfamily ( CYP26A1 ) involved in RA degradation along with dehydrogenase/reductase 7B ( DHRS7B ) which is involved in RA inactivation were also highly expressed in the UL cluster (cluster 7). Additionally, a number of genes encoding orthologs to secreted signaling factors including Fgf9/16/20 and Fgf-tun2 displayed enriched expression. We also noted enriched expression of the C. robusta ortholog to the Hedgehog-signaling associated gene smoothened ( SMO ). Notably, the presumed progenitor UL cluster displayed expression for numerous orthologs to genes associated with canonical Wnt signaling. These included Wnt receptors such as Frizzled ( Fzd-3/6/7 ), Fz4 and Fz-a , the co-receptor LDL Receptor Related Protein ( LRP-4/5/6 ), along with Wnt10A and β-Catenin-1 ( CTNNB1 ). UL clusters also displayed enriched expression of orthologs to three genes that are characteristically up-regulated in response to cWnt signaling to mediate auto-negative feedback, Dickkopf ( DKK ), secreted frizzled-related protein 1 ( sFRP1 ) and Axin [96, 97]. As various cWnt signaling components were highly expressed in the UL, we were particularly interested in examining the expression of orthologs to cWnt pathway genes within the UL sub-clusters ( Fig 3I ). UL-A and UL-C both expressed relatively high levels of orthologs to LRP5/6 along with Axin , indicating that these cells are subjected to cWnt signaling ( 118 ). Based on these findings we began to assess the contribution of cWnt signaling to cardiac progenitor proliferation using pharmacological assays. Wnt signaling suppresses cardiomyocyte progenitor division Based on previously characterized roles for canonical Wnt signaling in retention of proliferative stem cell populations ( 118 ), we hypothesized that this pathway promotes C. robusta cardiomyocyte progenitor proliferation. To test this hypothesis, we treated juveniles with IWR-1-endo (IWR), a cell-permeable molecule that stabilizes Axin and thus promotes β-catenin degradation to disrupt cWnt signaling ( 119 ). We then assayed the impact on progenitor proliferation using a 30-minute EdU pulse assay. Strikingly, application of IWR led to a significant increase in the frequency of cardiomyocyte progenitor proliferation ( Fig 4A and 4B ). As seen in our clonal analysis, the majority of control juvenile hearts displayed no EdU+ progenitors and nearly all of the remaining control hearts displayed one EdU+ progenitor. In contrast, following a 24-hour treatment with IWR, the majority of juvenile hearts displayed two or more EdU+ progenitors. A robust and significant increase in the incidence of EdU+ progenitors was seen in both midline and distal UL cells (Fig4 C and 4D). These results contradict our initial hypothesis, indicating that cWnt suppresses cardiomyocyte progenitor proliferation. To determine whether IWR also impacts the rate of cardiomyocyte progenitor division, we conducted EdU pulse/chase experiments. This analysis indicated that division rates were similar in control and IWR-treated cardiomyocyte progenitors (approximately one division per day, Fig 4E ). Thus, cWnt signaling appears to specifically decrease the frequency of cardiomyocyte progenitor division and not the ensuing rate of division. To further test this hypothesis, we applied two cWnt agonists with distinct mechanisms of action, 6-Bromoindirubin-3′-oxime (BIO) ( 120 ) which blocks GSK3 activity, and SKL2001 which blocks the interaction between Axin and ß-catenin ( 121 ). Although we extended the EdU pulse length to 4 hours, labelling of progenitors remained infrequent in control juvenile hearts ( Fig 4B ). Despite this low baseline, we were still able to detect a significant reduction in the number of juvenile hearts displaying EdU+ progenitors in both treatment conditions ( Fig 4A and B ). To alleviate concerns regarding the specificity of these pharmacological treatments, we used a targeted transgenic approach to disrupt the cWnt pathway. In these experiments, we deployed a characterized Foxf enhancer element ( 122 ) to drive a c-terminal truncated form of TCF specifically in embryonic heart lineage cells ( FoxfΔepi>TCFdn ). FoxF enhancer driven expression begins at the tailbud stage, well before heart morphogenesis and growth which occur after metamorphosis. Because the c-terminus of TCF mediates interaction with β-catenin, truncated TCF functions in a dominant negative fashion, suppressing cWnt-dependent gene expression ( 123 , 124 ). Comparisons between hearts in transgenic FoxfΔepi>TCFdn juveniles versus control juveniles ( FoxfΔepi>LacZ ) revealed that transgenic disruption of the cWnt pathway in the heart lineage from an early time point consistently and significantly increased heart size along with the number of cardiomyocyte progenitors ( Fig 4F and 4G ). Quantification confirmed that there was a robust and significant increase in midline UL numbers in the experimental samples. Strikingly, despite this substantial increase in midline UL numbers, distal UL clusters did not appear to be expanded. Furthermore, there were no detectable abnormalities in the structure of these enlarged hearts ( Fig 4F ). Taken together, these results strongly support a role for cWnt signaling in suppressing cardiomyocyte progenitor proliferation, potentially contributing to the low proliferative index observed in these cells during early stages of heart growth. Download figure Open in new tab Figure 4. Inhibition of the canonical Wnt pathway leads to an increase in the frequency of cardiomyocyte progenitor division. ( A ) Representative images showing the incidence of EdU+ UL nuclei (magenta) in juveniles treated with DMSO, the cWnt antagonist IWR-1-endo, or the cWnt agonist BIO. Dotted lines demarcate the UL of each heart. Cyan represents DAPI staining. ( B-D ) Graphs showing the % of hearts displaying specific numbers of all EdU+ UL cells ( B ), EdU+ distal UL cells ( C ) or EdU+ midline UL cells ( D ) in control and experimental samples as indicated. ( E ) UL division rates. To calculate these rates, juveniles were pulsed for 30 minutes with EdU followed by a chase period of 36 hours and assessed for the number of EdU+ nuclei. The average number of EdU+ nuclei in these chase experiments was then divided by the average number of EdU+ nuclei in the pulse-only experiments, then divided by 1.5 to get a per 24-hour value. A value of 1 indicates one division per day. 0 values were not included (see supplementary data). ( F ) Representative images of hearts from juveniles electroporated with the control construct FoxfΔepi>LacZ (left) or experimental construct FoxfΔepi>TCFdn (right). ( G ) Graph depicting the percent of juvenile hearts with substantially increased numbers of midline UL cells (greater than one standard deviation above the mean in controls). See methods for statistical analysis. T-tests performed for B, C, and D. Bar and asterisks (*) in B indicate p-values of less than .05, and (***) in C, D, and G indicate p-values of less than 0.0001. Transcriptional profile of neuron-like cells in the C. robusta heart We noted that one of the cardiac cell clusters in our scRNA-seq data set (Cluster 8) displayed enriched expression for a number of orthologs to neuron-associated transcription factors including One Cut Homeobox 1 ( Onecut1) and Achaete-scute complex-like-a ( Ascl.a) ( Fig 2B ) ( 125 – 127 ). Additionally, this cluster was distinguished by unique and highly enriched expression of the gene encoding the sole C. robusta ortholog for the tachykinin receptor ( TACR, Fig 2C ). Intriguingly, none of the heart cells in our scRNA-seq dataset display expression of tachykinin , suggesting that this neuropeptide is produced by extrinsic neurons. To begin exploring the identity and function of the cells in this presumptive neural-like cardiac cluster we examined the expression of orthologs to other neural markers, including genes encoding neurotransmitter and neuropeptide signaling components within Cluster 8 ( Fig 2D ). This analysis revealed highly enriched expression for a number of orthologs to well-established neuronal markers including Neurexin-1 ( NRXN1 ), and Neural Cell Adhesion Molecule ( NCAM ) (need citations). Cluster 8 also displayed highly enriched expression of orthologs to a number of genes involved in the synthesis and processing of dopamine including - tyrosine hydroxylase ( TH ), dopa decarboxylase ( DDC ) and dopamine beta-hydroxylase ( DBH ) which participates in converting dopamine into noradrenaline. This cluster also displayed enriched expression for orthologs to a number of genes encoding neurotransmitter receptors including serotonin receptor subunit 5-hydroxytryptamine receptor 1D ( HTR1D ) and nicotinic alpha 2 subunits-2 & 7 ( CHRNA2 and CHRNA7) ( 128 ). Additionally, Cluster 8 displayed enriched expression of an ortholog to Solute Carrier Family 6 Member 11 ( SLC6A11 ), involved in uptake of the inhibitory neurotransmitter gamma-aminobutyric acid (GABA). Lastly, this neuronal-like cluster displayed enriched expression of the ortholog to Choline Acetyltransferase ( ChAT ), which is necessary for the biosynthesis of acetylcholine. Thus, the gene expression profile for this cluster clearly indicates representation of neural-like cells associated with the heart. Strikingly, Cluster 8 displayed highly enriched expression of the ortholog to prohormone convertase 2 ( PC2 ), which is a neuropeptide enzyme necessary for cleaving tachykinin and other neuropeptides into functional forms. PC2 reporter signal has previously been