New insights into unique anatomical structures of the ascidian Halocynthia papillosa obtained by multimodal imaging

New insights into unique anatomical structures of the ascidian Halocynthia papillosa obtained by multimodal imaging | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (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],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article New insights into unique anatomical structures of the ascidian Halocynthia papillosa obtained by multimodal imaging Mareike Huhn, Lukas Hessel, Jonas Albers, Annika Michalek, Elizabeth Duke, and 3 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6701261/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 22 Apr, 2026 Read the published version in Communications Biology → Version 1 posted You are reading this latest preprint version Abstract Current understanding of the anatomical structures of ascidians remains limited, particularly in regionally confined but abundant species such as Halocynthia papillosa from the Mediterranean Sea. This study employed multimodal imaging techniques, including Light, Thunder, and fluorescent confocal microscopy, to investigate selected neural structures and the tunic of H. papillosa at various magnifications. We utilized advanced 3D imaging methods, such as Magnetic Resonance Imaging (MRI) and High-Throughput Tomography (HiTT) at a synchrotron beamline. Additionally, chemical analysis using Fourier-Transformation Infrared-Spectroscopy (FTIR) was conducted. Our 3D and isotropic high-resolution renderings revealed significant structural differences in this species compared to others, notably the absence of a cerebral thickening. HiTT imaging identified three distinct suborders of oral tentacles, each with its own innervation and blood supply. Using Thunder and fluorescent Confocal microscopy, we also documented autofluorescence in ascidian cuticular sheds for the first time in vivo and ex vivo in greater detail. Furthermore, HiTT X-ray imaging of the tunic revealed a spiralized structure emerging from the tunicin layers, which could be verified in this species for the first time using FTIR. The high-resolution, state-of-the-art imaging techniques presented in this study establish a strong foundation for future studies on H. papillosa and other solitary ascidians, and highlight the need to expand research beyond traditional model species. Biological sciences/Zoology/Animal physiology Biological sciences/Biological techniques/High-throughput screening Biological sciences/Biological techniques/Imaging/3-D reconstruction Biological sciences/Biological techniques/Imaging/Fluorescence imaging Biological sciences/Biological techniques/Imaging/Magnetic resonance imaging Neuroanatomy MRI HiTT Synchrotron Autofluorescence H. papillosa Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Introduction The ascidian Halocynthia papillosa (Linnaeus, 1767) is a solitary and sessile benthic filter feeder found in the Mediterranean Sea and along the Portuguese coast of the Northeast Atlantic 1 . Ascidians have become significant model organisms in recent years, primarily due to phylogenetic analyses that identify them as the closest relatives of vertebrates, forming a sister group within chordates 2,3 . Their classification as an evolutionary link between chordates and invertebrates has made them valuable subjects for various biological studies 4 . Ascidians are particularly interesting in ecological and developmental research, as well as in pharmaceutical studies and drug discovery. Numerous bioactive compounds and symbiotic interactions with bacteria in ascidian tissues contribute to their highly developed immune systems 5,6 . Ascidians play a crucial role in benthic ecosystems, particularly in the vertical transport of organic material 7 . Following filtration, ascidians assimilate essential nutrients while expelling waste as excretory pellets through the atrial siphon. These pellets subsequently settle on the seafloor, serving as a food source for scavengers and deposit feeders 8 . Therefore, ascidians, which significantly contribute to reef biomass, facilitate the aggregation of suspended organic matter, enriching nutrient cycles and supporting reef ecosystem stability. Their role in these processes highlights their ecological importance in marine environments. Despite its high abundance in the Mediterranean Sea and the accessibility of its habitat in shallow coastal waters, H. papillosa has not been widely adopted as an experimental model organism. However, research on the anatomy and biology of this species was conducted from the early 1970s through the 2000s 9–13 . These studies primarily focused on histochemical and microscopic examinations of the tunic. In contrast, extensively studied ascidian model organisms such as Ciona intestinalis have been the subject of numerous investigations covering nearly all physiological functions, including the nervous system and embryonic development 14–17 . Comparative analyses of the tunic across different ascidian species and detailed examinations of their neural structures reveal that while the fundamental anatomical features are conserved, they show significant variations in morphology and developmental patterns 18 . Comprehensive microscopic analyses have characterized the structural organization of ascidian tunics, identifying distinct layers. The outermost region, known as the outer cuticle, contains species-specific scales, while the inner layer, called the fundamental layer, consists of the primary structural tissue 9 . Molecular studies have identified the main components of the H. papillosa tunic as sulfated acids, mucopolysaccharides, tunicin, and proteins 10 . The fundamental layer can be divided into distinct sub-layers. While most research has focused on developmental stages, investigations have been limited to individuals 30 days post-morphogenesis, with no comprehensive assessments of adult specimens 11–13 . Even less is known about the neural system in H. papillosa , however, it has been well studied in other ascidian species 18 . Originating from the tentacles, the subcoronal nerve (SCN) extends into the velar sphincter (VS), which encircles the entire lumen of the oral siphon. Nerves extending from the VS into the tentacles form a plexus where motor and sensory components are integrated and indistinguishable. Only a few afferents, with nuclei located in the central ganglion (CG), are distinguishable 19 . The ascending nerves of the coronal organ connect directly to the CG via a nerve bundle, forming a reflex loop with the velar sphincter and squirt muscles 20 . A study by Braun and Stach (2019) provides a comprehensive reference for an essential overview of the CG 18 . Their investigation, which examined 18 ascidian species using microscopic data, enabled the reconstruction of three-dimensional models, revealing variations in the number and organization of brain nerves. However, the study did not include any Halocynthia species. The present study aimed to investigate various sections of the H. papillosa tunic and the neural structures thought to be linked to sound perception and general neural processing. To thoroughly analyze tissue diversity, we employed a combination of microscopic techniques, molecular approaches, and high-throughput tomography X-ray imaging (HiTT) for three-dimensional reconstructions. Initial examinations used low-magnification microscopy, including light microscopy and Thunder microscopy, to provide an overview of neuronal structures and selected body regions, create true-color representations, and visualize autofluorescence in vivo . Subsequently, high-resolution imaging techniques, such as confocal microscopy, were applied to further investigate fine structural details. Autofluorescence imaging was used to generate three-dimensional maximum intensity projections (MIP) of thick tunic sections. In addition to the molecular characterization of tunic components using Fourier-transform infrared spectroscopy (FTIR), high-resolution three-dimensional imaging was conducted using the advanced HiTT technology from EMBL at DESY in Hamburg. The integration of these multimodal imaging methods offers a detailed and multidimensional perspective on neuronal structures and tunic architecture, providing novel insights into their organization and composition. Results Imaging techniques at various scales were utilized to enhance understanding of the anatomical structures of the solitary ascidian Halocynthia papillosa . The imaging