Abstract
Muscle-based movement is a hallmark of animal biology, but the evolutionary origins of myocytes are unknown. Although believed to lack muscles, sponges (Porifera) are capable of coordinated whole-body contractions that purge debris from internal water canals. This behavior has been observed for decades, but their contractile tissues remain uncharacterized with respect to their ultrastructure, regulation, and development. We examine the sponge Ephydatia muelleri and find tissue-wide organization of a contractile module composed of actin, striated-muscle myosin II, and transgelin, and that contractions are regulated by the release of internal Ca2+ stores upstream of the myosin-light-chain-kinase (MLCK) pathway. The development of this contractile module appears to involve myocardin-related transcription factor (MRTF) as part of an environmentally inducible transcriptional complex that also functions in muscle development, plasticity, and regeneration. As an actin-regulated force-sensor, MRTF-activity offers a mechanism for how the contractile tissues that line water canals can dynamically remodel in response to flow and can re-form normally from stem-cells in the absence of the intrinsic spatial cues typical of animal embryogenesis. We conclude that the contractile module of sponge tissues shares elements of homology with contractile tissues in other animals, including muscles, indicating descent from a common, multifunctional tissue in the animal stem-lineage.
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Introduction
Animal movement is predominantly enabled by the coordinated activity of two cell types: myocytes (contractile cells of the muscular system) and neurons (signaling cells of the nervous system). But not all animals have these cell types—sponges lack both yet are still capable of coordinated tissue movements during whole-body contractions. Understanding how sponges contract has key relevance for understanding the evolutionary origins of animal movement.
Strictly sessile, sponges must actively pump water through an internal canal system for feeding, gas exchange, waste removal, and sexual reproduction. The canal system is partitioned into incurrent cavities (atria) and canals, feeding chambers lined by flagellated cells (choanocytes) that generate direction flow and phagocytose bacteria, and excurrent canals that channel wastewater to exhalant openings (oscula) (Fig. 1A). Contractions serve to clear canal blockages and maintain water flow. At the initiation of a contraction, incurrent pores close and canals narrow, and internal water pressure increases to dislodge debris1 (Fig. 1A).
Lab-grown freshwater sponges are useful models for studying contractions because they are small, transparent, and amenable to microscopy. In Ephydatia muelleri, contraction cycles can be described as biphasic. During phase I, atrial volume decreases, and incurrent canals narrow as water is displaced into excurrent canals which expand in diameter. During phase II, atrial volume and incurrent canal diameter increase as excurrent canals narrow (Fig. 1B). This is a progressive sequence, leading from the incurrent tissues of the system to the excurrent tissues, and propagates towards oscula1.
From a signaling perspective, it is known that canal blockage (natural or induced by the addition of Sumi ink) activates ciliated sensory cells, which release nitric oxide (NO). Nitric oxide diffuses across tissues and is modulated by glutamate and gamma-aminobutric acid (GAGA), initiating contraction2,3,4. A major gap in this model is that the cellular mechanisms of the contractile response remain essentially uncharacterized. How is the contractile force generated structurally, and how are contractions regulated at the level of cellular physiology? Answers to these questions are needed to clarify whether sponge contractile tissues are similar to non-muscle contractile tissues in other animals such as epithelia that undergo apical constriction, or to muscles?
In well-studied bilaterian animal models with muscle tissues, fast-contracting somatic myocytes are involved in voluntary or reactive movements, and slow-contracting visceral myocytes are involved in organ movements and tissue tension5. In both, contractions require interactions between actin filaments and type-II myosin, composed of two myosin heavy chains (MyHCs), two regulatory light chains (RLCs), and two essential light chains. The contraction speed of different cell and tissue types reflects the kinetics of the MyHC expressed. For example, striated-muscle myosin heavy chain (stMyHC) is found in fast-contracting skeletal and cardiac muscles of vertebrates, and smooth and striated invertebrate muscles6,7,8 (Fig. 2B). Non-muscle myosin heavy chain (nmMyHC) is found in slow-contracting vertebrate smooth muscle, and some invertebrate visceral muscles, but also in non-muscle contractile contexts such as cell motility, cytokinesis, and apical constriction9.
Bilaterian myocytes can be further distinguished by their regulatory mechanisms (see below). In fast contracting myocytes, the tropomyosin/troponin C complex hinders stMyHC from binding to actin but is released by Ca2+10. Troponin C-regulation is unique to bilaterian muscle. A potentially more ancient mechanism is the MLCK pathway, in which cytoplasmic Ca2+ binds to calmodulin, activating MLCK to phosphorylate the RLC of myosin II11,12. The MLCK pathway functions in muscle and non-muscle contractile contexts, and appears to regulate non-bilaterian muscles7,13.
During development, unique transcription factor combinations specify the identity of either cardiac and smooth muscles, or striated muscles in vertebrates (see below)5. These are well-conserved between species and indicate cell type homology between lineages14. An interaction central to the developmental specification of all muscle types is between a myocardin-related transcription factor (MRTF) and a MADS-box transcription factor—either serum response factor (SRF) or myocyte enhancer factor 2 (Mef2). Vertebrate smooth/cardiac muscle development also involves transcription factors from GATA, NK homeobox, and Fox families, and vertebrate striated/skeletal muscle development also involves E12 and MyoD transcription factors (restricted to bilaterians)7.
Here, we examine tissues of the sponge Ephydatia muelleri and find evidence of a contractile module with evolutionary links to muscle. Specifically, contractions depend upon the motor-activity of striated-muscle myosin II and are regulated by MLCK pathway, and the development of the contractile module appears to involve MRTF as part of a possible transcriptional complex that includes SRF and Fox-family transcription factors.
Results
Contractile actin-bundles in pinacocytes contain stMyHC
The tissues lining the atrial cavity and canals are believed to play a primary role in the contraction in sponges. These tissues are regionally differentiated but are all composed of endothelial-like cells (pinacocytes) that form a watertight barrier, express genes involved in contraction15,16, and decrease in area during contraction17. Moreover, in E. muelleri, pinacocytes contain linear actin-bundles that align between adjacent cells, exhibiting tissue-wide organization (Fig. 2A).