shown to label neural plexuses that form rings around the distal ends of the C. robusta myocardial tube ( 96 ), leading us to hypothesize that Cluster 8 may contain cells from these cardiac neural rings ( 129 ). Intriguingly, Cluster 8 also displayed enriched expression of orthologs to vertebrate marker genes for both intrinsic cardiac neurons ( ChAT , DBH , and Calbindin ( CALB ) ( 130 – 132 ) and cardiac pacemakers ( hyperpolarization activated cyclic nucleotide-gated potassium channel 4 ( HCN4 ), Fibroblast growth factor 13 ( FGF13 ), and TH ( 130 , 131 ), along with markers associated with cardiomyocytes ( TTN , TNNL1 , and MYH6 , Fig 2D ). Taken together, our analysis suggests that Cluster 8 may contain a variety of cell types including both intrinsic neurons and pacemaker cells that comprise the cardiac conduction system ( 133 ). To determine if there are indeed transcriptionally distinct neural-like cell-types contained within Cluster 8, we performed an additional Louvain clustering analysis on this cluster alone leading to the identification of 3 sub-clusters ( Fig 3G ). We then examined the expression of a variety of orthologs for neural genes within these sub-clusters ( Fig 3H ). Neural/Pacemaker-A (NP-A) was distinguished by highly enriched expression of the orthologs to the neuronal peptidergic marker PC2 and Transient receptor potential cation channel subfamily C, member 6 ( TRPC6) , which is modulated by TACR signaling ( 134 ). Neural/Pacemaker-B (NP-B) was distinguished by highly enriched expression of the C. robusta orthologs to CHAT, PAX3 , and the tachykinin receptor (TACR). NP-B also displayed relatively high levels of expression for orthologs to the cardiac pacemaker markers, HCN4 and FGF13 . Neural/Pacemaker-C (NP-C) was distinguished by highly enriched expression of orthologs to the transcription factor DNAH12 as well as Tbx2/3 , the sole C. robusta ortholog to TBX2 and TBX3 , a pair of paralogous transcription factors that play a key role in development of the vertebrate cardiac conduction system. NP-C also expressed relatively high levels of the ortholog to the pacemaker marker HCN4 and was distinguished by enriched expression of orthologs for a number of genes associated with signaling pathways, including the RA, Wnt, hedgehog and TGF-B pathways. Based on this analysis, we hypothesized that NP-A represents a population of intrinsic cardiac neurons while NP-B and NP-C represent mixed populations of intrinsic neural and pacemaker cell types. Profile of cardiac intrinsic innervation in adult C. robusta hearts In order to begin characterizing the location of morphological distinct neural sub-types represented by Cluster 8 in our transcriptional atlas, we deployed stable transgenic lines to label neural-like cells in the adult C. robusta heart ( S8A Fig ). We first used the PC2>Kaede reporter to label peptidergic neurons potentially represented by sub-cluster NP-A (the only neural-like sub-cluster displaying highly enriched PC2 expression). As observed previously ( 96 ), this reporter labeled neural rings associated closely with the distal myocardium at both the dorsal (visceral) and ventral (hypobranchial) ends of the heart ( S8B Fig ). Based on scRNA-seq data for sub-cluster NP-A ( Fig 3G ), we hypothesize that these rings are primarily composed of intrinsic neurons. To label cholinergic neural-like cells potentially represented by sub-cluster NP-B (the only neural-like sub-cluster displaying highly enriched ChAT expression) we used the VACHT>CFP reporter. Both ChAT (choline acetyltransferase) and VACHT (vesicular acetylcholine transporter) are generally co-expressed in cholinergic neurons. Although this reporter also labeled both distal neural rings, the zone of expression appeared to be limited in comparison with PC2>Kaede ( S8C and S8E Fig ). VACHT>CFP positive cells appeared to form a proximal narrow band exhibiting high levels of reporter expression along with a more distal dispersed band exhibiting lower levels of reporter expression. Based on the scRNA-seq data for sub-cluster NP-B ( Fig 3G ), we hypothesize that these VACHT+ bands contain pacemaker cells. To further test this hypothesis, we examined expression of VACHT>CFP along with a Troponin reporter ( Troponin>RFP ) in D30 juveniles. This analysis revealed that the VACHT and Troponin reporters co-label a set of neural-like cells at either end of the heart ( S8D Fig ). Thus, it appears that a subset of neural-like cells associated with the distal ends of the myocardium may represent pacemaker cells that are derived from cardiomyocyte progenitors as observed in vertebrates. We also conducted FISH for the pacemaker marker gene HCN2/3/4 . Although it was difficult to detect staining, we did observe enriched staining in several cells located near the dorsal distal end of the heart, suggesting these are putative pacemaker cells (S7C Fig). We next observed the expression pattern of a TH>Kaede reporter. Strikingly, this reporter was only expressed in the ventral cardiac plexus (S7E Fig). Using double TH>Kaede , VACHT>CFP lines, we were able to discern that Kaede (TH+) and CFP (VACHT+) cells do not overlap within the ventral plexus ( S8E Fig ). Thus, these distal TH + cells may correspond to the NP-C sub-cluster which displays enriched expression of TH but not of VACHT . It also appears that the TH+ region of the plexus may include the distal-most portion of the ventral plexus, including areas that do not express the PC2>Kaede reporter ( S8F Fig ). Developmental profile of cardiac intrinsic innervation in C. robusta juvenile hearts We next employed PC2>Kaede , VACHT>CFP, and TH>Kaede stable transgenic lines to analyze initial innervation of juvenile hearts ( S9 Fig ). In early juvenile samples (3D) a single VACHT>CFP+ cell was observed at both the ventral and dorsal ends of the heart ( S9A Fig ). In D4 double transgenic VACHT>CFP - PC2>Kaede juveniles, we were able to observe multiple distinct VACHT-, PC2+ or VACHT+, PC2- cells ( S9B Fig ). Based on these patterns of reporter expression, these distinct neurons may be represented by sub-clusters NP-A or NP-B respectively ( Fig 2I ). There were also a number of VACHT+, PC2+ neural-like cells in these D4 hearts that may not be represented by one of the NP sub-clusters. By day five, the VACHT>CFP+ ventral plexus appeared to consist of two distinct, large foci ( S9C Fig ). As seen in the adult heart, TH>Kaede+ cells are restricted to the ventral plexus in early juvenile hearts and do not colocalize with VACHT>CFP+ cells in double transgenic ( VACHT>CFP - TH>Kaede ) juveniles ( S9D Fig ). In D7 juveniles, we were able to observe extensions from the ventral TH>Kaede+ neurons that appear to innervate adjacent progenitors in the distal UL ( S9E and S9F Fig ). In older juveniles (D10-D12), PC2>Kaede+ neurons form a small plexus at each end of the heart ( S9G and S9H Fig ). As seen in earlier stages, these plexuses are in close proximity to the distal UL clusters and exhibit thin neurite-like projections associated with the adjacent line of cardiomyocyte progenitors ( S9H Fig ). To further test the hypothesis that some of VACHT+ or TH+ neural-like cells represent pacemakers derived from a mesodermal cardiomyocyte progenitor population, we employed juveniles co-expressing two reporters, Phox2b>Unc76::GFP and Mesp>H2B::mCherry . In these juveniles the Phox2b reporter was used to label cells associated with the NP-A and NP-B sub-clusters (which both display enriched expression of Phox2b along with pacemaker marker genes) while the Mesp reporter was used to label the mesodermal cardiac progenitor lineage ( 97 ). Although most Phox2b>GFP+ neural-like cells did not display co-expression of Mesp>H2B::mCherry , a small subset of these cells (2 of 37) co-expressed both reporters ( S10 Fig ). These double-labeled cells may represent pacemakers that arise from a bi-potential cardiomyocyte/pacemaker lineage as seen in vertebrate hearts ( 135 , 136 ). In summary, our data indicates that neural-like cells associated with the distal ends of the forming myocardial tube arise in early juveniles, just as the heart is completing morphogenesis. Additionally, our reporter analysis combined with our scRNA-seq data indicates that there is a complex array of neural– like cell types in these early distal plexuses, including both intrinsic neurons and cardiac pacemakers. Assessing the role of neural inputs in regulation of C. robusta heart rate Based on the presence of VACHT+ and TH+ cardiac neural-like cells we predicted that heart rate would be modulated by cholinergic and dopaminergic inputs respectively. Additionally, our scRNA-seq data detected highly enriched expression of the sole C. robusta ortholog to the tachykinin receptor ( TACR ) in the presumed cardiac neural-like cluster (Cluster 8, Fig 2C ). Thus, we also hypothesized that C. robusta heart function was modulated by tachykinin signaling. To test these candidate neural inputs, we quantified heart rate in D5-D6 juveniles both before and after exposure to a set of drugs targeting these pathways (S11A Fig). Application of the TH inhibitor α-methyl-para-tyrosine (AMPT) lowered heart rate slightly, but this result was not significant. In contrast, application of the acetylcholine agonist acetylcholine chloride and the TACR inhibitor aprepitant had no discernible impact on heart rate. Based on these results we began to investigate other potential roles for these neural signals. Assessing the role of tachykinin signaling in heart growth