results presented here range from the level of the whole animal (3 x 6 cm) to the details of neural structures (200–400 µm). Magnetic resonance imaging (MRI) data was collected from a single individual ascidian. To achieve detailed visualization of internal structures, high-resolution 2D images (100x100 µm in-plane) were acquired in both transverse (Fig. 2 A-B) and sagittal (Fig. 2 C-D) orientations. Additionally, isotropic imaging was conducted at a lower resolution (250x250x250 µm) for three-dimensional renderings of prominent structures (supplementary video 1). The strong contrast in the T2-weighted images allowed for a clear differentiation of internal structures between the zooid and the surrounding tunic. Within the zooid, the pharyngeal basket (PB) was visible in both the sagittal and transverse images (Fig. 2 B-C). In the transverse view, the arrangement of PB structures at a 90° angle became clear. The endostyle extending from posterior to anterior was highly distinguishable against its surroundings (Fig. 2 A-C). Furthermore, the stomach and intestine, with the latter stretching to the atrial siphon, were identified (Fig. 2 D). With additional contrast adjustments, neural structures like the dorsal tubercle (DT) also became apparent, allowing for measurements of its dimensions. The DT measured 1.51 mm in width and 1.29 mm in length as determined from the transverse images (Fig. 2 A). For a general overview of the tunic, we imaged slices from five different individual ascidians using light microscopy. Light microscopy revealed the tunic structure in the cuticular zone (CZ), which contains the cuticular shed (CS) and the underlying fiber bundles (Fig. 3 B), the median zone (MZ) that houses numerous tunicin layers with varying densities of embedded color pigments (Fig. 3 B-C), the peri-epidermal zone (PE) characterized by a high density of nuclei and vacuolar cells 9 , and the epithelium (E), a thin layer that separates the zooid from the tunic itself (rose line in Fig. 3 A). The subcuticular zone (SC) could not be visualized using the magnification applied with light microscopy and is therefore only presented schematically (Fig. 3 A). The integration of various imaging techniques (x-ray, light microscopy, DAPI staining) enabled the creation of a comprehensive scheme summarizing all the information (Fig. 3 A). The appearance of the CS and MZ varied among samples from different individuals and across different sections of the same individual. In tunic sections that had not been exposed to sunlight (left or right side of the animal, depending on its orientation in the reef, and animals collected from caves or beneath overhangs), the MZ appeared pale white with barely distinguishable tunicin layers (Fig. 3 B), while in sections with intense light exposure, the MZ displayed an orange to deep red hue (Fig. 3 C). Overall, the MZ exhibited a color gradient from pale to dark as light exposure increased. When the red color pigments were visible in the tunicin layers, a gradient (decreasing from outside to inside across tunicin layer groups) and a sub-gradient (increasing from outside to inside within each tunicin layer group) were observed (Fig. 3 C). The CS appeared more consistent among slices from different individuals and various slices of the same individual in terms of color and overall shape. The scales were light orange, regardless of the animal's location and coloration (Fig. 3 B-C). Higher-magnification detailed images of the tunic, taken with a Thunder microscope in vivo (top view) and a Confocal microscope (in slice), showed strong autofluorescence exclusively in the CS (Fig. 3 D-L), but not in the underlying tissues (Fig. F&I). The fluorescent pattern of the CS allowed for comparisons between the tunic of relaxed individuals (Fig. 3 D-F) and contracted individuals (Fig. 3 G-I). At low magnification, dark, non-fluorescent spaces between the CS were evident in relaxed individuals (Fig. 3 D). In contracted individuals, no spaces were observed, and the fluorescence covered the entire tunic surface (Fig. 3 G). At higher magnification, a superposition of neighboring CS became clear (Fig. 3 H). The superposition of different CS was not uniform, with some neighboring CS overlapping more than others (white arrow in Fig. 3 H). This variation was also visible in the slices (Fig. 3 F&I). The different imaging angles and magnifications revealed a superposition of the CS during contraction, with some CS being subordinate to others. Various sectional planes of the CS could be depicted (Fig. 3 F&I) and illustrated in a scheme (Fig. 3 I'). Higher magnification images of the CS structure were obtained through confocal microscopy (Fig. 3 J-L). The CS structure appeared conical and tapered, with small tips aligned with the main axial tip on the surface (white arrowheads, Fig. 3 J-K). In the medial direction, rounded, bubble-like structures beneath the CS were visible, oriented in alignment with the overall orientation of the cone (white arrows, Fig. 3 J). The subcuticular structure and fluorescence resembled the outer surface of the cone. A channel was visible, narrowing towards the tip. Higher magnification (63x) revealed fluorescent particles arranged like a string toward the tip (white arrowheads, Fig. 3 L). The presence of fiber bundles had been established in earlier investigations, with the nuclei situated below the spine 10 . DAPI staining highlighted the cell nuclei and their distribution in the tunicin layers and around the CS. Nuclei were absent in the CS and within the CS channel (Fig. 4 A). They were regularly distributed in the tunicin layers and occurred in greater density around the CS. For the structural tunic analysis of the three-dimensional HiTT X-ray images, we selected a sectional plane in the middle of the tunic, approximately equidistant from the epithelial tissue and the spines. In this top view of the tunic, a parallel arrangement of lanes, 50–150 µm wide, was visible (Fig. 4 B-C). These lanes exhibited a crescent-shaped structure and spiraled in some areas. Neighboring lanes were aligned in the same direction, occasionally interspersed with white (i.e., very X-ray-dense) structures that appeared as tiny dots in the sectional image (white arrowhead, Fig. 4 B). The spiralization observed in the top view (Fig. 4 E-F) and the side views of similar positions (Fig. 4 D) suggests that the tunicin forms a cone-shaped depression (Fig. 4 A) from which the fiber bundles (white arrowheads, Fig. 4 B&E) and the CS emerge. The three-dimensional HiTT images supported this assumption. Even within the spiralized tunicin, the crescent-shaped structures remained intact. The dense, white dot-like structures, presumed to be the fiber bundles, were located precisely in the center of the spiralized tunicin (Fig. 4 E-F). In the side view, the less complex layered structure of the tunicin was visible (arrows, Fig. 4 D). This simple layered structure faded at the elevation of the CS (arrowheads, Fig. 4 D). In addition to the anatomical and structural investigations, Fourier transform infrared spectroscopy (FTIR) was conducted on the tunic slices. The analysis included 10 measurement points evenly distributed over a 10 µm thick tunic slice. Measurement points 1–8 were located in the MZ of the tunic, measurement point 9 in the PE, and measurement point 0 in the SC and CZ (Fig. 5 ). Absorbance peaks were found at 1060 cm − 1 , 1035 cm − 1 , 1336/7 cm − 1 , 1430 cm − 1 , and 985 cm − 1 , and were identified as cellulose-like or tunicin complex 21 . A protein compound was detected between 1500 cm − 1 and 1700 cm − 1 (absorbance peak at 1547 cm − 1 corresponding to amide band II absorption, and 1650 cm − 1 corresponding to amide I absorption). The second most prominent peak complex (3200 cm − 1 – 3500 cm − 1 ) exhibited numerous OH-bonds 21 . The central nerve (red box, Fig. 6 A) was identified as a continuous cord that bifurcates dichotomously twice before each siphon (white arrowhead and black outlined arrowhead, Fig. 6 B) and encircles the siphon like a ring. From this surrounding ring, individual strands branched off multiple times, innervating the individual tentacles. A row of adjacent muscle strands was arranged parallel to the nerve (Fig. 6 C-D, darker blue), while an outer layer of muscle strands was found perpendicular to both the nerve and the inner strands (lighter blue). These muscle strands formed bundles (Fig. 6 C-D), which were also visible in the light microscope images (black arrowheads, Fig. 6 B). Using a