Searching the transcriptome18 we identified two type II myosin heavy chains, which clearly fall into stMyHC and nmMyHC groups (Fig. 2C). We generated and validated a custom antibody targeting a divergent region in the motor head for stMyHC (Fig. 2D, Supplementary Fig. 9). To test whether these actin-bundles are associated with myosin II, we immunostained for stMyHC (Fig. 2E). Staining was initially diffuse but increased throughout development. In fully differentiated tissues, stMyHC was organized into linear structures that resemble actin-bundle orientation (Fig. 2F), but co-staining was incompatible with fixation conditions. To further test for association with actin-bundles we treated sponges with latrunculin B, an inhibitor of actin polymerization, and observed that loss of stMyHC staining mirrored the dynamics of actin-bundle disassembly (Supplementary Fig. 1).
Contractions are regulated by the MLCK pathway
Previous studies have shown that sponge contractions abate in Ca2+/Mg2+-free medium (CMFM)1,19. To test the inverse of this, whether elevated cytoplasmic Ca2+ can induce contractions, we treated E. muelleri in CMFM with the calcium ionophore, ionomycin, and with the SERCA pump inhibitor, thapsigargin. Both treatments induced a strong biphasic contraction cycle (Fig. 3A). In ionomycin-treated sponges, contractions appeared to be more drawn-out and less intense compared with thapsigargin-treated sponges, which closely mirrored mechanically induced contractions at the tested concentrations. We interpret this as a difference in the kinetics of the molecules, as well as the mechanism of action; thapsigargin likely requires sufficient intracellular stores to elicit a response.
Since treatments were administered globally, the observed response could be a secondary effect downstream of NO signaling from activated sensory cells. To decouple sensation and contraction, we treated sponges with the NO-synthase inhibitor L-NAME (Fig. 3B). Sponges deficient for NO signaling did not contract in response to treatment with Sumi ink (which activates sensory cells) but exhibited a strong response to thapsigargin (Fig. 3C). This suggests that thapsigargin is acting on cells downstream of the sensory cells, which is consistent with the presence of endoplasmic reticulum-associated Ca2+ stores in contractile tissues.
With respect to the regulation of Ca2+ dependent contraction (Fig. 4A), E. muelleri has homologs of calmodulin, MLCK, and an RLC ortholog with conserved functional residues (Fig. 4B). If the release of internal Ca2+ is upstream of MLCK signaling, then contractions should be disrupted by the MLCK inhibitor ML-7. To test this, we first established a concentration of Sumi ink that permanently blocked water flow in L-NAME treated sponges (i.e., sponges deficient for NO synthesis), but could be cleared by contraction of untreated sponges. Ink clearance and the reestablishment of flow was determined by post-treating sponges with DiI to see if it entered canals. We found that ML-7-treated sponges were unable to clear ink and restore flow, nor could they be induced to contract with thapsigargin (Fig. 4C, D). Whereas ML-7 targets MLCK and is predicted to affect only a subset of myosin II activity, treatment with more general myosin ATPase inhibitors is expected to have global effects on myosin activity. We found that blebbistatin and para-amino blebbistatin caused a complex phenotype similar to phase I of the contraction cycle, but sponges never entered phase II and could not be induced to do so with thapsigargin (Supplementary Fig. 2).
In vertebrates, the RLC is the substrate for MLCK during smooth-muscle contraction20. To test for RLC phosphorylation during contraction, we immunostained sponges for phosphorylated RLC (pRLC) and saw staining of contractile bundles at the sponge surface (Fig. 4E). Their length was negatively correlated (R2 = 0.216, n = 118) with pRLC staining, indicating that increased phosphorylation of the RLC is associated with their contraction (Fig. 4F).
Collectively, these results support that contractions depend upon the release of ER-associated Ca2+ stores, downstream of NO signaling. Elevated Ca2+ then activates MLCK, leading to increased phosphorylation levels of RLC associated with contractile actin-bundles in pinacoderm tissues.
MRTF drives development of the contractile module
To test for conserved developmental mechanisms involved in specifying contractile cell-fate in sponges and bilaterians, we focused on the transcriptional cofactor, MRTF (Fig. 5A). In bilaterians, MRTF homologs are broadly expressed, but inhibited through the interaction between G-actin with N-terminal RPEL (RPxxEL) repeats21. Actin polymerization disrupts this interaction, exposing a nuclear localization signal and leading to nuclear translocation22,23. In the nucleus, MRTF interacts with SRF or Mef2 to drive the expression of contractile genes and can induce the differentiation of myocytes24,25,26,27 and myofibroblasts28, and is involved in the regulation of epithelial-mesenchyme transitions29,30. Loss of function studies in mice have shown that the loss of myocardin results in early embryonic lethality due to improper formation of vascular smooth muscle31. Loss of MRTF-B also results in early embryonic lethality due to cardiac defects32,33. Though most ubiquitously expressed, loss of MRTF-A is not lethal but results in deficiencies in lactation, likely resulting from loss of myoepithelial tissue34.
E. muelleri has a single, broadly expressed (Supplementary Fig. 3) MRTF ortholog that is predicted to have conserved RPEL repeats, suggesting that its activity may also be regulated by interactions with G-actin (Fig. 5B). Using a custom antibody, we found that this protein was predominantly cytoplasmic in undifferentiated archeocytes (stem cells), and nuclear in pinacocytes (Fig. 5C and Supplementary Fig. 4), where it presumably acts as a transcriptional co-factor. Known targets of MRTF regulation in bilaterians include the transgelin family (e.g., calponin, SM22alpha, MP20), which can be muscle-specific21,35. E. muelleri has three transgelin paralogs. Of these, we found that transgelin 2 (EmTAGLN2) was specifically localized to contractile bundles in pinacocytes (Fig. 5D and Supplementary Fig. 5).
Because MRTF activity is regulated by G-actin, it is possible to pharmacologically manipulate this interaction. Highly specific MRTF inhibitors such as CCG-20397136,37 have been identified, as well as activators that act through less specific mechanisms. One of these, N-cyclopropyl-5-(thiophen-2-yl)isoxazole-3-carboxamide (ISX) is an MRTF activator38,39,40 that drives cardiomyocyte differentiation in vivo38. ISX also drives the expression of secretory programs and neuronal differentiation through the transcription factor NeuroD1 in a Mef-2-dependent manner41,42,43. However, as sponges lack neurons and NeuroD1, we reasoned that it may act primarily on MRTF in sponges. To test this, we monitored EmTAGLN2 (a predicted MRTF-target) expression in response to CCG-207319 and ISX treatment and observed corollary changes; CCG-207319 treatment caused a decrease in EmTAGLN2 expression, whereas ISX treatment caused an increase in EmTAGLN2 expression (Fig. 5D). Since ISX is less specific than CCG-207319, we also corroborated this result by treatment with cytochalasin D—a potent MRTF-activator through competitive binding of G-actin44—and similarly observed an increase in EmTAGLN2 expression. From a functional perspective, MRTF-inhibited sponges had a reduced ability to clear Sumi ink—an indicator of diminished contractile activity (Fig. 5E).