Based on the proximity of PC2+ neural-like cells to the distal progenitors at each end of the undifferentiated line in juvenile and adult hearts (S9F’ and S9G’’ Fig), we hypothesized that these neural-like cells may regulate distal progenitor cell behaviors. To more closely examine the relative positions of these two cell types in the adult heart, we dissected and flattened distal growth zones in transgenic PC2>Kaede adults. Strikingly, we observed PC2+ neurites interspersed within the densely clustered nuclei of the distal growth zone (S11B Fig). PC2+ neurites were also observed along the border of the distal cluster, potentially associated with the transition zone containing proliferative myocardial or pericardial precursors. Previous studies have shown that tachykinin family members substance P and substance K promote cell proliferation in cultured vertebrate muscle cells ( 137 ). Additionally, a more recent study found that substance P promotes cell proliferation in cultured cardiomyocyte progenitors ( 138 ). Thus, we hypothesized that tachykinin signaling may play a similar role in C. robusta , promoting heart growth by increasing proliferation of cardiomyocyte progenitors in the distal UL growth zones. To begin investigating a potential role for either tachykinin or cholinergic signaling in heart growth, we injected C. robusta adult hearts with the TACR inhibitor aprepitant, the cholinergic inhibitor atropine, or with control carrier solutions and cultured the otherwise intact animals for 24 hours. In order to assay myocardial proliferation, hearts were removed after the 24 hours of drug treatment and incubated in EdU for 24 hours. As predicted by our hypothesis, aprepitant-treated hearts possessed a significantly lower percentage of EdU+ cells in the distal region in comparison with matched controls, whereas inhibiting cholinergic signaling by applying atropine did not impact cell proliferation ( Fig 5A - 5E ). However, it was difficult to discern which cell populations were impacted. To more precisely examine the role of tachykinin signaling on heart growth, we treated D3 juveniles for four days with one of two distinct tachykinin receptor antagonists, aprepitant or spantide II, and assayed the number of midline progenitors in the resulting D7 juveniles. As predicted by our hypothesis, both aprepitant and spantide II significantly reduced the number of midline UL cells in comparison to controls ( Fig 5F - 5J ). To explore the specificity of this result, we treated juveniles with inhibitors against a number of other neural signaling pathways including the acetyl-choline receptor antagonist atropine, the alpha-1 adrenergic receptor antagonist Doxazosin Mesylate and the TH pathway inhibitor AMPT. None of these drugs had any discernible impact on heart growth or midline progenitor numbers (S11C-11I). Download figure Open in new tab Figure 5. Tachykinin signaling promotes distal progenitor proliferation. (A) Diagram of cardiac and neural anatomy in an adult. Green lines denote possible extensions of the central ganglion to each side of the heart. (B) Diagram of adult heart. Boxes indicate areas shown in panels C and D. (C) EdU (cyan) labeled distal UL section of control adult heart. (D) EdU (cyan) labeled distal UL section of an adult heart treated with aprepitant for 24hrs. In C and D, red indicates SYTO labeled nuclei. ( E ) Violin plot quantification of percent cells EdU+ per treatment, n= 3 hearts per condition. (F) D7 control heart. (G) D7 heart treated with aprepitant. (H) D7 heart treated with Spantide. (I) Enlargements of UL cells from boxed areas of F, G, and H. In F-I, cyan is DAPI and magenta is phalloidin. Asterisks mark individual UL nuclei in magnified panel. (J) Violin plot quantification of normalized midline UL cells per treatment. Midline UL count normalized to control per trial. 3 trials per condition. N=49 DMSO, N=49 Aprepitant, N=46 Spantide. (K) Diagram of relative cardiac and neural anatomy in a juvenile (left) and diagram of juvenile heart anatomy. (L) D11 control DMSO treated heart. (M) D11 heart treated with Aprepitant for 24hrs. In L and M, outline of the heart and UL depicted by dotted lines. Cyan indicates DAPI staining and magenta indicates EdU labeled nuclei. (N) Violin plot of percent animals per trial with one or more EdU+ progenitor in the dorsal, midline, or ventral UL per trial. 4 trials each condition, N= 228 DMSO, N=213 Aprepitant. A t-test was performed between the averages per trial. To more directly assay the impact of tachykinin pathway inhibition on cardiomyocyte progenitor proliferation, we treated D10 juveniles with aprepitant or DMSO for 24 hours and labeled proliferating cells using a 6-hour EdU pulse ( Fig 5L - 5N ). As predicted by our hypothesis, we observed a significant reduction in the number of EdU+ progenitors in both the ventral and dorsal distal regions in comparison to controls ( Fig 5N ). In contrast, no reduction was observed in the number of EdU+ midline progenitors. Taken together, our data strongly supports a role for tachykinin signaling in promoting distal myocardial progenitor proliferation. Expression patterns for the genes encoding tachykinin and its receptor Our analysis of the scRNA-seq data indicates TACR is uniquely expressed in the neural-like sub-cluster (Cluster 8, Fig 2C ). Additionally, further analysis of this cluster revealed that the Neural/Pacemaker-B sub-cluster displays highly enriched expression of TACR in comparison with the other two sub-clusters ( Fig 2I ). As described in previous sections, it appears that this sub-cluster represents a mixed population of VACHT+ intrinsic neuron or pacemaker cells that are found in both the ventral and distal plexuses in close association with distal cardiomyocyte progenitors. Based on this data, we hypothesized that tachykinin signaling impacts heart growth indirectly, impacting VACHT+ neural/pacemaker cells which then directly regulate proliferation in nearby distal progenitors. To begin investigating this hypothesis we used fluorescent in situ hybridization (FISH) to visualize TACR expression in juvenile hearts. Although the TACR expression pattern was often obscure, we did occasionally observe expression in cells adjacent to distal progenitors (S12 Fig). We next began to explore the expression of the sole C. robusta ortholog to the vertebrate tachykinin family neuropeptides ( Cr-TK ). Previous studies have documented that Cr-TK is expressed exclusively in the brain, intestine, endostyle and gonads ( 139 ). In line with this previous data, we did not detect expression of Cr-TK in any of the cardiac or blood cell clusters in our scRNA-seq dataset (S13C Fig). We therefore employed a Tachykinin reporter ( TK>CFP ) to document the source of tachykinin in D7 juveniles. As predicted by our scRNA-seq data, the TK reporter was not expressed in the heart (S13B Fig). Instead, in line with previous studies, we observed reporter expression in the central ganglion (S13A Fig). Characterization of central ganglion neurons innervating the C. robusta heart According to our current model, tachykinin produced by central ganglia cells interacts with TACR expressed by cardiac neural-like cells associated with the distal pools of cardiac progenitors. In vertebrates, cholinergic extensions of the vagal nerve innervate the heart ( 71 , 140 ). Examination of the C. robusta nervous system has not revealed any innervation from the central ganglion to the heart ( 96 , 141 ). To explore potential extrinsic innervation of the C. robusta heart by the CNS we used high-intensity confocal laser settings to visualize neurons labeled by PC2>Kaede in the hearts of young C. robusta adults and D10 juveniles ( Fig 6A and 6B ). This analysis revealed a network of thin PC2>Kaede+ projections that appear to branch off the relatively thick projections of the central ganglia and extend over a large surface area of the heart in both adult and juvenile stages. Although these presumably extrinsic neural projections were highly variable in morphology ( Fig 6A ’ vs Fig 6B ’) they consistently appear to connect with dorsal and ventral PC2>Kaede+ neural plexuses in both young adults and D10 juveniles ( Fig 6A ’, 6B’, and 6B’’). The ventral (hypobranchial) cardiac neural plexus appears to be innervated by neurites extending from central ganglion axons located along the longitudinal muscle band on the right side of the animal. The dorsal (visceral) cardiac neural plexus appears to be innervated by neurites extending from central ganglion axons that wrap around the posterior end of the stomach before reaching the heart. Download figure Open in new tab Figure 6. Neurites extending from the central ganglia appear to innervate cardiac distal neural plexuses. (A) Young adult displaying PC2>Kaede expression (white). A’ shows the magnified boxed area from A . Arrows highlight the connection between central ganglion projection to the ventral cardiac neural plexus. A’’ depicts a diagram of proposed cardiac innervation. Extrinsic neurons (purple) from the central ganglion connect to dorsal and ventral cardiac neural plexuses (green). (B) PC2>Kaede expression in a D10 juvenile (white). B’ shows the magnified boxed area from B . Arrows indicate connection from central ganglion projection to ventral and dorsal cardiac neural plexuses (arrowheads). B’’ shows a heart from a D30 juvenile displaying the same connections as in A and B . Yellow dotted line outlines the heart. (C) D10 transgenic juvenile displaying VACHT>CFP reporter