stitching protocol developed by EMBL Hamburg 2 , the entire data set (total length 8492.25 µm) acquired by HiTT in various orientations could be visualized in a three-dimensional reconstruction (Fig. 6 E-F). The dorsal strand plexus (DSP) could also be visualized and was found partially running parallel to the nerve while also wrapping around it until it lost contact with the nerve and disappeared from the field of view before the nerves divided dichotomously near the atrial siphon (Fig. 6 E-F). Towards the oral siphon, the DSP appeared to be covered by muscles (Fig. 6 E). At this point, the DSP seemed to emerge from the nerve, and together they appeared to form cavities (black arrowheads, Fig. 6 G). The nerve did not exhibit any apparent thickening or structural changes at any site that could be identified as the CG (supplementary video 2). The dorsal tubercle (DT) of H. papillosa was examined using various imaging techniques (Fig. 7 ). It was located above the nerve cord, just anterior to the oral siphon (OS), and became visible under a light microscope following dissection (Fig. 7 B). In the low-magnification image, both siphons (OS and AS, Fig. 7 B) and the entire central nerve cord (black arrowheads, Fig. 7 B), including the initial dichotomous branching of the nerve near the atrial siphon, were identifiable (white arrowheads, Fig. 7 B). Adjacent to the oral siphon, the DT obscured the branching of the central nerve in the light microscopic view. The DT had a diameter ranging from 2 to 5 mm (depending on the animal and measurement direction) and featured a ciliated funnel that appears horseshoe-shaped with elevated horns within a jelly-like shell (7 H–I), as observed and investigated previously in Microcosmus bitunicatus and Halocynthia roretzi 18,22 . The elevation of the horns became apparent in the lateral view (Fig. 7 G-H). Light microscopy displayed the transparent tissue along with a yellow-to-orange hue of the ciliated funnel (black arrowheads, Fig. 7 I). The HiTT acquisitions of the DT were conducted on tissue that underwent a dehydration series with ethanol to enhance contrast during scanning. Consequently, the DT appeared shrunken in the HiTT images (Fig. 7 C-F). Three-dimensional rendering visualized the DT's isolation from the surrounding tissue (Fig. 7 D), the rear view free of surrounding tissue (Fig. 7 E), and a segmentation of the central nerve cord beneath the DT (Fig. 7 E). This segmentation revealed the dichotomous nerve division located directly beneath the DT (Fig. 7 E-F). Another area of focus was the oral tentacles. The oral tentacles of H. papillosa were arranged in a ring within the oral siphon, oriented directly into the inflowing water stream through the siphon (OS, Fig. 8 A). Each tentacle featured smaller sub-tentacles emerging from its main structure (Fig. 8 A’ & B-C). These sub-tentacles were attached to the lower (posterior) end of the main tentacles (Fig. 8 B-C). The tentacles appeared rounder toward the outer side of the oral siphon (Fig. 8 B) and flatter toward the inner side of the zooid (Fig. 8 C). Generally, the tentacles tapered in the medial direction (Fig. 8 B-C). Segmentation of the HiTT images revealed nervous structures (green) and blood vessels (red) within the tentacles (Fig. 8 C-E). Blood vessels extended posteriorly inside the tentacle, branching into each sub-tentacle (Fig. 8 C). When the surrounding tissue was removed in the 3-dimensional projection, the segmented blood vessels (red) and nerves (green) became visible (Fig. 8 D-E). In partial isolation, where only half of the tissue was removed, additional details about the branching of nerves and blood vessels in the sub-tentacles became evident (black and black outlined arrowheads, Fig. 8 D). Many smaller vessels were present inside the tentacles but were not included in the visualization. For clarity, only the larger vessels (first and second order) were segmented and displayed (supplementary video 3). Discussion This study elucidates the anatomical features of the ascidian Halocynthia papillosa through comprehensive whole-animal imaging techniques. It provides an in-depth examination of neural structures, oral tentacles, and tunic sections, achieved through three-dimensional visualization and segmentation of High-Throughput Tomography images, magnetic resonance imaging, and confocal microscopy. For the first time, magnetic resonance imaging successfully visualized an entire ascidian, including its fragile endostyle and pharyngeal basket, within a fully liquid-submerged specimen. Additionally, we have precisely identified the autofluorescent regions within the cuticle sheds of the tunic for the first time in both this species and ascidians in general. Structural analyses utilizing Fourier-transform infrared spectroscopy delineated variations in compositional characteristics across the different layers of the tunic. Unlike in most other ascidian species, the central nerve of H. papillosa did not show any visible thickening that would allow locating the cerebral ganglion (CG). Braun and Stach reconstructed the central nervous system of 14 different ascidian species 18 . All four species investigated that belong to the same order as H. papillosa (Stolidobranchia) showed distinct thickening of the nerve at the CG. For the genus Halocynthia , only one study (on H. roretzi) has published images of the CG, and neither has found any thickening in the central nerve 23 . In the respective study, the authors refer to the entire nerve between the two ends of dichotomous branching as the central ganglion of H. roretzi 23 . In H. roretzi , this section was reported to be 10 mm long 23 . In the present study with H. papillosa , we measured the nerve section between the two dichotomous branches to be 65 mm long, which is ~ 10 times larger than the known sizes of cerebral ganglia in solitary ascidians 18 . We, therefore, suggest that the cerebral ganglion only comprises a subsection of the nerve between the dichotomous branches and is visually not distinct. Further investigations, including staining methods such as Nissl staining, MAP2, BP102, and Pan-Nav1, should clarify this assumption and are planned for future studies. Additional in vivo Ca 2+ signal imaging could clarify the axonal transport in behavioral stress experiments. The CG is possibly located near or at the anterior dichotomous branching underneath the dorsal tubercle (DT). As in other Stolidobranchia, the DT is also often located close to the CG 18 . Furthermore, dichotomous branching has not been observed other than polytomous branching in Stolidobranchia 18 , but it is present multiple times in H. papillosa. Emerging from the first dichotomous branch from the CG, the branching continues until the more peripheral nerves merge into the circular oral or atrial siphon nerve. Additionally, the current data uniquely reveals the three-dimensional internal characteristics of the nerve, as the cerebral nerve forms cavities at the emerging region with the dorsal strand plexus (DST), which may have formed during the development of the cerebral nerve. The anatomy and function of ascidian oral tentacles were first described in Botryllus schlosseri 24 and later detailed in the model organisms Corella inflata, Styela plicata , and Ciona robusta . This study is the first to image and characterize the oral tentacles of H. papillosa . Unlike the oral tentacles of most other species besides Molgula socialis 25 , we found that H. papillosa has branched oral tentacles. The tentacles were divided into three orders of sub-tentacles toward the posterior end, whereas Molgula socialis has four sub-orders 25 . Furthermore, the tentacles were rounded toward the superior direction, which we suggest contributes to lower resistance against the incoming water stream and allows the coronal organ to be present as a fringe on the inferior margin of the tentacles across all suborders. The findings regarding sub-tentacles appear inconsistent within the order of Stolidobranchia, as the family of Stylidae exhibits different shapes, suborders, abundance, and arrangement of the tentacles 24 . The dual abundance observed in different families within the order of Stolidobranchia leads us to hypothesize a secondary loss of sub-tentacles within this order. Further investigations of families within Stolidobranchia should be conducted to confirm this hypothesis. Through the segmentation of inner tentacular structures, we first outline a vascular network within the tentacles. To date, various papers have