To test the effects of MRTF-activation on contractile tissue differentiation, we dissociated juvenile sponges and treated archeocyte-enriched cell fractions with either DMSO or ISX and placed them in an attachment-free environment (this allows for the formation of primary aggregates—primmorphs—but delays differentiation). After three days, control primmorphs lacked contractile-bundles, whereas primmorphs treated with ISX developed linear actin-bundles that aligned at adhesion plaques (Fig. 6A). These stained positive for pRLC (Supplementary Fig. 6) and treated primmorphs contained elevated levels of stMyHC (Fig. 6B). Thapsigargin had no effect on control primmorphs but induced contractions in ISX-treated samples, which exhibited a 15.0 (+/−5.5)% reduction in cross-sectional area, followed by return to resting size (Fig. 6C).
MRTF activates myogenic factors, signaling, contractile, and adhesion genes
To understand the transcriptional response of primmorphs to MRTF activation, we sequenced mRNA from ISX or DMSO treated primmorphs. Differential expression analysis of 16,712 mapped transcripts revealed that 1390 were upregulated and 1091 were downregulated (Supplementary Figs. 7–8). We interpret upregulated genes as candidate targets of MRTF regulation, and downregulated genes as archeocyte-enriched. The ISX-treated samples had elevated expression of genes involved in contraction, signaling, development, and adhesion (Fig. 6D). Contraction-related genes included stMyHC (LogFC = 0.65, P = 0.038), which was validated by western blot (Fig. 6B), TAGLN2 (LogFC = 1.03, P = 1.0E−3), and components of the Ca2+-dependent MLCK pathway including MLCK-like serine/threonine kinase (LogFC = 1.58, P = 6.29E-5), calmodulin (LogFC = 1.55, P = 0.002), sodium/calcium exchanger 1 (SLC8A1) (LogFC = 1.05, P = 0.029), and Phospholipase C gamma (PLCγ) (LogFC = 1.01, P = 0.005) (Fig. 6D). Upregulated signaling genes included metabotropic glutamate receptors and GABA receptor subunits (Fig. 6D), consistent with a role for glutamatergic and GABAergic signaling in contractile behavior2,3. Upregulated developmental factors included the myogenic transcription factor SRF (LogFC = 1.18, P = 0.001). Four Forkhead transcription factors showed increased expression, including Fox-L2 (LogFC = 1.07, P = 0.008), FoxG (LogFC = 1.31, P = 4.16E−5), which is expressed in myocytes of invertebrates45, FoxO, and FoxJ1 (LogFC = 0.60, P = 0.016 and LogFC = 1.86, P = 0.001 respectively). The phylogenetically broad muscle marker Crip/Csrp46 was also upregulated (LogFC = 1.10, P = 0.001).
Some of the most highly upregulated genes belonged to the collagen family, suggesting a role for pinacocytes in the secretion of extracellular matrix (Fig. 6D). As many as fourteen collagens had elevated expression in ISX-treated samples. Adhesion molecules, including cadherins, integrins, and down syndrome cell adhesion molecule, had increased expression levels as well (Fig. 6D). Though many upregulated genes correspond to the transcriptional profile of pinacocytes based on scRNA-seq data47, upregulation of silicatein—a sclerocyte marker—suggests that ISX treatment caused differentiation of other cell types as well, directly or indirectly.
Discussion
A challenge for tracing the ancestry of myocytes is that muscles are very diverse in modern animals. For example, like vertebrates, the ascidian Ciona robusta and the annelid Platynereis dumerilii have striated myocytes that express stMyHC, are regulated by the troponin C/tropomyosin complex, and are developmentally specified by a skeletal muscle-like transcriptional complex. They also have smooth muscles that express nmMyHC and are patterned by a smooth/cardiac muscle-like transcriptional complex5,48. However, in a divergence from the vertebrate paradigm, the flatworm Schistosoma mansoni has smooth muscles that express stMyHC and are patterned by a skeletal-muscle like complex that includes MyoD49,50, but contractions are regulated by MLCK phosphorylation of the RLC. Among non-bilaterian animals, cnidarians have smooth and striated myocytes with epithelial-like organization that express either stMyHC or nmMyHC and are regulated by the MLCK pathway (where known). Their developmental specification is not yet characterized, but they lack the skeletal muscle transcription factor MyoD7. Also, ctenophore myocytes predominantly have smooth ultrastructure (with one exception51), express stMyHC, but lack troponin C and MyoD7,52.
Here, we establish that E. muelleri also contains a contractile module with elements of homology to contractile tissues in other animals, including myocytes. Actomyosin bundles containing stMyHC and transgelin exhibit tissue-wide organization in pinacoderm tissues that line internal body cavities and canals. Contraction depends upon the release of ER-associated Ca2+ stores and MLCK regulation of stMyHC, and the development of the module appears to depend upon MRTF-activity. Our interpretation of this contractile module as muscle-related is corroborated by single-cell sequencing data from the related species, Spongilla lacustris, which indicate that pinacocytes, and another cell type—myopeptidocytes—cluster with myocytes from other animals (myopeptidocytes are solitary cells found between tissues that express contractile genes including nmMyHC)45,47. Single-cell RNA sequencing of the demosponge Amphimedon queenslandica also revealed co-expression of key components of the actin-based contractile apparatus in pinacocytes15.
Although sponge contractile tissues and muscles have similarities that can only be explained by common ancestry, we do not assert that this reflects their one-to-one homology. This is partly due to the similarity of muscle and non-muscle contractile mechanisms, but also because invertebrate muscles are often multifunctional. For example, the epitheliomuscles of Hydra function in contraction, the formation of an epithelial barrier, innate immunity, and regeneration53. In the planarian Schmidtea mediterranea, muscle also acts as a connective tissue that secretes ECM proteins (including 19 collagens) and signaling molecules that provide spatial cues for regeneration from neoblasts54. High expression of numerous collagens has also been found in the epitheliomuscles of Nematostella vectensis (30 in the circular body muscles), which is interpreted as a sign of multifunctionality55. Similarly, sponge pinacoderm tissues form an endothelial-like barrier to the environment, are capable of phagocytosis56, and express genes involved in sensation, metabolism, and defense47. Also, in an intriguing parallel with S. mediterranea and N. vectensis, MRTF-induction of contractile tissue development was associated with the upregulation of as many as 14 collagen genes.