expression (magenta). C’ shows the magnified boxed area from C, including brightfield to reveal the position of the heart. C’’ Depth-encoded view of the same heart reveals close proximity between a central ganglion neurite (arrows) and a cell in the VACHT+ ventral cardiac neural plexus (arrowhead). To further explore potential similarities between vertebrate and C. robusta cardiac innervation we determined whether extrinsic neurons that appear to innervate the distal intrinsic plexuses of the C. robusta heart are cholinergic using the stable VACHT>CFP reporter lines. As observed previously, the VACHT reporter labeled cardiac neural plexuses on both ends of the heart ( Fig 6C and 6C ’). Additionally, we observed a network of extrinsic cholinergic neurons (VACHT>CFP + ) that extended along the longitudinal muscle and appeared to innervate the ventral intrinsic plexus ( Fig 6C , C’ and C’’). Targeted knockdown of either tachykinin or the tachykinin receptor severely disrupts formation of the myocardium To more definitively disrupt cardiac tachykinin signaling we used CRISPR-Cas9 to knockdown either tachykinin (Cr-TK) or the tachykinin receptor ( TACR ) and examined the impact on juvenile heart growth. Based on our model we predicted that both of these manipulations would reduce distal progenitor proliferation, leading to decreased heart growth. To knockdown Cr-TK , we used the PC2 enhancer to drive Cas9 expression in the PC2 positive neurons and co-expressed a pair of Cr-TK -specific sgRNAs ubiquitously using the U6 promoter ( 142 ). As documented above, the PC2 enhancer drives reporter expression in both extrinsic central ganglia neurons and distal cardiac plexuses ( Fig 6 ), however Cr-TK is only expressed in the central ganglia (S12 Fig). Thus, we used this approach to determine whether TK produced by extrinsic neurons is required for heart growth. As predicted by our model, we found that 37.5% of D10 TK crispant juveniles showed significantly higher levels of abnormal heart morphology in comparison to controls ( Fig 7A - 7B , and S1-S2 Movies). The partial penetrance of this phenotype aligns well with other CRISPR-Cas9 assays in C. robusta and is considered to reflect mosaic incorporation of plasmids ( 143 , 144 ). Surprisingly, abnormal hearts were not markedly reduced in size but instead appeared to consist of a nearly hollow pericardial cavity with no discernible UL and in which beating myocardial cells were either completely absent or restricted to a small rudiment ( Fig 7A and 7B ). High resolution confocal imaging confirmed this phenotype ( Fig 7C and 7D ). To knockdown Cr-TACR , we used the Mesp enhancer to drive Cas9 expression in cardiac lineage cells and co-expressed a Cr-TACR -specific sgRNA ubiquitously using the U6 promoter. As observed in the TK knockdown assay, loss of TACR severely disrupted heart morphology, generating a similar hollow pericardial cavity phenotype ( Fig 7B, E and F, S3-S4 Movies). Strikingly, loss of the myocardium in these TACR mutant hearts was associated with the absence of PC2+ distal neural plexuses ( Fig 7G and 7H ). Due to the severe nature of the hollow pericardial phenotype, it was difficult to assess the presence of the UL. To determine whether any remnant of the UL was still present in these abnormal hearts we visualized the expression of the UL marker gene CrUL1 in TK mutant hearts. While all control hearts displayed strong CrUL1 expression just posterior to the UL ( Fig 7I ), all abnormal TK mutant hearts lacked any discernible CrUL1 expression ( Fig 7J ). These data provide strong support for our model, indicating that both tachykinin secreted by central ganglia neurites, and tachykinin receptors in cardiac neural-like cells, are required for cardiac progenitor proliferation. Indeed, these assays indicate that tachykinin-dependent progenitor proliferation is not only required for heart growth, but also plays a critical, earlier role in formation or maintenance of the myocardium. Download figure Open in new tab Figure 7. Targeted knockdown of tachykinin in extrinsic neurons or of tachykinin receptor in cardiac lineage cells abrogates formation of the myocardial tube. (A) Transmitted light D10 control heart ( A ) and a CRISPR-Cas9 TACR crispant heart ( A’ ). Note the hollow, empty pericardium lacking an internal myocardial tube. (B) Violin plot of treated and control hearts scored for morphology. When targeting TK , 2/64 control hearts were abnormal and 24/64 TK crispant hearts were abnormal. When targeting TACR , 1/60 control hearts were abnormal and 22/70 TACR crispant hearts were abnormal (three trials per group, p-values displayed in graph.) (B’) Schematic illustrating normal and abnormal heart phenotypes. (C) Representative micrograph of D10 juvenile control heart showing normal heart anatomy including myocardial tube, pericardium, and UL. (D) Representative micrograph of a TK crispant heart showing lack of normal myocardial tube and UL. (E) Representative micrograph of a D4 control juvenile heart stained with DAPI. Panel on right is magnified showing UL (dotted line) and pericardial nuclei (asterisks). (F) Representative micrograph of a TACR D4 crispant heart showing lack of normal myocardial and UL cells within the pericardium (asterisks). 8/8 TACR crispants with abnormal heart morphology had absent UL cells, 3/3 controls had normal UL morphology. (G) D10 control juvenile electroporated with PC2>Kaede (yellow) labeling the dorsal and ventral intrinsic neuronal-like clusters (D, V). (H) Representative D10 TACR crispant demonstrating reduction in PC2 reporter signal in the heart in otherwise PC2+ juveniles. 8/8 TACR crispants with abnormal morphology had absent PC2 reporter in the heart, 3/3 controls had normal PC2 expression. ( I) Representative control showing CrUL1 expression (magenta) in a D7 juvenile heart, all 21 samples displayed normal CrUL1 expression across two trials. ( J) Representative D7 TACR crispant showing lack of CrUL1 expression (magenta) as observed in 11/26 samples across two trials. (K) Schematic model in which tachykinin secreted by central ganglion neurites is received by TACR-expressing cardiac neural-like cells (either intrinsic neurons or pacemakers), causing them to secrete a hypothetical downstream signal that promotes proliferation of adjacent distal myocardial progenitors. Discussion In this paper, we have begun to establish the C. robusta heart as a platform for investigating the contributions of paracrine and neural signals to cardiogenesis along with subsequent growth. In particular, our data indicate that both canonical Wnt and tachykinin signaling play critical roles in heart formation and growth ( Fig 4 and Fig 7K ). We first demonstrated that the undifferentiated line serves as a stem-like progenitor population capable of producing new myocardial cells ( Fig 2 ). Through clonal analysis we delineated distinct division patterns in distal vs. midline progenitor population and generated a model regarding the role of asymmetric divisions in sustaining the distal reserve stem population while promulgating elongation of the UL and myocardial tube ( Fig 2 ). Through scRNA-seq of the adult heart we identified eight distinct cell states, including a cluster of cells that represent the UL ( Fig 3 and S5 Fig ). Analysis of expression patterns within this presumptive UL cluster led us to investigate the role of canonical Wnt signaling, revealing that cWnt negatively regulates proliferation in the UL, potentially contributing to the low proliferative index of these cells and constraining heart growth ( Fig 3 and Fig 4 ). We also began to investigate a cardiac neural-like cell state which appears to include both cardiac intrinsic neurons and pacemaker cells ( S8 - S10 Figs ). Using high resolution microscopy and reporter constructs we mapped distinct cardiac neural sub-types in both adult and juvenile C. robusta hearts ( S8 Fig ). Remarkably, we found that distal neural plexuses appear to innervate adjacent pools of distal progenitors ( Fig 6 ). Based on high levels of tachykinin receptor ( TACR ) expression in this cluster, we investigated the role of this neuropeptide revealing that tachykinin signaling promotes proliferation within the distal myocardial progenitors ( Fig 7 ). Additionally, our data indicates that central ganglion neurites innervate the distal cardiac intrinsic plexuses ( Fig 6 and S13) and that these extrinsic neurons serve as the source of tachykinin required for heart morphogenesis and growth ( Fig 7 ). Progenitor and precursor lineages contribute to growth of the C. robusta myocardium According to our current model of C. robusta heart growth, new myocardial cells are derived from three distinct populations - distal and midline progenitors that reside in the undifferentiated line along with myocardial precursors that divide as they migrate anteriorly before differentiating. Furthermore, we posit that both distal and midline progenitors behave like cardiac stem cells, dividing asymmetrically to self-renew while producing a midline or precursor daughter respectively. We are currently working to identify genetic markers that distinguish between these two progenitor populations. These markers would allow us to identify which of the UL scRNA-seq sub-clusters represent each type of progenitor and thus assess their different transcriptional profiles. Additionally, these markers could be used to generate reporter constructs required for more precise lineage tracing and behavioral analysis. Reporters