discussed the occurrence and characteristics of blood or hemolymph systems within tunicates. Consequently, we strongly endorse the concept of a closed or semi-closed vascular system based on the finely distributed capillary system of the tentacles, in alignment with the findings described in Konrad 2016 26 . The cross-sectional elliptical capillaries vary in diameter (height/width) from 79.0 µm/68.1 µm (± 8.40 µm/±9.65 µm) in the first-order tentacle to 33.3 µm/34.0 µm (± 2.83 µm/±4.23 µm) in the second order. Vessels of the third order were not measurable. The blood supply is generated via a single intratentacular vessel located inferiorly, which subdivides into one vessel per sub-tentacle (second order), alternating to the right and left. A similar single subdivision on alternating sides into the sub-tentacles of the third order was also observed. In contrast, an innervating structure was found on the superior side of the tentacle. Similarly, in the vessel distribution, the nerve formed sub-structures to the corresponding sub-tentacles, establishing the innervation of the coronal organ located on the inferior edge of the tentacles. Smaller capillaries emerging between the main branches and the sub-tentacles supply the nerve. The diameters of the first-order nerves are comparable in width, measuring 62.4 µm (± 11.60 µm) and height, 82.53 µm (± 8.45 µm), to the respective blood vessels of a similar order. The tunic of ascidians shows considerable variation across different orders, suborders, and families of ascidians. For example, in Ciona intestinalis , the tunic consists of a gelatinous and thin tissue 27 , whereas in Halocynthia spp. , it has a leatherier consistency 28 . Notably, even within the same genus, such as Halocynthia spp. , distinct differences in the tunic’s structure have been observed 29 . The cuticular sheds found in the tunic of H. papillosa feature spikes that are absent in the closely related species H. roretzi . When the animal contracts, these cuticular sheds form a strong armor that conceals the softer underlying tissues. Furthermore, our findings reveal, for the first time, a significant presence of autofluorescent tissue in solitary tunicates within the cuticular sheds (CS). This autofluorescence is highly concentrated in the cuticular sheds of the tunic, effectively obscuring all non-fluorescent parts of the tissue while the animal is contracted (see Fig. 3 G, H). Studies comparing tunics of different species have been limited, as the tunics of the well-known model organisms within the ascidians do not appear to contain any unique properties. In H. papillosa , the first investigations of the tunic were carried out in the 1980s 10,11 , while the most recent findings on the tunic structure of H. papillosa date back to the 1990s 12,13,30 . Thus, they no longer represent the current state of research and technology. In the present study, anatomical examinations were performed to analyze the structure and alignment of tunic fibers, along with a detailed investigation of the CS. The findings revealed that the autofluorescent CS, with an average diameter of approximately 500 µm in the animal's relaxed state, contains small gaps of up to 50 µm that do not exhibit autofluorescence. In the contracted state, however, these spaces are closed by an overlay of the CS, which completely conceals the deeper soft tissue and results in a seemingly uninterrupted fluorescent image of the outer tunic. In the approximately 300 µm-long conical cuticular spines, cavities were visible beneath the fluorescent cuticle layer. These cavities contained a chain of fluorescent particles arranged like beads on a string, which are transported up and/or down the spine’s center by fiber bundles. Similar observations have been reported in the embryogenesis of the same species, where granules were found along fiber bundles in the developing tunic two days after morphogenesis 13 . Unfortunately, the static image could not reveal the transport direction and requires further investigation. Using HiTT imaging combined with DAPI staining, we could also show that tunicin forms spiralized fibers to create a support structure for the cuticular sheds. This spiraling was visible only within the subcuticular zone (SC), which lies directly beneath the CS and originates from the parallel-arranged tunicin fibers. We suggest that this configuration allows the fiber bundles to extend through the center of the spiral, reaching the tip of the CS. The stained DAPI nuclei of the tunicin fibers also illustrated the dense spiral layering of the fibers in the SC area. Additional FTIR investigations were conducted on a thin slice of tunic. Previous studies on tunic slices of the closely related species H. roretzi and C. intestinalis showed similar results regarding tunicin detection in the central layers of the tunic 29,31 . Minor differences in the overall appearance of the FTIR output between H. roretzi and the current study likely stemmed from slightly different scanner settings. Nevertheless, the current spikes in the range of 1060 cm − 1 to 1035 cm − 1 are consistent with cellulose determinations previously published 21,29 . As anticipated from past light microscopy studies, the outer and inner layers exhibit distinct differences in the tunicin/cellulose spectrogram and show complex superimpositions of other tissue types over the tunicin that are not easily identifiable. The findings of this study provide a comprehensive anatomical examination of the tissues that make up the tunic, oral tentacles, and selected central nervous system structures in ascidians. A detailed analysis of these tissues is essential for understanding the ecological significance of ascidians, which have long been recognized for their role in environmental studies due to their species diversity and widespread presence across various marine ecosystems 17,32,33 . Solitary ascidians, in particular, offer valuable insights into the effects of anthropogenic stressors, such as noise pollution 34,35 , rising temperatures 36 , and variations in water quality 37,38 . Previous research on H. papillosa has established a correlation between elevated temperatures and stress responses, as evidenced by changes in heat shock protein expression and behavioral modifications 36 . Ascidians have also been acknowledged as essential bioindicators in environmental monitoring 38 . Since ascidians are filter feeders, their physiological condition, settlement behavior, and survival rates in coastal environments can serve as indicators of water quality, particularly in relation to pollution levels and nutrient availability. The present study highlights two structural adaptations that may enhance the ability to perceive sound or vibrations. Given the increasing anthropogenic noise pollution in the oceans 39 , especially in coastal regions and harbors where these organisms are predominantly found, there is an urgent need for comprehensive research using indicator species in this area. 40 Consequently, examining the anatomical features and sound perception capabilities of ascidians alongside the traditional indicator species (mammals 41–44 , fish 45 , selected invertebrates 35,40 ) is crucial for a broad view of the oceans' changing environments. Additionally, our findings indicate the necessity of conducting investigations not solely on selected model organisms 17 . While established model organisms like C. intestinalis offer significant advantages—attributed to their known whole genome sequencing data, the availability of genetically modified strains, and the remarkable transparency of their tunic 46 — it is essential to acknowledge that even closely related species within the same phylogenetic family may display considerable anatomical variations, leading to functional and behavioral differences. Anatomical investigations and an enhanced understanding of the cerebral system in various ascidians are also greatly significant. Previous studies reveal findings of rare chemical compounds and secondary metabolites unique to ascidians, many of which demonstrate toxic properties 47,48 or possess anti-tumor potential, making them highly valuable for medical research and drug development 49,50 . Beyond biomedical research, evolutionary interest in ascidians stems from their larvae, which serve as an optimal model for studying early chordate development. Their developmental features closely resemble those of vertebrates while retaining the genomic simplicity of invertebrates 51 , providing insights into the