Our findings can inform hypotheses about the organization of contractile tissues in the first animals, as well as the sequence of events that gave rise to early muscles. Specifically, the epithelial-like nature of contractile tissues in both sponges and cnidarians suggests that this represents the ancestral state, and that narrowly specialized myocytes emerged later. This fits well with the view of epithelial tissues as the “building blocks” of animal body plans57, which may have been among the first tissues to evolve. It has been hypothesized that myoepithelia represent a retained feature of an ancient muscle precursor17,58. The first animals were undoubtedly aquatic, so myoepithelia may have functioned to maintain tissue tension in the context of a hydrostatic skeleton or may have functioned in peristalsis-like behaviors like those seen in burrowing of the anemone Nematostella vectensis, or canal constriction in E. muelleri.
Depending on the phylogenetic position of sponges, which is contentious59,60, the long-held view that they lack muscles has been interpreted as evidence that myocytes evolved after sponges diverged from other animals, or that myocytes were lost in sponges61. Our results clarify that the contractile module of muscle tissues predates modern animals. Consistent with this interpretation, the stMyHC, and nmMyHC paralogs diverged in the holozoan stem lineage7,62, and the choanoflagellate Choanoeca flexa forms colonies that resemble a polarized epithelium with coordinated contractile behaviors63.
Finally, the possible role of MRTF in specifying contractile tissues in E. muelleri helps to explain the plasticity and regenerative capacity of sponges. Evidently, without the need for intrinsic spatial cues characteristic of embryogenesis, sponges can develop from archeocyte-enriched aggregates and gemmules, and adult tissues can remodel in response to flow dynamics64,65,66,67. In vertebrates, MRTF is an actin-regulated force-sensor involved in muscle plasticity and regeneration22,68,69, and our data indicate that similar mechanisms are operating in E. muelleri. This supports a model in which environmental-feedback mechanisms drive contractile tissue development, and we speculate that this could be common in sponges. Even during sexual reproduction, there is limited correlation between embryonic patterning and adult tissue identity70. It is conceptually plausible that the evolution of animal developmental mechanisms involved a transition in which ancient environmental feedback mechanisms were later harnessed by genetically encoded, intrinsic patterning mechanisms.
Methods
Sponge collection and cultivation
Gemmules of E. muelleri were collected from an unnamed lake in the Brainard Lake Recreation Area in the Colorado Rocky Mountains (40°04'48.0“N 105°32'34.6“W) and stored in autoclaved lake water (LW) at 4 °C. Before use, gemmules were treated with 1% hydrogen peroxide for 5 min, washed thoroughly with autoclaved lake water, and plated in either six-well plates (CellTreat #229106) or coverslip-bottom dishes (MatTek Corporation #P35G-1.5-10-C) in LW containing 100 µg/mL ampicillin. Plates/dishes were then placed in a dark cabinet at room temperature (RT) until hatching (~3 days). After hatching, water was renewed daily without the addition of ampicillin.
Induction of contraction by mechanical agitation, Sumi ink, and elevated Ca2+
Sponges were induced to contract using mechanical agitation by placing the plate or dish on a vigorous rocking platform for 3 min before transfer to an inverted microscope for imaging. Alternatively, contractions were induced with Sumi Ink (Yasutoma #KY6) by incubating sponges in 1:1000 solution for 10 min, followed by LW washes. To test for Ca2+ dependence of contractions, sponges were placed in 2 mL LW and left on the microscope stage for 1 h to confirm they were not contracting prior to treatment. After 5 min of imaging, treatments were applied by gently removing 1 mL of LW and replacing it with 1 mL of either thapsigargin (Tocris #1138) or ionomycin (Sigma-Aldrich #I0643) at twice the desired final concentrations of 1 or 30 nM, respectively. To induce contractions in the absence of NO signaling or external Ca2+, sponges were pretreated with either 50 µg/mL L-NAME (Cayman Chemical #80210), 0.1 mM EDTA, or deionized water (Milli-Q, Millepore). Time lapse images were collected, generally, at a rate of one image every 20 s and stacks were compiled into videos using ImageJ software. Major excurrent canals were manually annotated from still images. For changes in canal diameter, at least six incurrent and excurrent canal areas were chosen for each sponge and diameter across the same plane was measured every 6th frame over the course of the video. The ratio of diameter at time t to the initial diameter at t = 0 was then taken to represent change in diameter over the course of treatment. Graphs of canal dynamics were generated using Microsoft Excel (version 16)™.
Antibody production and validation
Polyclonal antibodies against stMyHC and MRTF were produced in rabbits (Syd Labs), and polyclonal antibodies against TAGNL2 and TAGNL3 were produced in chickens (Pacific Immunology). The coding region of each antigen (Supplemental Sequences) was cloned into the pET His6 GST TEV LIC cloning vector (Addgene, plasmid #29655) from synthesized gBlocks (Integrated DNA technologies) (stMyHC) or from the E. muelleri cDNA library. Recombinant proteins were expressed in Escherichia coli (Rosetta strain DE3; EMD Millipore) and purified with Pierce Glutathione resin (ThermoFisher Scientific #16101) following the manufacturer’s protocol. Antibodies were affinity purified on a column made by coupling each recombinant protein to AminoLink Resin (ThermoFisher Scientific #20381) and validated by Western blotting and by Peptide Competition Assay (Supplementary Figs. 9–12).
Immunostaining
Sponges were grown in glass-bottom dishes for 3–4 days post hatching and then fixed using one of two methods: (1) to prepare samples for pRLC (Cell Signaling Technology #3671 T), MRTF, TAGNL2, TAGNL3, and vinculin71 staining, sponges were fixed in 3.7% formaldehyde in EtOH for 50 min at room temperature; (2) to prepare sponges for stMyHC staining, sponges were fixed in Carnoy’s solution (60% EtOH, 30% chloroform, 10% glacial acetic acid) for 3 min at RT, followed by a gentle 10 min wash in 100% EtOH at RT. All samples were then washed in PBST and blocked with PBST containing 3% BSA for 1 h at RT. Primary antibodies in blocking solution were applied for 1 h at RT, or overnight at 4 °C (1:100 for anti-pRLC, 1:200 for anti-MRTF, 1:250 for anti-TAGLN2, 1:500 for anti-TAGLN3, 1:1000 for anti-EmVin1 and 1:500 for anti-stMyHC). Secondary antibodies, either anti-rabbit AF488 (Life Technologies) or anti-Chicken IgY AF 647 (Invitrogen #A32933), were applied at a 1:1000 dilution, together with 1:120 phalloidin AF 568 (ThermoFIsher Scientific #A12380), and 1:100 Hoechst dye in blocking solution. Samples were incubated for 45 min at RT in the dark, washed in PBST, and mounted in mounting medium (0.1% Propyl gallate, PBS pH 7.6, 90% glycerol). Samples were imaged on an Olympus Fluoview FV3000 confocal microscope.