could also be used for cell sorting and re-sequencing to more definitively assess progenitor cell transcriptional profiles. Once these tools are in place, we will be able to characterize the developmental origins of these distinct progenitor cell types. They will also facilitate exploration to determine whether either of these progenitors exhibit patterns of gene expression or cell behavior reminiscent of characterized stem cells in other organisms, or if they bear any notable similarities to cardiac progenitor populations that reside at the distal ends of the vertebrate embryonic heart tube ( 145 – 147 ). We are particularly interested in discerning whether there are niche-like structures that maintain stemness in these populations. Along these lines, it is intriguing that the anterior side of the UL exhibits enriched actin, WGA, and localized accumulation of UL1 mRNA ( Fig 1 and S7A Fig). These features lead us to hypothesize that interactions of the UL with myocytes or the ECM on this anterior side may serve to promote stemness. In line with this hypothesis, the division of midline progenitors perpendicular to the UL axis may allow only one of the daughters to remain in contact with this presumed anterior niche. It is important to point out that our model of heart growth is primarily based on data from juvenile hearts. Widespread mitosis within the adult myocardium (S2 Fig) suggests that differentiated myocardial cells may be capable of undergoing mitosis or dedifferentiating into precursors. Alternatively, a population of precursors may remain embedded in the myocardium. Although we did not observe any cells in the myocardium that exhibited distinctive morphological features indicative of a precursor, further studies are needed to address this point. The role of canonical Wnt signaling in heart growth Though our data indicate that canonical Wnt signaling suppresses UL proliferation, the underlying mechanisms remain to be explored. In particular, based on our scRNA-seq data we can begin investigating candidate receptors and ligands that mediate activation of this pathway. We are also interested in using characterized reporters of Wnt activity to determine the spatiotemporal pattern cWnt activation within different progenitor populations. Further studies will also focus on determining whether cWnt signaling acts directly to drive progenitors into a mitotic arrest or if it acts indirectly, for instance by specification of newly born progenitor daughter cells as a less proliferative cell type. In vertebrate embryos, Wnt-dependent contributions to heart development are complex and vary depending on the model organism, impacted cell population and developmental stage ( 148 – 150 ). A more coherent understanding regarding the role of cWnt signaling in C. robusta heart growth will allow productive comparisons with vertebrate cardiogenesis, potentially delineating specific functional similarities. One of the most exciting results of this part of our study was that inhibition of cWnt signaling led to an overall increase in heart size without apparently impacting heart morphology. Thus, we anticipate that insights into the role of cWnt signals in C. robusta heart growth will inform a broader understanding of proportional organ growth. A complex range of neural and neural-like cell types are associated with the C. robusta heart Our data suggest that the C. robusta heart is innervated by extrinsic neurons originating from the central ganglia that connect with two previously described distal intrinsic neural plexuses located at either end of the heart ( Fig 7 ). It was previously thought that PC2 serves as a pan-neuronal reporter for these cardiac plexuses. However, we have identified additional cardiac neural-like populations associated with these plexuses including some that do not co-express PC2 ( Fig 6 and S9 Fig). Additionally, we have observed striking differences in composition of the ventral vs. the dorsal plexus (S8, S9). Our data also suggests that these plexuses consist of intrinsic cardiac neurons presumably ectodermal in origin along with mesoderm-derived pacemaker cells produced from a common cardiomyogenic progenitor, as seen in vertebrate embryos ( Fig 7 ) ( 135 , 136 ). Future work will focus on more thoroughly distinguishing each of these distinct cardiac neural-like cell populations through the use of lineage-specific reporters. These reporters would also allow us to sort and re-sequence distinct cardiac neural-like cell populations along with tracing their distinct developmental origins. Additionally, these reporters could be used to drive optogenetic construct expression in central ganglion neurons or different cardiac neural-like cell types, permitting functional analysis regarding the contributions of these different neural cell types to cardiac function or growth. Neural control of progenitor cells proliferation and organ growth The most striking finding in this study relates to the presumed role of intrinsic neurons or pacemaker cells in promoting cardiomyocyte progenitor proliferation ( Fig 7K ). Although our studies strongly support a key role for tachykinin signaling in neural-dependent heart growth, many key aspects of our model remain ambiguous. One major gap relates to the specific identity of cells that respond to tachykinin signaling. Characterization of cell-type specific regulatory elements would allow us to perform targeted CRISPR-Cas9 knockdown of TACR in distinct populations of cardiac neural-like cell types and identify which cells mediate tachykinin-dependent cardiac progenitor proliferation. Another major gap relates to the identity of the proliferative signal produced by cardiac neural-like cells. Based on signaling factor expression in the scRNA-seq cardiac progenitor cluster ( Fig 2F ), we are particularly interested in investigating potential downstream roles for the RA, EGF, Insulin, Notch and TGF-beta pathways. Activity sensors for a range of developmental signaling pathways could be deployed to assess TK-dependent activation or inactivation of these pathways in distal progenitor cells. We are also interested in investigating whether TK signaling instructs neural-like cells to secrete cWnt inhibitors. It is also possible that TK promotes proliferation indirectly through regulation of peristaltic rate. This hypothesis could be tested by paralysing the juvenile heart and examining whether this impacts progenitor proliferation. Lastly, a role for neural signaling in cardiomyocyte progenitor proliferation suggests that environmental or physiological conditions could mediate changes in heart growth. This hypothesis could be tested by examining the impact of nutritional availability or other external factors on heart growth and determining whether any such impacts require extrinsic cardiac innervation. Though many studies have revealed roles for innervation in promoting organ development, few studies focus on the impact of innervation on progenitor or stem cell proliferation( 15 , 16 , 18 ). Literature on potential roles for cardiac innervation or neuropeptides in progenitor proliferation generally focus on post-natal growth or injury ( 22 , 80 , 138 , 151 , 152 ). Thus, insights gained from further exploration of heart formation and growth in C. robusta may inform efforts to better understand the role of tachykinin and other neural signals in vertebrate heart growth. Methods EdU EdU staining was performed using the manufacturer’s protocol (Invitrogen Click-iT®). Briefly, juveniles or adult heart tissue was incubated in filtered artificial seawater (FSW) and the EdU staining solution in 35mm dishes and reared at 18°C for the desired amount of time (between 30 min and 24 hours). Samples were rinsed 3 times with FSW and then either pulled for fixation or left for a pulse phase. EdU-incorporated samples were fixed in 4% PFA in FSW overnight at 4°C, rinsed twice with PBS, and incubated in the Click-iT® reaction buffer for 30min. Samples were then rinsed twice with PBS, incubated in 1X DAPI/PBS for 20 min at room temperature, then rinsed in PBS prior to mounting. Phalloidin and DAPI Staining Juveniles were first relaxed by adding a small (∼3mm length) crystal of menthol directly to a 60 mm dish of FSW for 15-30 min and were monitored until unresponsive. Juveniles were then fixed using 4% PFA in seawater for 15 minutes at room temperature. Juveniles were then washed twice with PBS. To permeabilize, juveniles were incubated for 1 hr in 0.5% Triton-X in PBS while rotating on a nutator. Following permeabilization, juveniles were incubated in DAPI (Thermofisher 62248) (1:1000) and Phalloidin (1:100) overnight at 4°C on a nutator covered in foil. Juveniles were then incubated for 5 min twice with PBS. Juveniles were mounted on slides in PBS using coverslip spacers. Electroporation protocol Gravid Ciona robusta were obtained from MREP and Marinus Scientific, collected from various locations in southern California. Animals were maintained under constant illumination in a recirculating refrigerated tank to promote gamete accumulation. For electroporation, gametes from a minimum of three animals were combined. Transgene DNA constructs in water were pooled and mixed to a final concentration of 0.77 M D-mannitol. DNA concentrations were kept at or below 125 ug/mL and performed as in ( 144 ). Electroporated embryos were subsequently cultured at 18°C in FSW supplemented with 10 U/mL penicillin and 10 μg/mL streptomycin (FSW+pen/strep) in gelatin-coated dishes. Juvenile