ancestral origins of chordates 11,52 . In addition to the general importance of understanding the differences in anatomical structures among various ascidian species, as previously explained, it is crucial to recognize the role of H. papillosa as a species endemic to the Mediterranean Sea and Eastern Atlantic 53 . Despite its endemism, it is widely distributed and can be regarded as one of the most significant solitary ascidians in the Mediterranean Sea in terms of biomass. It holds economic value due to its popularity among scuba divers and plays a vital ecological role in benthic communities, particularly within coralligenous assemblages 54 . As a non-selective filter feeder, H. papillosa consumes a broad range of microorganisms, including heterotrophic bacteria, phytoplankton, and ciliates, effectively regulating phytoplankton levels and contributing to water clarity 55 . Its filtration activity also establishes it as an important carbon sink, helping to mitigate eutrophication in marine ecosystems. Due to its sensitivity to environmental disturbances, such as sediment resuspension caused by diving activities, H. papillosa serves as a valuable bioindicator for assessing the health of the benthic zones 54 . A decline in its population or size may indicate ecological stress before such stress becomes apparent at higher levels of the food web. Conclusion This study underscores the limitations of concentrating detailed anatomical investigations exclusively on model species within a taxon. To achieve a more comprehensive understanding, well-established imaging and histological techniques should also be utilized for species that, while not widely distributed, play essential roles within their ecosystems. Integrating anatomical and physiological data into extensive genetic databases (e.g., GenBank) has already gained acceptance as standard practice. Biological sciences should further enhance the connection between anatomical and physiological findings with existing nucleotide and protein databases to develop a more holistic understanding of species and their ecological interactions. This strategy will become increasingly important as environmental changes quicken, resulting in shifts in species distributions and range expansions. Materials and Methods Study organism Halocynthia papillosa is a common solitary ascidian found in the Mediterranean Sea. According to the Society for Laboratory Animal Science (GV-SOLAS), no animal testing application was necessary since the animal used is an invertebrate (non-cephalopod). However, the experiments were conducted only by individuals trained and educated in animal experimentation (EU function A). All experiments were performed to the best of our knowledge and in a manner intended to minimize stress and suffering to the animals. All invasive procedures were carried out only after anesthesia. Animal Transportation The organisms were retrieved from the Adriatic Sea near Pula on August 21st, 2022, at a depth ranging from 10 to 25 meters through scuba diving. After consultations with the Croatian Ministry of Environmental and Nature Protection, it was determined that a sampling permit was unnecessary for ascidians at the designated collection site near Pula (N44.83°, E13.84°). The collection process was carried out with precision, using a knife to carefully detach the substrate to which the organisms were adhered, thereby minimizing any potential injury to the specimens. Subsequent transportation to the aquatic laboratory at Ruhr-University Bochum (RUB) was achieved using plastic containers fitted with individual ventilation via air stones and provisions for continuous cooling to maintain a temperature of 16°C. Animal care and husbandry Ascidians were placed in a FLUVAL Flex aquarium with a capacity of 123 liters, equipped with a continuous water flow system driven by an EHEIM compactON 1000 pump (15W), which has a throughput capacity of 1000 liters per hour. This setup was enhanced by an external chilling unit (TECO TK150, maintaining a temperature of 16°C) to ensure optimal thermal conditions. A comprehensive filtration system included an active carbon filter (AquaMedic carbolit 4mm), a biological filter using plastic balls (AquaMedic miniballs), and a ceramic filter (AquaNova NCR-0.5). Feeding protocols consisted of six daily feeds, providing a total of 20 ml (~ 2 x 10 8 cells) of Nannochloropsis salina per ascidian each day. Clean artificial seawater with a salinity of 37 to 39 PSU was supplied through a 30% water exchange every week. The light cycle was set to 12h/12h (light/dark) and was provided by an LED lamp (Aquasky, 6500K, 21W) integrated into the aquarium. Light microscopy A Leica EZ4 was used for light microscopic examinations. The selected lens configuration permits magnifications ranging from 8x to 35x, enabling various illumination techniques, including transmitted light, top light, and side light. This flexibility provided a thorough visualization of the diverse tissue specimens. The light microscope primarily captured consecutive images depicting tissues and dissections. MRI Magnetic resonance imaging (MRI) was conducted ex vivo using a 9.4 T horizontal small animal scanner ( BioSpec® 94/20 USR , Bruker BioSpin GmbH & Co. KG, Germany), equipped with the B-GA12S HP gradient system (Bruker BioSpin GmbH & Co. KG, Germany) at the Leibnitz Institute for Neurobiology in Magdeburg, Germany. A 1 H transmit-receive volume coil with an inner diameter of 40 mm (Bruker BioSpin GmbH & Co. KG, Germany) was utilized for all measurements. The 4% paraformaldehyde (PFA) pre-fixed H. papillosa was incubated in Fomblin® (a hydrogen-free, high-performance precision mechanics pump oil) for two hours to enhance the contrast between the specimen and the surrounding fluid. Based on preliminary experimental testing (not shown here), T2-weighted multi-slice multi-echo (MSME) sequences were found to be the most effective method for visualizing the ascidian. Before the anatomical scans were conducted, a B 0 -map was acquired (TR 15 ms, four avg.; image matrix 64’, FoV 40x40x40 mm 3 ). The following imaging parameters were used: Sequence type: MSME sequence; TR: 13921.8 ms; TE: 16.1/80.5/187.7 ms; Echo averages: 2/10/10; transverse imaging (Fig. 2 A/B) with 25 contiguous slices of 0.5 mm; sagittal imaging (Fig. 2 C/D) with 55 contiguous slices of 0.5 mm; Image size: 3.2x3.8 cm²; Matrix size: 320x380 (resolution: 100x100 µm²); Scan time for transverse images: 0h33m44s; Scan time for sagittal images: 1h28m10s. Isotropic images for 3D visualization (supplemental video 1) were acquired with the same sequence and parameters, but with a matrix size of 320x380 (resolution: 250 x 250 x 250 µm³) and a scan time of 20 h 41 min. Contrast adjustments of the images were performed using ImageJ, and measurements of dimensions were obtained with Osirix®. 3D visualization and explosion videos were produced with VGStudio Max (2024.4). HiTT High-throughput tomography (HiTT) was conducted at the European Molecular Biology Laboratory (EMBL) beamline P14, utilizing the Petra III storage ring at the Deutsches Elektronen-Synchrotron (DESY) in Hamburg 56 . Given the constraints imposed by a field of view (FOV) limited to a width of 1.3 mm, the dissection of the ascidian was undertaken to isolate specific sections of interest. These included the oral tentacles, the complete nerve between the two siphons, a 1 mm punch biopsy of tunic tissue near the oral siphon, and muscle strands between the siphons. The isolated segments were subsequently fixed in 4% PFA overnight, followed by a series of increasing alcohol concentrations according to the protocol established by Zhanmu et al. (2020) 57 to achieve a terminal concentration of 99.8% ethanol, thereby enhancing image contrast in the HiTT data. After preparation, the samples were carefully placed into a 200 µm pipette tip, sealed at the end, fixed to a magnetic goniometer base (MiTeGen Type B5), and covered with the corresponding magnetic lid. The samples were loaded into SPINE pucks and then placed into a sealed sample dewar maintained at room temperature within the beamline hutch. A computer-controlled robotic arm (MARVIN 58 ) was used to mount the samples onto the diffractometer. The optimal scanning position was precisely determined using an integrated light microscope within the diffractometer setup (Arinax MD3). The X-ray energy at beamline P14 is adjustable within the range of 7–30 keV, with our measurements conducted at 12.7 keV. A 10-fold magnifying objective was