In order to understand the developmental dynamics of stMyHC, sponges were fixed in 8 h time-intervals after attachment to the dish. The only change to the fixation protocol described above was that the glacial acetic acid component of Carnoy’s solution was added dropwise during the first minute to help preserve the delicate structure of the newly formed tissues. Since actin filaments were not preserved using this fixation method, we instead performed a timed sequence of latrunculin B treatments to test whether stMyHC staining was disrupted in conjunction with actin-bundle dissolution. Specifically, gemmules were plated in glass-bottom dishes and grown until canals and choanocytes were clearly visible. Sponges were then treated with 20 μM latrunculin B in LW for either 1, 5, 10, or 30 min, then fixed and stained using appropriate methods for either stMyHC or F-actin.
To test for elevated pRLC staining in contracted tissues, 10 z-stacks were taken through comparable regions of the pinacoderm using the same confocal settings for relaxed and contracted samples. Projections of the images were made in Fiji72 and the F-actin and pRLC images were merged. pRLC staining intensity was measured as pixel intensity and expressed as a ratio to actin-bundle length (nine biological and three technical replicates). Measurements were obtained from nine biological and three technical replicates, and a simple regression analysis was performed for pixel intensity and actin-bundle length in RStudio73.
HCR RNA fluorescent in situ hybridization of MRTF and SRF
Multiplexed HCR RNA-FISH was performed with custom split initiator probes ordered from Molecular Instruments®. Sponges were grown in coverslip-bottom dishes (Mattek) for 6 days. Cellbrite-fix 640 (Biotium) was added at 1:1000 dilution for 15 min, and sponges were then washed once in lakewater. Immediately after washing, samples were fixed in ice-cold 4% (v/v) paraformaldehyde in ¼ Holtfreter’s solution overnight at 4 C and then washed with ¼ Holtfreter’s. A dehydration series was performed to transfer the samples to 100% MeOH and then transferred to 100% EtOH before rehydration into 1× PBST. Samples were digested with 5 μg/mL protease K for 90 s and digestion was halted with 2 mg/mL glycine. Samples were then postfixed in 4% (v/v) paraformaldehyde in 1× PBST for 1 h at room temperature and were washed into 2× SSC. Hybridization and amplification were performed using buffers supplied by Molecular Instruments® and following the general protocol with minor modifications. Samples were prehybridized for 30 min in hybridization buffer at 37 C. Samples were then hybridized with 0.5, 2, 5, 10, and 10 pmol probes for each target for ~16 h at 37 C in humidified chambers, on a rocking platform. Samples were then washed into 2× SSC and brought to room temperature. Samples were pre-amplified in provided amplification buffer for 30 min at room temperature and 24 pmol of amplification hairpins for each target were prepared by snap cooling, separately, in amplification buffer. These were combined and added to the sample for 16 h at room temperature in the dark. Samples were then washed into 1× PBST and stained with 1:100 Hoechst dye before mounting.
Testing the role of MLCK and MRTF in physiological contractions
Gemmules were plated in 24-well dishes and treated with the 1 μM ML-7 (MLCK inhibitor74,75), or 50 μg/mL L-NAME for 2 h, or 20 μM or 50 μM CCG-203791 (MRTF inhibitor76,77) for 3 days beginning at tent stage, with treatment refreshed daily. To disrupt flow, sponges were each treated with 1:20 (v/v) Sumi ink for 10 min, carefully washed with LW, and allowed to sit, undisturbed, for 3 h (long enough to allow ink blockages to be cleared in sponges capable of contraction). To test if water flow was restored by contraction, samples were then treated with 1:1000 (v/v) DiI (ThermoFisher Scientific #D282) in LW for 10 min in the dark and washed with LW, which under normal flow conditions strongly stains choanocyte chambers. Samples were manually screened and imaged via epifluorescence microscopy for DiI staining within water canals. Laser-level and exposure time was set based on control sponges and DiI staining was judged based on overlap with Sumi ink-stained choanocyte chambers. Washout experiments and extended treatments were performed to confirm sponges remain viable under treatment conditions.
qRT-PCR
Gemmules were plated in six-well format on 22 mm coverslips with ~10 sponges per well. For MRTF inhibition, sponges were treated with 20 μM CCG-20397176,77 in LW, beginning at ‘tent-stage’ (i.e., before choanocyte chambers formed) and with treatment refreshed daily. For MRTF activation, sponges were treated with 50 μM ISX938,39 beginning at tent-stage and with treatment refreshed daily, or with 10 μM cytochalasin D (ThermoFisher Scientific #PHZ1063) for 30 min prior to harvesting (cytochalasin D disrupts the MRTF/g-actin association through competitive binding of g-actin78). RNA was extracted by washing sponges off of coverslips with Trizol Reagent (ThermoFisher Scientific #15596-018) in a 50 mL tube, followed by ethanol precipitation and rehydration in nuclease-free water. The concentration of purified total RNA was measured by spectrophotometry and an equal mass from each extraction was used to generate cDNA using Takara RNA to cDNA EcoDry mix (Takara #639549). qRT-PCR was performed using a BioRad iQ5 Real-Time PCR detection system (BioRad #170-9780), with an initial denaturing step at 95 °C for 30 s followed by a two-step cycle; 95 °C for 10 s, 60 °C for 55 s, repeated 45 times. A melt-curve was performed after the run to verify the signal was not due to the formation of primer-dimers. Prior to performing the qRT-PCR reaction, a dilution series of purified cDNA library template was used to calculate the amplification efficiency for each of the primer sets in order to verify that it was >1.8 per cycle (2.0 equates to 100% efficiency or doubling each cycle).
Activation of MRTF in archeocytes
Approximately 50 gemmules were grown to 3 days post-hatching, then dissociated in LW containing 1 mM EDTA and passed through a 70 µm cell strainer. Cell suspensions were centrifuged at 1000 × g for 5 min to enrich for archeocytes. The supernatant was removed by pipetting and the cell pellet was resuspended in LW containing 50 µM ISX or DMSO. Using a wide-bore pipette tip, 25 µL of the cell suspension was plated as a hanging drop and maintained for 2–3 days.