growth Protocols for fertilization, dechorionation, and electroporation were carried out as described. At ∼21 hours post fertilization (post-hatching stage), larvae were moved into scratched dishes and incubated at 18°C. Two days after settlement, metamorphosed juveniles were moved into culturing aquaria. Approximately 2L of seawater from our adult C. robusta holding tank was added to 2L of FSW. A small 5V USB pump was added for circulation. Scratched dishes containing the D3 juveniles were transferred to the 4L aquarium and kept at 18°C. The scratch dishes are engineered to hang vertically within the aquarium using Styrofoam secured to one end and an Eppendorf weighted down with glass beads secured to the opposite end. Aquaria were covered with foil to prevent evaporation. Every 2 days, 2L of water was replaced with FSW, and juveniles were fed 0.5 mL MicroVert (Kent Marine) and 0.5 mL Roti-Feast (Reef Nutrition). Adult heart in vivo pharmacological inhibition Glass needles were prepared for injection by pulling capillary tubes with a needle puller. Either 25 µL of Aprepitant (Sigma SML2215), Atropine (Cayman 12008), or molecular grade DMSO plus Fast Green dye was loaded into the needle. The final working concentration was 20µM for Aprepitant and 50 µM for atropine. Micro-dissection scissors were used to make a short and shallow incision in the tunic superior to the heart. The epicardial layer and longitudinal muscles that surround the heart and stomach were kept intact. Only if Fast Green dye was visible in the interior of the heart was an injection considered successful. Injected animals were then incubated overnight in a 400 mL container with FSW+ Pen/strep at 18° C. Hearts were then dissected out and were incubated with EdU for 24 hours before fixation. Hearts were stained with EdU Click-iT® kit (Invitrogen C10637) according to manufacturer’s protocol and co-stained for DAPI. EdU- and DAPI-stained nuclei were quantified using a custom pipeline in CellProfiler ( 153 , 154 ). Single cell sequencing Entire hearts were dissected in dishes in ice-cold FSW in 100 mm dishes. Micro-dissection scissors were used to cut hearts into ∼2 mm 2 pieces of tissue. All tissue pieces were transferred to an Eppendorf tube using forceps to minimize transfer of excess FSW. 100ul of Papain solution (Thermofisher 88285E) + 1mg/ml collagenase (Gibco 17100-017) + 0.5mg/mL cellulase (Thermofisher J64019) (kept on ice prior to use) was added to tissue, and subsequently left at room temperature for a 30 min incubation with occasional mixing. 15-minute trituration was performed using a P1000 pipette set to 1000 µL, using slow deliberate pipetting at room temperature. A cell mesh filter was used to strain out clumps of cells and debris (Falcon 352235). 20% Fetal Bovine Serum was added to ice cold magnesium and calcium-free FSW to inhibit the protease. Samples were then spun at 800 x g (∼2800 RPM) for 3 min at 4°C. Cells were washed with ice cold Mg/Ca2+ free FSW twice and resuspended in 100uL magnesium and calcium-free FSW. A hemocytometer and trypan blue staining was used to count cells and assess cell death percentage. Cells were diluted to 700 cells/uL using magnesium and calcium-free FSW. Cells were then kept on ice until 3p Capture and library preparation by the University of Pennsylvania Next-Generation Sequencing Core. Two replicates were performed. 3p Library Prep was performed by the University of Pennsylvania Next-Generation Sequencing Core and sequenced first on a MiSeq, and then a HiSeq 2000 resulting in 195M reads for ∼8000 captured cells. Kaede Photoconversion Electroporation was carried out with standard protocol using 100 µg of Mesp>Kaede per electroporation. Embryos were kept in the dark using foil. Juveniles were grown according to the juvenile culturing protocol until D5. FSW+menthol was prepared by adding menthol crystals up into the 5mL line of a 50 mL falcon tube and then filled to the 50 mL line with FSW, incubated for 30 minutes, and then placed on ice. Individual juveniles were placed in a small imaging dish with the FSW+menthol. A Zeiss LSM confocal was used for photoconversion and imaging using 40x. The stage was cooled to 4°C. Zeiss Smart Setup was used to select Kaede-green and Kaede-red and transmitted light (PMT-T). A range of ∼1-10% laser for 488 nm and 5-10% 568 nm laser as used with 650 V gain for the fluorescent PMTs, and ∼150-230 V gain for the transmitted light PMT. Photobleaching using the ROI tool in Zen was used, with the 405nm laser for conversion and set to 5%. Cells were converted until the green signal was no longer visible (∼10-30 seconds). A 96 well plate was then used to culture individual juveniles (1 juvenile/well) after washing juveniles with fresh FSW+pen/strep (Gibco 15070-063). Animals were fed and imaged once every 24 hours for several days after initial photoconversion. Single Cell Heart Sequencing Analysis Genomics single-cell RNA sequencing. Reads were aligned to the Ciona intestinalis ( C. robusta) Ciona3 genome assembly using gene annotations from the KH2012 gene models. Gene expression matrices were generated using the Cell Ranger count pipeline with default parameters and further processed using Scanpy ( 155 ). Cells expressing fewer than 400 genes were filtered out. The data were then normalized to a target sum of 1e4 per cell and log-transformed using the log1p function. Technical effects were regressed out using regress_out, and data were subsequently scaled to unit variance with a maximum value of 10. Principal component analysis (PCA) was performed using the ARPACK solver. The log variance ratio was used to determine the number of informative principal components. A nearest neighbour graph was computed, followed by UMAP dimensionality reduction. Louvain clustering was performed on the neighbourhood graph to identify transcriptional clusters. Two biological replicates were included and ingested together into the final displayed UMAP( 155 ). Single-cell Blood analysis Preprocessing A standard Scanpy pre-processing and clustering pipeline was used for each single cell dataset. First quality control metrics were calculated to determine threshold values. An appropriate threshold was set to eliminate cells that did not meet the minimum number of genes. This threshold was dynamically set as determined by the depth and calculated distribution of the number of genes detected per cell, however across all datasets a minimum of >200 genes/cell was required. Scatterplots of the number of genes by counts and total counts were generated to ensure dataset quality after the filtering step. Data were normalized to a target sum of 1e4 and then log-transformed to reduce variability of gene expression. Logₑ(1 + x) was used, where x is the normalized count. Regression was then applied to the data to reduce technical variability and batch effects across sequencing libraries. Finally, the data were scaled to a max value of 10 for intuitive comparison of gene expression. Dimensionality reduction Principal Component Analysis (PCA) analysis was performed individually on the normalized datasets. The variance ratio was calculated and plotted to determine the ranking and impact each principal component contributed to the variance in the dataset, which was then used to determine the number of principal components to include for subsequent nearest neighbour analysis. Uniform Manifold Approximation and Projection (UMAP) was then used to visualize the sequencing datasets to visualize local and global structures in the datasets. Standard Louvain clustering was then performed on the UMAP. Manual inspection of known marker genes was used as a quality control-step of Louvain and UMAP settings for stages where known markers are well established in the literature (see references therein). Elimination of blood clusters To identify blood cells, we compared to published scRNA-seq data of adult C. robusta circulating cells ( 156 ). We identified cell clusters in the heart dataset which were transcriptomically similar to the blood dataset’s published Leiden clusters. Specifically, for every pair of heart and blood dataset cluster pair, we calculated the correlation across highly variable genes of the mean log-transformed expression within each cluster. Cluster pairs with a correlation above a threshold of 0.5 are considered transcriptomically similar, and the heart clusters of those pairs were labeled as blood cells. This correlation threshold was chosen by running an equivalent analysis to compare biological replicates in the blood dataset, picking a threshold that identified cell states represented in both replicates. Two manual adjustments were then made to the set of clusters which passed this threshold: ( 1 ) One cluster pairing, matching heart cluster 6 to blood cluster 10, has a correlation just under the0.5 threshold (correlation=0.494). Manual inspection of marker genes showed significant overlap, so we also labeled this heart cluster as blood cells. ( 2 ) One rare blood cell cluster, cluster 31 (0.17% of the observed blood cells) has a high correlation to heart clusters 1 and 8 (together, 12.1% of the observed heart cells). Given the large discrepancy in cell abundance, we did not label heart clusters 1 and 8 as blood cells. The final set of heart clusters matched to blood cell states is listed in Table Sx [attached csv, also plotted in attached png]. Identification of marker genes and genes of interest Potential marker genes for adult C. robusta heart clusters were