used for each scan, resulting in an effective voxel size of 650 nm. Samples were imaged at four different propagation distances of 73 mm, 77 mm, 83 mm, and 92 mm. For each distance, a total of 1810 projection images were obtained over an 181° rotation angle, along with 100 additional flat-field frames 56 . The total acquisition time for all four distances was 136 seconds. Following the acquisition phase, reconstruction was completed fully automatically within one minute using the in-house TOMO-CTF software package. Phase retrieval was performed using the contrast-transfer function approach, employing a beta-delta ratio of 0.1. In the case of larger samples, a tiled acquisition of multiple adjacent data sets was performed. Reconstructed volumes were stitched together later using NRStitcher 59 . Data analysis, including segmentation and 3D visualization, was conducted using VG Studio Max (Version 2022.4, 64-bit) following the methodologies outlined in Albers et al. (2024) 56 . Confocal microscopy Confocal microscopy was performed at Ruhr University Bochum (RUB) using a Leica TCS SP5 system. For this analysis, the tunic tissue specimens were fixed in a 4% paraformaldehyde (PFA) solution and subsequently embedded in Tissue-Tec®. Tissue slicing for visualizing cuticular sheds was carried out with a CM 3050 S microtome, producing sections with a thickness of 50 µm. The imaging process involved positioning the tissue slices, which were affixed to super frost sample holders and covered with a cover slip, in an inverted orientation within the microscope. A 40x objective lens was utilized for imaging CS sections, while a 63x oil immersion lens provided enhanced magnification. To generate three-dimensional maximum intensity projections of the spines with significant depth and minimal background noise, virtual slice stacks comprising up to 101 optical planes were recorded. Additional CS sections were imaged in a non-fixed state to further confirm the absence of autofluorescence induced by PFA fixation. All acquired images relied solely on the tissue's inherent autofluorescence without introducing any artificial fluorescence. The GFP laser used for imaging was a DPSS 561. Thunder Thunder imaging was conducted at Ruhr-University Bochum (RUB) using a Leica microscope (model M205 FCA) equipped with a DFC9000 GT camera. This advanced microscopy system offers a magnification range from 0.75x to 16.0x, facilitating extensive imaging applications. Thunder imaging techniques were employed to create whole-animal images and perform initial assessments of auto-fluorescence within the CS at low magnification in the living specimen ( in vivo ). Throughout the study, the straightforward microscope design proved most suitable for two-dimensional imaging with DAPI staining, as light and filters can be easily adjusted. Additionally, conventional mounting fluids were replaced with Roti®-Mount FluorCare DAPI solution (Roth) for tissue sample preparation, allowing for image acquisition with a high signal-to-noise ratio. FTIR spectroscopy Fourier-transform infrared spectroscopy was performed at the Chair of Applied Electrodynamics and Plasma Technology (AEPT) at Ruhr University Bochum, Germany, using a Bruker Hyperion 3000 equipped with an Infinity 1 video camera. The tunic was sliced immediately after anesthesia using a vibratome (Leica VT 1200) without any fixing detergents in salt water (38 PSU). The resulting slice thickness was calibrated to 10 µm. The specimens were promptly transferred to silicon wafers for drying following the slicing procedure. The samples underwent overnight incubation before measurement. In a nitrogen-precooled scanning environment, the scanning process utilized an LN-MCT-D316-025 detector with a total scanning duration of 14 minutes and 17 seconds. It included 32 sample scans and 32 background scans in the microscopic scanning position. Declarations Data availability statement The datasets generated during and/or analyzed during the current study are available from the corresponding author upon request. Acknowledgments HiTT data were collected on EMBL Beamline P14 on the Petra III synchrotron in Hamburg. We thank Diving Pula for support with logistics. Author contributions L.H.: Hypothesis generation, conceptualization, literature search, methodology, data scanning, image analysis, data evaluation and analysis, drawing, manuscript writing. J.A.: Methodology, data scanning, image analysis, data evaluation, and analysis. A.M.: Methodology, data scanning, image analysis, data evaluation, and analysis. E.D.: Methodology, data scanning. I.S.: Methodology S.H.: Conceptualization, funding acquisition. J.G.: Hypothesis generation, conceptualization, methodology, data scanning, image analysis, manuscript writing. 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The same MRI data is shown in Figure 1 (c). Grey shows the tunic, green shows the endostyle, dark red shows the tentacle ring with an incision to free the view on the tentacles, yellow shows the dorsal tubercle, brown shows the compromise of digestive and genital tubes, and turquoise shows the pharyngeal basket. Video2wholenervesegmented.mov Supplementary video 2 Whole central nerve cord segmentation animation. The animations show the whole data set with segmentations in green (nerve cord), yellow (dorsal strand plexus), and blue (muscle fibers). Different angles and cropping of the nerve show cavities emerging at the beginning of the dorsal strand plexus. The same HiTT data is shown in Figure 6. The rendering and segmentation were performed with VG Studio Max (Version 2022.4, 64-bit). Video3OralTentacleSegmentation.avi Supplementary video 3 Tentacle animation. Animation of an oral tentacle with segmentation of nerve tissue (green) and vascularization (red). The rendering and segmentation were performed with VG Studio Max (Version 2022.4, 64-bit). Cite Share Download PDF Status: Published Journal Publication published 22 Apr, 2026 Read the published version in Communications Biology → Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. 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13:02:42","extension":"html","order_by":20,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":136484,"visible":true,"origin":"","legend":"","description":"","filename":"earlyproof.html","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/143492fd9a402aaebfe5fdc4.html"},{"id":94860826,"identity":"9fee7e4e-e547-451f-9906-fa27affb24d4","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":622913,"visible":true,"origin":"","legend":"\u003cp\u003eGeneral anatomy of \u003cem\u003eH. papillosa\u003c/em\u003e. \u003cstrong\u003ea\u003c/strong\u003e In situ picture of \u003cem\u003eHalocynthia papillosa\u003c/em\u003e taken during scuba diving in Pula, Croatia, June 2022. \u003cstrong\u003eb\u003c/strong\u003e Scheme of \u003cem\u003eH. papillosa\u003c/em\u003e, including water inflow (blue arrow, top), water outflow (blue arrow, left), genital flow (small brown arrows), and digestive flow (green arrows). The drawing shows a cross-section of the sagittal plane with the atrial siphon pointing to the left. \u003cstrong\u003ec\u003c/strong\u003e Three-dimensional rendering of the acquired MRI data (raw data shown in Fig. 2). Grey shows the tunic, green shows the endostyle, dark red shows the tentacle ring with an incision to free the view on the tentacles, yellow shows the dorsal tubercle, brown shows the compromise of digestive and genital tubes, and turquoise shows the pharyngeal basket. The figure shows a perspective illustration created with VG Studio Max, which, for the scale, is just an approximation. Abbreviations can be found in the following. A, atrium; AS, atrial siphon; OS, oral siphon; DT, dorsal tubercle; CF, ciliated funnel; AN, anus; GD, genital ducts; EP, esophagus; T/O, testis/ovary; S, stomach; E, epithelium; H, heart; PG, pyloric gland; TU, tunic; ES, endostyle; PS, pharyngeal slit; PB, pharyngeal basket; TE, tentacles; VS, velar sphincter;\u003c/p\u003e","description":"","filename":"floatimage1.