Contraction assays were performed by transferring aggregates to the center of a coverslip-bottom dish and adding thapsigargin to a final concentration of 1 nM. Aggregates were photographed every 30 s for 60 min to create time-lapse videos. Time-lapse series were analyzed in Fiji to measure the changes to aggregate size in response to thapsigargin. Immunostaining of aggregates was conducted as described above, except aggregates were stained in 1.5 mL tubes instead of in dishes.
RNAseq and differential gene expression analysis
Approximately 100 gemmules were grown till 3 days post-hatching, then dissociated as described above. Cells were resuspended in treatment solution (LW, 50 μM ISX) or in control solution (LW, DMSO), and were maintained in 10 mL of solution in a 60 mm petri dish on an orbital rocker to prevent attachment. After 24 h, primmorphs were collected and total RNA was extracted using Trizol Reagent. Experiments were performed in triplicate. Generation of a poly-A-selected library and paired-end sequencing on Illumina HiSeq 4000 platform was performed by Novogene (Sacramento California). Adapter sequences were removed from the raw reads and were quality filtered using fastp79. Cleaned reads were mapped to the E. muelleri genome80 using HISAT281, and transcripts were assembled using StringTie82 and htseq-count83. Differential expression analysis was performed using EdgeR84,85 and was visualized from normalized expression using HeatMap286. Gene ontology analysis was performed using BLAST2GO Pro87. The identity of proteins of interest was assessed by performing reciprocal BLAST88, searching Pfam89, and by phylogenetic analysis using phylogeny.fr90. Phylogenetic trees and analysis of proteins of interest in this study are available in the supplement (Supplementary Figs. 13–15).
Reporting summary
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Data availability
Sequences and accession numbers for all genes and proteins analyzed in this study are made available in the supplement. Raw RNAseq reads have been deposited at the NCBI SRA (accession number/BioProject PRJNA718521, BioSamples SAMN18537458, SAMN18537459, SAMN18537460, SAMN18537461, SAMN18537462, SAMN18537463). Raw confocal image files are available at figshare (https://doi.org/10.6084/m9.figshare.20003945). Source data are provided with this paper.
References
Elliott, G. R. D. & Leys, S. P. Coordinated contractions effectively expel water from the aquiferous system of a freshwater sponge. J. Exp. Biol. 210, 3736–3748 (2007).
Elliott, G. R. D. & Leys, S. P. Evidence for glutamate, GABA and NO in coordinating behaviour in the sponge, Ephydatia muelleri (Demospongiae, Spongillidae). J. Exp. Biol. 213, 2310–2321 (2010).
Ellwanger, K., Eich, A. & Nickel, M. GABA and glutamate specifically induce contractions in the sponge Tethya wilhelma. J. Comp. Physiol. A 193, 1–11 (2007).
Ludeman, D. A., Farrar, N., Riesgo, A., Paps, J. & Leys, S. P. Evolutionary origins of sensation in metazoans: functional evidence for a new sensory organ in sponges. BMC Evolut. Biol. 14, 3 (2014).
Brunet, T. et al. The evolutionary origin of bilaterian smooth and striated myocytes. eLife 5, e19607 (2016).
Hooper, S. L. & Thuma, J. B. Invertebrate muscles: muscle specific genes and proteins. Physiol. Rev. 85, 1001–1060 (2005).
Steinmetz, P. R. H. et al. Independent evolution of striated muscles in cnidarians and bilaterians. Nature 487, 231–234 (2012).
Schiaffino, S. & Reggiani, C. Molecular diversity of myofibrillar proteins: gene regulation and functional significance. Physiol. Rev. 76, 371–423 (1996).
Murrell, M., Oakes, P. W., Lenz, M. & Gardel, M. L. Forcing cells into shape: the mechanics of actomyosin contractility. Nat. Rev. Mol. Cell Biol. 16, 486–498 (2015).
Gordon, A. M., Homsher, E. & Regnier, M. Regulation of contraction in striated muscle. Physiol. Rev. 80, 853–924 (2000).
Hong, F. et al. Biochemistry of smooth muscle myosin light chain kinase. Arch. Biochem. Biophys. 510, 135–146 (2011).
Sanders, K. M. Regulation of smooth muscle excitation and contraction. Neurogastroenterol. Motil. 20, 39–53 (2008).
Tanaka, H., Ishimaru, S., Nagatsuka, Y. & Ohashi, K. Smooth muscle-like Ca2+-regulation of actin-myosin interaction in adult jellyfish striated muscle. Sci. Rep. 8, 7776 (2018).
Arendt, D. et al. The origin and evolution of cell types. Nat. Rev. Genet. 17, 744–757 (2016).
Sebé-Pedrós, A. et al. Early metazoan cell type diversity and the evolution of multicellular gene regulation. Nat. Ecol. Evol. 2, 1176–1188 (2018).
Musser, J. M. et al. Profiling cellular diversity in sponges informs animal cell type and nervous system evolution. Science 374, 717–723 (2021).
Nickel, M., Scheer, C., Hammel, J. U., Herzen, J. & Beckmann, F. The contractile sponge epithelium sensu lato–body contraction of the demosponge Tethya wilhelma is mediated by the pinacoderm. J. Exp. Biol. 214, 1692–1698 (2011).
Peña, J. F. et al. Conserved expression of vertebrate microvillar gene homologs in choanocytes of freshwater sponges. Evodevo 7, 13 (2016).
Prosser, C. L. Ionic analyses and effects of ions on contractions of sponge tissues. Z. Vgl. Physiol. 54, 109–120 (1967).
Kamm, K. E. & Stull, J. T. The function of myosin and myosin light chain kinase phosphorylation in smooth muscle. Annu. Rev. Pharmacol. Toxicol. 25, 593–620 (1985).
Hinson, J. S., Medlin, M. D., Lockman, K., Taylor, J. M. & Mack, C. P. Smooth muscle cell-specific transcription is regulated by nuclear localization of the myocardin-related transcription factors. Am. J. Physiol. Heart Circ. Physiol. 292, H1170–H1180 (2007).
Olson, E. N. & Nordheim, A. Linking actin dynamics and gene transcription to drive cellular motile functions. Nat. Rev. Mol. Cell Biol. 11, 353–365 (2010).
Miralles, F., Posern, G., Zaromytidou, A.-I. & Treisman, R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113, 329–342 (2003).