identified through a variety of methodologies to converge into a final list. First known transcription factors were assessed for enrichment across clusters and then manually evaluated for biological relevance in each cluster. A Wilcoxon Rank-Sum Test was performed across clusters to identify genes of interest and potential marker genes in an unbiased manner, and then manually curated to highlight relevant genes. In addition, the top highest expressed genes were determined for each cluster and then manually selected depending on specificity. Lastly, specific and low-expressed genes were also identified for each cluster to identify any potentially important genes not identified through the other analyses. Known marker genes for cell types in C. robusta and across other species were used to begin to glean the cell type identity across the various heart clusters. Sub-clustering of adult C. robusta heart clusters After reporter and FISH analysis of various C. robusta stages (see section on cluster validation), the TACR cluster and UL clusters were further processed for analysis of sub-types of cells within each of these broader clusters. The TACR and UL clusters were subset in Scanpy, and then Louvain clustering was performed again on the UL and TACR cluster individually. The previously calculated PCA value for all adult C. robusta clusters was re-used to preserve the structure of the individual TACR and UL clusters. The Louvain clustering was recalculated for each new cluster using a resolution 0.83. WMISH/FISH In situ hybridizations were performed as in ( 122 ), with slight modifications for fluorescence in situ detection: After juveniles were phased out of hybridization buffer, they were washed three times with TNT (0.1 M Tris–HCl pH 7.4, 0.15 M NaCl, 0.1% Tween-20), then blocked for 1 hour in TNB buffer (0.1 M Tris–HCl pH 7.4, 0.15 M NaCl, 1% BSA). Juveniles were subsequently incubated overnight at 4 °C with a 1:1000 dilution of POD-conjugated anti-DIG primary antibody (Roche). Following antibody incubation, embryos were washed three times with TNT and exposed for 5 minutes to FITC-tyramide working solution (Perkin Elmer) to fluorescently label antisense RNA probes. Juveniles were incubated in TNT for 10 minutes three times prior to mounting. Molecular cloning and reporter transgene information Syt15>GFP was generated by PCR-amplifying approximately 1500 bp of DNA upstream from the start of transcription using the forward oligo GGTGTTAAATTATCCACGATAAACCG and the reverse oligo TATCGTAGTATAACAACGACAAACTTGG. This fragment was subcloned into Mesp>GFP ( 97 ) after restriction digest removal of the Mesp enhancer fragment. Mesp>Kaede::NLS ( Mesp>Kaede ) was generously provided by Alberto Stolfi. Mesp>RFP was previously described in ( 157 ). Tg[MiCiPC2K]2, Tg[MiCiPC2K]3, Tg[MiCiTnIG]2, Tg[MiCiTnIG]3, Tg[MiCib2TBC]3, Tg[MiCib2TBC]4, Tg[MiCiVACHTC]2 , were provided by Yasunori Sasakura ( 141 , 158 – 161 ), and pCiTachykinint;CF from Honoo Satake. VACHT>eCFP cassette ( 162 ) of pSPCiVACHTC ( 163 ) was amplified by PCR using with PrimeSTAR HS DNA polymerase (Takara Bio Japan). The fragment was inserted into the BamHI site of pMiLRneo ( 164 ) by In-Fusion HD Cloning kit (Clontech) to create pMiCiVACHTC . pMiCiVACHTC was electroporated with Minos transposase mRNA to create stable transgenic line according to the previous method ( 165 ). FoxfΔepi>dnTCF Dominant-negative TCF (dnTCF), the C. robusta TCF coding sequence which lacks the N-terminus, was generated by PCR amplification of genomic DNA with the oligos CACCATGTACGATGTTCCGGCAAAAGTA and GCTGATGTTGCACGGCGG, as in ( 123 ). FoxfΔepi>dnTCF was constructed by subcloning the dnTCF sequence into our FoxfΔepi vector (described in ( 122 )) after restriction-digest removal of the LacZ sequence. Constructs for CRISPR-Cas9 targeting of Tachykinin and Tachykinin Receptor To generate PC2>Cas9 , we obtained mMiCiPC2K from CITRES ( https://marinebio.nbrp.jp/ciona/top/top.jsp ). The MESP>Cas9 vector was digested using AscI and NotI. The mMiCiPC2K enhancer was PCR amplified with added overhangs matching the Mesp>Cas9 vector backbone using ACGTATTAATTAAGGCGCGTAACACCACGATATTAAATC and GGCTAGCCATGGTTGCGGCCCATTCAAATAAAATGCTGCT. The following sgRNA targeting sequences were obtained by using CRISPOR ( 166 ) to maximize target efficiency and low off-target predictions. Kinased and annealed sgRNA oligos were cloned into an empty U6>sgRNA vector after digestion with BsaI as previously described in( 167 ). The sgRNA sequences used for targeting Tachykinin were TTGATGGGAAAACGATCAAT ( TKsgRNA2 ), and CATCGTTCACTGGCTTGATG ( TKsgRNA3 ). The sgRNA sequences used for targeting Tachykinin Receptor ( TACR ) were AGTAACAAGAGACGTGGGAC ( TACRsgRNA2 ) and CGTATGGGCTCTGGGCGAAC ( TACRsgRNA3 ). The control sgRNA used targeted GFP, as in ( 122 ). Electroporation was carried out using 25 µg PC2>Cas9 or Mesp>Cas9 , 25 µg of each TK or TACR sgRNA and 25 µg PC2>Kaede resulting in 100 µg total DNA per electroporation. Mesp>Cas9 and the U6>sgRNA constructs were generously provided by Lionel Christiaen (described in ( 142 )). Pharmacological Inhibition of Tachykinin Signaling Immediately after metamorphosis, D3 juveniles were subjected to continuous pharmacological treatment until D7. In separate dishes, juveniles were either grown in 100 μΜ Aprepitant, 1 µM Spantide II (Sigma SCP0241), or control DMSO (Invitrogen C10337) for 4 days, followed by a 6 hr EdU pulse after the drug treatment. Juveniles were fixed and stained according to the above protocols and UL counts were assessed using FIJI( 168 ). Pharmacological Inhibition of Neurotransmitters in Juveniles Juveniles were cultured according to protocol. Drugs were at the following concentrations: 100 μΜ AMPT, 9.13μΜ doxazosin mesylate, or 50 µM for atropine. Atropine, DMSO, or doxazosin mesylate was added to D3 juveniles for 4 days and UL length was counted in D7 animals after staining with DAPI and subsequent confocal imaging. AMPT or FSW control was added to D6 juveniles for 24 hours. Animals were fixed and stained with DAPI then imaged. UL counting was performed using FIJI. Pharmacological inhibition of the cWnt pathway IWR-1-endo was dissolved in DMSO and applied at a final concentration of 2.5µM. BIO (6-Bromoindirubin-3′-oxime (Sigma B1686); ( 120 ) was dissolved in DMSO and applied at a final concentration of 2.5uM. Statistical analysis For statistical analysis of CRISPR of TK and TACR juveniles, a Fisher’s Exact Test was performed using scipy.stats.fisher_exact. Analysis of normalized UL counts for Aprepitant, Spantide-II, Atropine, AMPT, and doxazosin mesylate was performed by first normalizing each trial by the counts for an animal in the DMSO condition, and then taking the average normalized UL count of each trial. A t-test was performed on the normalized trial values to obtain a p-value using excel. A t-test was also performed using excel on the UL EdU anatomical region (ventral, middle, dorsal) measuring the average percent of animals per each trial that had one or more EdU positive cells per anatomical region of UL. For the adult heart growth by region, and for the injection assay of DMSO, Aprepitant, or Atropine, first the number of nuclei were counted in total using CellProfiler, then the number of EdU nuclei were counted to give a percent of cells that are EdU positive. A t-test in excel was performed comparing the percent of EdU+ cells per adult heart for each anatomical region (distal, middle, apical) or drug (DMSO, Aprepitant, Atropine). A paired t-test was used on heart rate data before and after each drug treatment on individual juveniles for Aprepitant, Acetylcholine, and AMPT using scipy.stats.ttest_rel in Python. Contributions B.D, H.N.G, and J.Y.B. conceived of the project. B.D., H.N.G, and C.J.P. designed experiments. J.Y.B. and H.N.G. and G.B. performed heart microanatomy and proliferation experiments. C.J.P, R.G. and M.G. performed Wnt experiments. J.Y.B. performed and analysed pulse-chase juvenile experiments and C.J.P. interpreted data. T.S. performed blood cell analysis with A.M.K input. C.J.P. generated heart reporter and all FISH data. H.N.G and A.H. documented neural reporter expression, and Y.S. provided feedback for neural reporters. T.O. and H.S. generated tachykinin reporter. H.N.G. performed and analysed heart single cell experiments, tachykinin CRISPR, cloned PC2-Cas9, inhibition of tachykinin signalling in juvenile and adult, and statistical analysis. C.J.P. and H.N.G. performed Kaede photoconversion experiments. S.N. performed statistical analysis of Wnt data. K.M. performed adrenergic experiments and collected developmental juvenile time course data. K.L., H.N.G, O.C., G.L., S.Y.C, and T.G. performed adult heart atropine experiment. K.L. and H.N.G performed atropine juvenile studies. M.C.D. and C.J.P. performed TH inhibition experiments. E.D.G. performed Phox2b experiments. B.D., H.N.G., and C.J.P. wrote the paper. Funding This work was supported by two grants received by BD, and the F32 fellowship (1F32HL170997) supporting H.N.G. The American Heart Association grant number 20AIREA35080013 ( https://professional.heart.org/en/research-programs/aha-funding-opportunities ) along with the National Science Foundation grant number 8077804 ( https://www.nsf.gov/ ). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Download figure Open in new tab S1 Fig. Microanatomy of C. robusta and C. savignyi adult hearts. (A) Cross-section (X-Y) view of adult C. robusta heart. (B) Cross-section (Y-Z) view of C. robusta adult heart. (C-F) C. savignyi cell types as labeled, arrows in ( F ) indicate the UL. Middle panels are magnified areas from the left panels, right panels are magnified areas from middle panels. (G) Previously unidentified ordered pericardial, F-actin-rich fascicles spanning the heart from posterior (distal) to anterior (apex) on the pericardial layer. These fibers were only prominent in young adult C. robusta hearts. In all images cyan represents DAPI staining and magenta represents phalloidin staining. Download figure Open in new tab S2 Fig. Regional cell division frequency in the adult heart. (A) Graph displaying mean proliferation index (% dividing cells) in distal, middle and apical regions of the heart as assayed by 24-hour pulses of EdU, N=15 adult hearts. Error bars represent the S.E.M., and p-value calculated from t-test. (B-C) Representative images of distal and midline UL proliferation respectively as assayed by a 6-hour pulse of EdU (magenta), DAPI stained nuclei (cyan). Download figure Open in new tab S3 Fig. Juvenile midline UL and myocardial growth rates. (A) Quantity of myocyte (blue) and midline UL cells (orange) from D3 to D15 using standard culturing conditions. (B) Quantity of myocyte (blue) and midline UL cells (orange) from D3 to D8 days using optimized culturing conditions (see Methods section). In A and B, D3-D8 along the x-axis are highlighted for comparison. Orange dots and lines, number of UL cells per time point. Download figure Open in new tab S4 Fig. Lineage tracing of myocardial progenitors in the UL using Kaede conversion. In all panels, photoconverted cells (yellow) are tracked over multiple days in living juveniles starting with photoconversion at D5, timepoints indicate hours after conversion. See text for details. (A) Blue arrows track a single labeled midline UL cell that appears to divide symmetrically to produce two midline UL daughters. (B, C) Red arrows track single labeled midline UL cells that appears to divide asymmetrically to produce a midline UL daughter that remains in the same confocal plane as the rest of the UL, bottom row, along with a presumptive myocardial precursor that moves into a different confocal plane (top row). (D) Red arrows track a single labeled presumptive myocardial precursor that appears to migrate anteriorly, away from the UL. (E) Red arrows track a single labeled presumptive myocardial precursor that appears to mature over a 72-hr time-course as evidenced by elongation of the nucleus. Download figure Open in new tab S5 Figure. Pearson’s correlation coefficient (PCC) analysis to assess cluster similarities. PCC suggests Cluster 2 correlates strongly with Cluster 3 while Cluster 1 correlates strongly with Cluster 7. Overlapping expression patterns between these two clusters may contribute to the lack of uniquely enriched genes in Cluster 1 (Main text Figure 3A , B). Additionally, PCC revealed Cluster 8 did not correlate strongly with any other cluster suggesting this cluster is transcriptionally unique. Download figure Open in new tab S6 Figure. Synaptotagmin 15 reporter expression in the epicardium of a D15 juvenile. Mesp>RFP labels pericardium, myocardium, and the UL (magenta). Syt15>GFP detected in the overlying epicardium (green). Download figure Open in new tab S7 Figure. Expression patterns of marker genes associated with the presumptive UL and cardiac neural-like clusters. (A) CrUL1 expression in a D5 heart. Colorimetric in situ hybridization (left), FISH (middle), and cartoon schematic (right). (B) Gata.a expression in a D8 heart. FISH (left) and cartoon (right). Note that in the micrograph a Z-plane containing myocardial cells in the anterior region of the heart is shown on the left while the Z-plane containing the UL in the posterior region of the heart is shown on the right. (C) HCN2/3/4 expression in a D5 heart. FISH (left) and cartoon depiction (right). Blue (A and B) or cyan (C) represents DAPI staining. Yellow (A and B) or magenta (C) represents probe detection for each transcript. All images shown anterior to the left and dorsal up. Download figure Open in new tab S8 Figure. Location of cardiac neural-like/pacemaker cells in the adult C. robusta heart. (A) Summary diagram of observed localization patterns for PC2+, VACHT+, and TH+ neural-like cells in the distal plexus of the adult heart. Note the ventral-exclusive presence of TH (green. (B) PC2>Kaede expression. (C) VACHT>CFP expression. ( D,D’, D’’ ) VACHT>CFP and Troponin>RFP expression in a young D30 adult as indicated. D’’ shows magnified regions from D’. (E) VACHT>CFP and TH>Kaede double-labeled adult hearts. Left column is VACHT>CFP (blue), middle column is TH>Kaede (green), and right column is merged. Middle and bottom rows correspond to enlarged areas of two different Z-planes of the boxed area in the top row. (F) TH>Kaede reporter expression at the distal end of the ventral plexus. Top panel is Kaede fluorescence, middle panel is brightfield, bottom panel is merged. Line in F indicates where the myocardial tube ends relative to the TH+ neural-like ring. Download figure Open in new tab S9 Figure. Location of cardiac neural-like/pacemaker cells in the juvenile heart. (A) VACHT>CFP (red) in live D3 juvenile heart. (B) PC2>Kaede (red) and VACHT>CFP (blue) in a fixed D4 juvenile heart, outlined in yellow. (C) VACHT>CFP (red) in live D5 juvenile heart. (D) VACHT>CFP (blue) and TH>Kaede (yellow) in a fixed D5 juvenile heart, outlined in white. (E) TH>Kaede in a live D7 juvenile heart. (F) TH>Kaede (magenta) in a fixed juvenile heart. DAPI staining in cyan. F’ shows a single Z-plane from F, note the overlap in magenta and cyan that appears to be associated with a UL cell (arrow). (G) DAPI stained nuclei of a D10 transgenic PC2>Kaede juvenile heart. G’ shows PC2>Kaede expression. G’’ shows merged view. Note PC2> Kaede expression at the dorsal and ventral ends of the UL. (H) PC2>Kaede expression (red) in a D12 juvenile heart. Note staining at the dorsal and ventral ends of the UL as well as expression along the UL (arrow and dotted line). In A, C, E, and H, fluorescence merged with brightfield. Download figure Open in new tab S10 Figure. Identification of presumptive pacemaker cells that co-express Phox2b and Mesp reporters. (A-D) Juvenile MESP>H2B:mCh (cyan) Phox2b>Unc76::GFP (yellow) heart stained with phalloidin (magenta). The only Phox2b+ cell shown in this substack is also Mesp+ (arrowhead). (E-H) Another juvenile MESP>H2B:mCh (cyan) Phox2b>Unc76::GFP (yellow) heart stained with phalloidin (magenta). Of the two Phox2b+ cells shown in this substack, one is Mesp- (arrow) while the other is Mesp+ (arrowhead). Note that the morphology of the Mesp+ cells is distinct from that of the Mesp-cell. A total of 9 transgenic hearts were examined and these are the only two double positive cells observed among a total of 37 Phox2b+ cells that were detected in these hearts. Download figure Open in new tab S11 Figure. The impact of neural signaling modulators on heart rate and UL cell numbers. (A) Violin plots of recorded heart rates in response to acetylcholine chloride (top), AMPT (middle), and aprepitant (bottom). (B) PC2>Kaede (magenta) labels cells that are interspersed with densely clustered DAPI stained nuclei (cyan) within the distal UL (outlined in white) in an adult heart. (C) D7 control heart. (D) D7 heart treated with Atropine. (E) D7 control heart. (F) D7 heart treated with doxazosin mesylate. (G) D7 control heart. (H) D7 heart treated with AMPT. (I) Violin plot of normalized UL counts per treatment. Counts normalized to the DMSO control for each trial. In C-H, cyan represents DAPI stained nuclei. Images in E-H also show phalloidin staining (magenta). For I, a t-test was performed on normalized data, averaged across trials. Doxazosin Mesylate: N= 60 control and 58 experimental samples, 2 trials. Atropine: N=13 control and 14 experimental samples, 2 trials. AMPT: N= 46 control, 33 experimental samples, 2 trials. Download figure Open in new tab S12 Figure. Fluorescent in situ hybridization of TACR expression. Blue represents DAPI and magenta displays fluorescent probe detection (left). Cartoon model of expression pattern (right), UL cells outlined in blue, myocardial cells outlined in red, green cells represent presumptive neural-like/pacemaker cells. Download figure Open in new tab S13 Figure. Tachykinin reporter and scRNA-seq analysis confirms lack of Cr-TK expression in the heart. (A) TK>CFP reporter electroporation expression in the central ganglion (red). (B) Magnified region of the heart. Note absence of reporter expression. (C) UMAP showing Cr-TK expression levels, note absence of expression. S1 Movie. Control D10 juvenile exhibiting stereotypical peristalsis. Juvenile expressing PC2>Cas9 and a GFP-targeting sgRNA. S2 Movie. Experimental D10 juvenile exhibiting a pericardial bubble absent a prominent inner myocardial tube. Juvenile expressing PC2>Cas9 and a pair of TK -targeting sgRNAs. S3 Movie. Control D10 juvenile exhibiting stereotypical peristalsis. Juvenile expressing Mesp>Cas9 and a GFP-targeting sgRNA. S4 Movie. Experimental D10 juvenile showing dramatically disrupted myocardial tube along with abnormal peristalsis. Juvenile expressing Mesp>Cas9 and a pair of TACR -targeting sgRNAs. Acknowledgements The neural transgenic lines were supported by the National BioResource Project, Japan. The authors would like to thank Alberto Stolfi for his feedback. H.N.G. thanks M.B. for their support. 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