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/55a97612819a490343fa65b3.png"},{"id":94986031,"identity":"c7ef87df-fee4-490a-a154-53a9cc27e735","added_by":"auto","created_at":"2025-11-03 06:59:37","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":420223,"visible":true,"origin":"","legend":"\u003cp\u003eOverview of MRI data. T2-weighted MRI images of an ascidian submerged in Fomblin® in transversal (\u003cstrong\u003ea-b\u003c/strong\u003e) and sagittal slices (\u003cstrong\u003ec-d\u003c/strong\u003e) with an in-plane resolution of 100 µm. The number of the displayed slice is given in the lower left of each image, orientation of the ascidian within the images is shown in the lower right. Red dotted lines in \u003cstrong\u003ea\u003c/strong\u003e and \u003cstrong\u003ec\u003c/strong\u003e show the corresponding orthogonal section planes (\u003cstrong\u003ea’\u003c/strong\u003e-\u003cstrong\u003ed’\u003c/strong\u003e). In the top transversal slice (\u003cstrong\u003ea\u003c/strong\u003e), a dotted circle marks the OS; the DT and ES are marked by arrows. Measured DT dimensions are included in green (width) and blue (length). In a more posterior section (\u003cstrong\u003eb\u003c/strong\u003e), the ES, the PBL, the M, and the AS are visible and marked accordingly. The structure of the PBL is outlined by black arrowheads throughout the images \u003cstrong\u003eb-d\u003c/strong\u003e, while white arrowheads show the tunic (\u003cstrong\u003eb, c\u003c/strong\u003e). In the sagittal midline slice of the ascidian (\u003cstrong\u003ec\u003c/strong\u003e), the ES, OS, PBL, and AS are displayed. \u003cstrong\u003ed\u003c/strong\u003e Right lateral side of the ascidian with marked M, INT, and S. OS, oral siphon; DT, dorsal tubercle; ES, endostyle; PBL, pharyngeal basket lumen; M, mantle; AS, atrial siphon; INT, intestine; S, stomach; a, anterior; p, posterior; d, distal; v, ventral; rl, right lateral; ll, left lateral\u003c/p\u003e","description":"","filename":"floatimage2.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/d227ba028b1788c58516114c.png"},{"id":94985060,"identity":"e17f5053-e956-4669-bdc8-6a2dedf6cae0","added_by":"auto","created_at":"2025-11-03 06:57:20","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":397322,"visible":true,"origin":"","legend":"\u003cp\u003eTunic imaged with different microscopes at different magnifications. Tunic slices displayed as scheme (\u003cstrong\u003ea\u003c/strong\u003e)\u003cstrong\u003e,\u003c/strong\u003e light microscopic images (\u003cstrong\u003eb\u003c/strong\u003e-\u003cstrong\u003ec\u003c/strong\u003e), and Thunder microscopic images (\u003cstrong\u003ed\u003c/strong\u003e-\u003cstrong\u003ee\u003c/strong\u003e, \u003cstrong\u003eg\u003c/strong\u003e-\u003cstrong\u003eh\u003c/strong\u003e, \u003cstrong\u003ei\u003c/strong\u003e) and Confocal imaging (\u003cstrong\u003ef\u003c/strong\u003e, \u003cstrong\u003ej\u003c/strong\u003e-\u003cstrong\u003el\u003c/strong\u003e) in different magnifications. The scheme (\u003cstrong\u003ea\u003c/strong\u003e) comprises all features displayed in Fig. 3 (\u003cstrong\u003eb\u003c/strong\u003e-\u003cstrong\u003el\u003c/strong\u003e) and Fig. 4. \u003cstrong\u003eb\u003c/strong\u003e) Thick tunic section from an individual located in a sunlight-protected environment in a relaxed state. \u003cstrong\u003ec\u003c/strong\u003e Individual from a location with regular light exposure in a more contracted state. The numbers (Fig. 3 \u003cstrong\u003ec\u003c/strong\u003e) indicate the four tunicin layer groups with decreasing intensity of color (outside to inside, with increasing numbers). The increasing intensity and color (from outside to inside) are visible within each group. \u003cstrong\u003ed\u003c/strong\u003e) Low magnification of the autofluorescent CS in a not-contracted (relaxed) state with non-fluorescent spaces in between. \u003cstrong\u003ee\u003c/strong\u003e) Same tunic view as (\u003cstrong\u003ed\u003c/strong\u003e) in a higher magnification at location (\u003cstrong\u003ee’\u003c/strong\u003e) with white arrowheads pointing to the non-autofluorescent spaces between the CS. \u003cstrong\u003eg)\u003c/strong\u003e Tunic section similar to (\u003cstrong\u003ed\u003c/strong\u003e-\u003cstrong\u003ee\u003c/strong\u003e) in a contracted state. \u003cstrong\u003eh\u003c/strong\u003e) Same tunic as (\u003cstrong\u003eg\u003c/strong\u003e) in a higher magnification at position (\u003cstrong\u003eh’\u003c/strong\u003e) with a white arrowhead pointing at the overlap of two CS and a white arrow pointing at the submerged Spine of a CS. \u003cstrong\u003ef)\u003c/strong\u003e MIP of a thin tunic section acquired with a confocal microscope, visualizing three different cross-sections of a CS (S1-S3) as indicated in (\u003cstrong\u003ei’\u003c/strong\u003e). \u003cstrong\u003ei\u003c/strong\u003e) Contracted tunic cross-section imaged with a Thunder microscope, visualizing the superimpositions of the CS. Annotations S2 and S3 show the respective cross-sections illustrated in (\u003cstrong\u003ei’\u003c/strong\u003e). \u003cstrong\u003ej\u003c/strong\u003e,\u003cstrong\u003ek\u003c/strong\u003e) High magnification (40x oil lens) MIP of a thin tunic section (50µm) with white arrowheads pointing to small sub-spines on the central spine. White arrows show the bubble-like structure below the spine (\u003cstrong\u003ej\u003c/strong\u003e), while (\u003cstrong\u003ea\u003c/strong\u003e) shows an artifact not to be misinterpreted. \u003cstrong\u003el\u003c/strong\u003e) Higher magnification (63x oil lens) of (\u003cstrong\u003ek\u003c/strong\u003e)\u003cstrong\u003e \u003c/strong\u003ewith white arrowheads pointing to rounded particles lined up like a string. (a), artifact; CS, cuticular shed; CZ, cuticular zone; FB, fiber bundle; MZ, median zone; PE, peri-epidermal zone; SC, subcuticular zone\u003c/p\u003e","description":"","filename":"floatimage3.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/6dff5f05fe6800bf41abff19.png"},{"id":94860833,"identity":"3c25447e-4e22-4a5a-bac4-4f6455a7faf9","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":253585,"visible":true,"origin":"","legend":"\u003cp\u003eTunicin fine structure. \u003cstrong\u003ea\u003c/strong\u003e) DAPI-stained slice of the tunic acquired with a Thunder microscope. The dotted line shows the CS that is not visible in the DAPI staining. White arrowheads show a higher density of stained nuclei at the position of the spiralized tunicin. \u003cstrong\u003eb\u003c/strong\u003e) Tunicin structure in the top view within the MZ acquired with HiTT, with a white arrowhead pointing at a fiber bundle in the cross-section. \u003cstrong\u003ec\u003c/strong\u003e) Same section of tunicin as (\u003cstrong\u003eb\u003c/strong\u003e) with additional annotations to visualize the parallel aligned tunicin fibers (coarse dotted line) and the crescent-shaped fine structure of the tunicin fibers within a bundle (fine dotted line). \u003cstrong\u003ed\u003c/strong\u003e) Part of a cross-section of the tunic of \u003cem\u003eH. papillosa\u003c/em\u003e, imaged with a Thunder microscope in transmitted light to visualize the parallel tunicin fiber layers in the middle of the CS (black arrows), which spiral towards the CS (black arrowheads). Additionally, two different cross-sections of the CS itself are visible. \u003cstrong\u003ef\u003c/strong\u003e,\u003cstrong\u003eg\u003c/strong\u003e) Same section of tunicin directly below a spine in a cross-section from the top view. The white arrowhead shows the fiber bundle in the center of the rising spiral. The dotted white line shows the spiralized tunicin structure with a crescent-shaped fine structure similar to the parallel regions (\u003cstrong\u003eb\u003c/strong\u003e-\u003cstrong\u003ed\u003c/strong\u003e). AS, atrial siphon; CS, cuticular shed; OS, oral siphon;\u003c/p\u003e","description":"","filename":"floatimage4.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/d22abb84f8f8304623fcebe2.png"},{"id":94860835,"identity":"b35f5926-1dd4-4630-857f-525868cf254d","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":700877,"visible":true,"origin":"","legend":"\u003cp\u003eFT-IR measurement of the tunic thin section. Whole spectrum of the FT-IR measurement performed on a thin tunic section. The slice in the upper center of the figure shows the thin tunic section with its exact measurement points shown as colored dots in the magnified section (box on the right). The orientation is from outside (left) to inside (right), and measurement points are numbered from outside to inside. The Diagram shows the Wavenumber (cm-1) on the x-axis and the absorbance units on the y-axis. The graphs are numbered according to the measurement position indicated at the top.\u003c/p\u003e","description":"","filename":"floatimage5.