Long, X., Creemers, E. E., Wang, D.-Z., Olson, E. N. & Miano, J. M. Myocardin is a bifunctional switch for smooth versus skeletal muscle differentiation. Proc. Natl Acad. Sci. USA 104, 16570–16575 (2007).
Creemers, E. E., Sutherland, L. B., Oh, J., Barbosa, A. C. & Olson, E. N. Coactivation of MEF2 by the SAP domain proteins myocardin and MASTR. Mol. Cell 23, 83–96 (2006).
Han, Z., Li, X., Wu, J. & Olson, E. N. A myocardin-related transcription factor regulates activity of serum response factor in Drosophila. Proc. Natl Acad. Sci. USA 101, 12567–12572 (2004).
Cenik, B. K. et al. Myocardin-related transcription factors are required for skeletal muscle development. Development 143, 2853–2861 (2016).
Fan, L. et al. Cell contact–dependent regulation of epithelial–myofibroblast transition via the rho-rho kinase-phospho-myosin pathway. Mol. Biol. Cell 18, 1083–1097 (2007).
Gomez, E. W., Chen, Q. K., Gjorevski, N. & Nelson, C. M. Tissue geometry patterns epithelial-mesenchymal transition via intercellular mechanotransduction. J. Cell. Biochem. https://doi.org/10.1002/jcb.22545 (2010).
Gjorevski, N., Boghaert, E. & Nelson, C. M. Regulation of epithelial-mesenchymal transition by transmission of mechanical stress through epithelial tissues. Cancer Microenviron. 5, 29–38 (2012).
Li, S., Wang, D.-Z., Wang, Z., Richardson, J. A. & Olson, E. N. The serum response factor coactivator myocardin is required for vascular smooth muscle development. Proc. Natl Acad. Sci. USA 100, 9366–9370 (2003).
Oh, J., Richardson, J. A. & Olson, E. N. Requirement of myocardin-related transcription factor-B for remodeling of branchial arch arteries and smooth muscle differentiation. Proc. Natl Acad. Sci. USA 102, 15122–15127 (2005).
Li, J. et al. Myocardin-related transcription factor B is required in cardiac neural crest for smooth muscle differentiation and cardiovascular development. Proc. Natl Acad. Sci. USA 102, 8916–8921 (2005).
Li, S., Chang, S., Qi, X., Richardson, J. A. & Olson, E. N. Requirement of a myocardin-related transcription factor for development of mammary myoepithelial cells. Mol. Cell. Biol. 26, 5797–5808 (2006).
Wang, Z., Wang, D.-Z., Pipes, G. C. T. & Olson, E. N. Myocardin is a master regulator of smooth muscle gene expression. Proc. Natl Acad. Sci. USA 100, 7129–7134 (2003).
Hutchings, K. M. et al. Pharmacokinetic optimitzation of CCG-203971: Novel inhibitors of the Rho/MRTF/SRF transcriptional pathway as potential antifibrotic therapeutics for systemic scleroderma. Bioorg. Med. Chem. Lett. 27, 1744–1749 (2017).
Bell, J. L. et al. Optimization of novel nipecotic bis(amide) inhibitors of the Rho/MKL1/SRF transcriptional pathway as potential anti-metastasis agents. Bioorg. Med. Chem. Lett. 23, 3826–3832 (2013).
Russell, J. L., Goetsch, S. C., Aguilar, H. R., Frantz, D. E. & Schneider, J. W. Targeting native adult heart progenitors with cardiogenic small molecules. ACS Chem. Biol. 7, 1067–1076 (2012).
Velasquez, L. S. et al. Activation of MRTF-A-dependent gene expression with a small molecule promotes myofibroblast differentiation and wound healing. Proc. Natl Acad. Sci. USA 110, 16850–16855 (2013).
Alajbegovic, A. et al. MRTFA overexpression promotes conversion of human coronary artery smooth muscle cells into lipid-laden foam cells. Vasc. Pharmacol. 138, 106837 (2021).
Petrik, D. et al. Functional and mechanistic exploration of an adult neurogenesis-promoting small molecule. FASEB J. 26, 3148–3162 (2012).
Schneider, J. W. et al. Small-molecule activation of neuronal cell fate. Nat. Chem. Biol. 4, 408–410 (2008).
Dioum, E. M. et al. A small molecule differentiation inducer increases insulin production by pancreatic β cells. Proc. Natl Acad. Sci. USA 108, 20713–20718 (2011).
Panayiotou, R. et al. Phosphorylation acts positively and negatively to regulate MRTF-A subcellular localisation and activity. Elife 5, e15460 (2016).
Tarashansky, A. J. et al. Mapping single-cell atlases throughout Metazoa unravels cell type evolution. https://doi.org/10.1101/2020.09.28.317784 (2021).
Henderson, J. R. et al. The LIM protein, CRP1, is a smooth muscle marker. Dev. Dyn. 214, 229–238 (1999).
Musser, J. M. et al. Profiling cellular diversity in sponges informs animal cell type and nervous system evolution. https://doi.org/10.1101/758276 (2021).
Johnson, C. J., Razy-Krajka, F. & Stolfi, A. Expression of smooth muscle-like effectors and core cardiomyocyte regulators in the contractile papillae of Ciona. EvoDevo 11, 15 (2020).
Sulbarán, G. et al. An invertebrate smooth muscle with striated muscle myosin filaments. Proc. Natl Acad. Sci. USA 112, E5660–E5668 (2015).
Diaz Soria, C. L. et al. Single-cell atlas of the first intra-mammalian developmental stage of the human parasite Schistosoma mansoni. Nat. Commun. 11, 6411 (2020).
Mackie, G. O., Mills, C. E. & Singla, C. L. Structure and function of the prehensile tentilla of Euplokamis (Ctenophora, Cydippida). Zoomorphology 107, 319–337 (1988).
Dayraud, C. et al. Independent specialisation of myosin II paralogues in muscle vs. non-muscle functions during early animal evolution: a ctenophore perspective. BMC Evol. Biol. 12, 107 (2012).
Buzgariu, W. et al. Multi-functionality and plasticity characterize epithelial cells in Hydra. Tissue Barriers 3, e1068908 (2015).
Cote, L. E., Simental, E. & Reddien, P. W. Muscle functions as a connective tissue and source of extracellular matrix in planarians. Nat. Commun. 10, 1592 (2019).
Cole, A. G. et al. Muscle cell type diversification facilitated by extensive gene duplications. Preprint at bioRxiv https://doi.org/10.1101/2020.07.19.210658 (2020).
Imsiecke, G. Ingestion, digestion, and egestion in Spongilla lacustris (Porifera, Spongillidae) after pulse feeding with Chlamydomonas reinhardtii (Volvocales). Zoomorphology 113, 233–244 (1993).
Tyler, S. Epithelium–the primary building block for metazoan complexity. Integr. Comp. Biol. 43, 55–63 (2003).
Leclère, L. & Röttinger, E. Diversity of cnidarian muscles: function, anatomy, development and regeneration. Front. Cell Dev. Biol. 4, 157 (2016).
Kapli, P. & Telford, M. J. Topology-dependent asymmetry in systematic errors affects phylogenetic placement of Ctenophora and Xenacoelomorpha. Sci Adv 6, eabc5162 (2020).
Redmond, A. K. & McLysaght, A. Evidence for sponges as sister to all other animals from partitioned phylogenomics with mixture models and recoding. Nat. Commun. 12, 1783 (2021).
O’Malley, M. A., Wideman, J. G. & Ruiz-Trillo, I. Losing complexity: the role of simplification in macroevolution. Trends Ecol. Evol. 31, 608–621 (2016).
Sebé-Pedrós, A., Grau-Bové, X., Richards, T. A. & Ruiz-Trillo, I. Evolution and classification of myosins, a paneukaryotic whole-genome approach. Genome Biol. Evol. 6, 290–305 (2014).
Brunet, T. et al. Light-regulated collective contractility in a multicellular choanoflagellate. Science 366, 326–334 (2019).
Lavrov, A. I. & Kosevich, I. A. Sponge cell reaggregation: mechanisms and dynamics of the process. Russian J. Dev. Biol. 45, 205–223 (2014).
Soubigou, A., Ross, E. G., Touhami, Y., Chrismas, N. & Modepalli, V. Regeneration in the sponge partly mimics postlarval development. Development 147, dev193714 (2020).
Ereskovsky, A., Borisenko, I. E., Bolshakov, F. V. & Lavrov, A. I. Whole-body regeneration in sponges: diversity, fine mechanisms, and future prospects. Genes 12, 506 (2021).
Colgren, J. & Nichols, S. A. The significance of sponges for comparative studies of developmental evolution. Wiley Interdiscip. Rev. Dev. Biol. 9, e359 (2020).
Mokalled, M. H., Johnson, A. N., Creemers, E. E. & Olson, E. N. MASTR directs MyoD-dependent satellite cell differentiation during skeletal muscle regeneration. Genes Dev. 26, 190–202 (2012).
Zhang, M. et al. HDAC6 regulates the MRTF-A/SRF axis and vascular smooth muscle cell plasticity. JACC Basic Transl. Sci. 3, 782–795 (2018).
Nakanishi, N., Sogabe, S. & Degnan, B. M. Evolutionary origin of gastrulation: insights from sponge development. BMC Biol. 12, 26 (2014).
Mitchell, J. M. & Nichols, S. A. Diverse cell junctions with unique molecular composition in tissues of a sponge (Porifera). Evodevo 10, 26 (2019).
Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
RStudio Team. RStudio: Integrated Development for R. RStudio (PBC, Boston, MA, 2020).
Lin, H.-B., Cadete, V. J. J., Sawicka, J., Wozniak, M. & Sawicki, G. Effect of the myosin light chain kinase inhibitor ML-7 on the proteome of hearts subjected to ischemia–reperfusion injury. J. Proteom. 75, 5386–5395 (2012).
Gu, X. et al. Cardiac functional improvement in rats with myocardial infarction by up-regulating cardiac myosin light chain kinase with neuregulin. Cardiovasc. Res. 88, 334–343 (2010).
Hayashi, K. ’ichiro., Watanabe, B., Nakagawa, Y., Minami, S. & Morita, T. RPEL proteins are the molecular targets for CCG-1423, an inhibitor of Rho signaling. PLoS ONE 9, e89016 (2014).
Yu-Wai-Man, C. et al. Local delivery of novel MRTF/SRF inhibitors prevents scar tissue formation in a preclinical model of fibrosis. Sci. Rep. 7, 518 (2017).
Medjkane, S., Perez-Sanchez, C., Gaggioli, C., Sahai, E. & Treisman, R. Myocardin-related transcription factors and SRF are required for cytoskeletal dynamics and experimental metastasis. Nat. Cell Biol. 11, 257–268 (2009).
Chen, S., Zhou, Y., Chen, Y. & Gu, J. fastp: an ultra-fast all-in-one FASTQ preprocessor. Bioinformatics 34, i884–i890 (2018).
Kenny, N. J. et al. The genomic basis of animal origins: a chromosomal perspective from the sponge Ephydatia muelleri. https://doi.org/10.1101/2020.02.18.954784 (2020).
Kim, D., Langmead, B. & Salzberg, S. L. HISAT: a fast spliced aligner with low memory requirements. Nat. Methods 12, 357–360 (2015).
Pertea, M. et al. StringTie enables improved reconstruction of a transcriptome from RNA-seq reads. Nat. Biotechnol. 33, 290–295 (2015).
Wen, G. A Simple Process of RNA-Sequence Analyses by Hisat2, Htseq and DESeq2. Proc. 2017 International Conference on Biomedical Engineering and Bioinformatics - ICBEB 2017. https://doi.org/10.1145/3143344.3143354 (2017).
Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).
Liu, R. et al. Why weight? Modelling sample and observational level variability improves power in RNA-seq analyses. Nucleic Acids Res. 43, e97 (2015).
Campagne, F. & Simi, M. MetaR Documentation Booklet. (Fabien Campagne, 2015).
Götz, S. et al. High-throughput functional annotation and data mining with the Blast2GO suite. Nucleic Acids Res. 36, 3420–3435 (2008).
Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J. Basic local alignment search tool. J. Mol. Biol. 215, 403–410 (1990).
Finn, R. D. Pfam: clans, web tools and services. Nucleic Acids Res. 34, D247–D251 (2006).
Dereeper, A. et al. Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res. 36, W465–W469 (2008).
Acknowledgements
We thank Jennyfer Mitchell for help with experimental design and collection of E. muelleri, and Pawel Burkhardt for feedback on the initial manuscript. J.C. and S.A.N. were supported by grants from the National Aeronautics and Space Administration (16-EXO16_2-0041) and the National Science Foundation (IOS:2015608).
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Colgren, J., Nichols, S.A. MRTF specifies a muscle-like contractile module in Porifera. Nat Commun 13, 4134 (2022). https://doi.org/10.1038/s41467-022-31756-9
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DOI: https://doi.org/10.1038/s41467-022-31756-9
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