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/78e98a7635ea17cd9be609b0.png"},{"id":94985739,"identity":"d2da0dda-71ca-483b-9536-e88d16193efc","added_by":"auto","created_at":"2025-11-03 06:58:49","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":972543,"visible":true,"origin":"","legend":"\u003cp\u003eNerve, dorsal strand plexus, and muscles between the siphons. \u003cstrong\u003ea\u003c/strong\u003e) Schematic representation of the region investigated in the sea squirt. The red box shows the area from the DT on the right to the dichotomous split of the central nerve on the left. \u003cstrong\u003eb\u003c/strong\u003e) Light microscopic image of the atrial siphon ring muscles (black arrowheads) and the first (white arrowhead) and second (black outlines arrowhead) dichotomous branching of the central nerve after resection of the tunic. \u003cstrong\u003ec\u003c/strong\u003e-\u003cstrong\u003eg\u003c/strong\u003e) Three-dimensional reconstructed renderings made with VG Studio Max in a perspective view. Therefore, no scale is given. \u003cstrong\u003ec\u003c/strong\u003e,\u003cstrong\u003ed\u003c/strong\u003e) Reconstructed and segmented subregion of the muscles in proximity to the nerve strand. Light blue muscle strands show the parallel to the central nerve-aligned muscle strands, while dark blue muscle shows the orthogonal muscle strands. The large bracket marked in (\u003cstrong\u003ee\u003c/strong\u003e) shows the subsection. \u003cstrong\u003ee\u003c/strong\u003e) Stitch of the acquired HiTT data set of the nervous system between the two siphons with a total length of 8492.25 µm, which is equivalent to the red box (\u003cstrong\u003ee’\u003c/strong\u003e) in figure (\u003cstrong\u003ea\u003c/strong\u003e). \u003cstrong\u003ef\u003c/strong\u003e) Different perspective of the same dataset with a visualization of the non-segmented whole FOV in the right part, including a legend that applies to all subfigures (\u003cstrong\u003ec\u003c/strong\u003e-\u003cstrong\u003eg\u003c/strong\u003e). \u003cstrong\u003eg\u003c/strong\u003e) Specific region where the DSP emerges from the nerve and builds cavities in both structures, which are pointed out by black arrowheads. AS, atrial siphon\u003c/p\u003e","description":"","filename":"floatimage6.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/e7d5c81866fef723460c15b7.png"},{"id":94860850,"identity":"63c96dcd-ab76-483d-a6e4-acc45a456b48","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":717084,"visible":true,"origin":"","legend":"\u003cp\u003eDorsal tubercle visualized by different techniques. \u003cstrong\u003ea\u003c/strong\u003e) Schematic drawing of the position of the dorsal tubercle and the central nerve. The dorsal tubercle is additionally marked with a red box. \u003cstrong\u003eb\u003c/strong\u003e) Entire central nerve from the atrial siphon (AS – left) to the oral siphon (OS – right). The course of the nerve is shown with black arrowheads. The nerve splits dichotomously on the left (white arrowheads), resulting in the ring nerve surrounding the AS. The same happens on the right side (OS) but is covered by the dorsal tubercle (DT). \u003cstrong\u003ec\u003c/strong\u003e) Three-dimensional rendering of the DT from the HiTT data. Due to the dehydration process, the structure shrunk slightly. The same structure, without the surrounding tissue, is shown in (\u003cstrong\u003ed\u003c/strong\u003e). The position of (\u003cstrong\u003ec\u003c/strong\u003e-\u003cstrong\u003ef\u003c/strong\u003e) is shown by the red box in (\u003cstrong\u003ea\u003c/strong\u003e). \u003cstrong\u003ee\u003c/strong\u003e) (\u003cstrong\u003ed\u003c/strong\u003e) with a longitudinal rotation angle of 180°. The green color shows the segmented nerve below the DT. \u003cstrong\u003ef\u003c/strong\u003e) Segmented nerve separated with the dichotomous split visible. \u003cstrong\u003eg\u003c/strong\u003e) Side view of the DT acquired with transmitted light from the Thunder microscope. \u003cstrong\u003eh\u003c/strong\u003e,\u003cstrong\u003ei\u003c/strong\u003e) Light microscopic pictures demonstrating the structure's color and texture. Black arrowheads point at the ciliated funnel. Figures (\u003cstrong\u003ec\u003c/strong\u003e-\u003cstrong\u003ef\u003c/strong\u003e) were segmented and rendered with VG Studio Max and were shown in a perspective view without a scalebar. AS, atrial siphon; CG, central ganglion; DT, dorsal tubercle; OS, oral siphon; OT, oral tentacle\u003c/p\u003e","description":"","filename":"floatimage7.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/26d63b764596d652ef08c052.png"},{"id":94860846,"identity":"1b776fc6-9ead-410c-bebc-6ba1486bbd33","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"png","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":293170,"visible":true,"origin":"","legend":"\u003cp\u003eTentacle visualization. Different sections of the oral tentacles. \u003cstrong\u003ea\u003c/strong\u003e) Upper part of a sea squirt with oral (OS) and atrial siphon (AS). \u003cstrong\u003ea’\u003c/strong\u003e) Magnified part of \u003cstrong\u003ea\u003c/strong\u003e with more details in the sub-tentacle structure of the tentacles. \u003cstrong\u003eb\u003c/strong\u003e) Top view of a 3D-rendered tentacle image from HiTT. \u003cstrong\u003ec\u003c/strong\u003e) Same tentacle from below. The red structures in (\u003cstrong\u003ec\u003c/strong\u003e) show the blood vessels segmented and colored within the dataset. \u003cstrong\u003ed\u003c/strong\u003e,\u003cstrong\u003ee\u003c/strong\u003e) 3D image of the blood vessels (red) and the nerval structure (green). While (\u003cstrong\u003ed\u003c/strong\u003e) only shows vessels and nerves, (\u003cstrong\u003ee\u003c/strong\u003e) shows half of the surrounding tissue. Black outlined arrowheads in (\u003cstrong\u003ed\u003c/strong\u003e) show the blood vessels of second order that supply single sub-tentacles. Black arrowheads show nerves of the second order that innervate the sub-tentacles of the first order from the main nerve. Images (\u003cstrong\u003eb\u003c/strong\u003e-\u003cstrong\u003ed\u003c/strong\u003e) were segmented and rendered with VG Studio Max. While (\u003cstrong\u003eb\u003c/strong\u003e) and (\u003cstrong\u003ec\u003c/strong\u003e) represent a parallel view, including a scale bar, (\u003cstrong\u003ed\u003c/strong\u003e) and (\u003cstrong\u003ee\u003c/strong\u003e) represent a perspective view, so no scale bar is shown. AS, atrial siphon; OS, oral siphon\u003c/p\u003e","description":"","filename":"floatimage8.png","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/caf16d02cd56d5e08730c61d.png"},{"id":107604276,"identity":"c5af7a96-78dd-4d92-adb9-f6a639bc3673","added_by":"auto","created_at":"2026-04-23 07:22:56","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":5053489,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/2c06dece-ecd8-4fbb-9e6f-b9c33ecb703f.pdf"},{"id":94860842,"identity":"5a536a69-81cd-4e77-ab9d-cabdd9570c82","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"mp4","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":19537230,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSupplementary video 1\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eExplosion animation of the whole \u003cem\u003eH. papillosa\u003c/em\u003e. The video shows an explosion rendering of the MRI data segmented and animated with VG Studio Max (Version 2022.4, 64-bit). The same MRI data is shown in Figure 1 (\u003cstrong\u003ec\u003c/strong\u003e). Grey shows the tunic, green shows the endostyle, dark red shows the tentacle ring with an incision to free the view on the tentacles, yellow shows the dorsal tubercle, brown shows the compromise of digestive and genital tubes, and turquoise shows the pharyngeal basket.\u003c/p\u003e","description":"","filename":"Video1Explosionrenderingwholeanimal.mp4","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/2a7e46ffc6684efab7b61e72.mp4"},{"id":94860857,"identity":"b79e8b0b-ee72-42f6-8c19-1bdfe7363671","added_by":"auto","created_at":"2025-10-31 13:02:42","extension":"mov","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":33687729,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSupplementary video 2\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWhole central nerve cord segmentation animation. The animations show the whole data set with segmentations in green (nerve cord), yellow (dorsal strand plexus), and blue (muscle fibers). Different angles and cropping of the nerve show cavities emerging at the beginning of the dorsal strand plexus. The same HiTT data is shown in Figure 6\u003cstrong\u003e.\u003c/strong\u003e The rendering and segmentation were performed with VG Studio Max (Version 2022.4, 64-bit).\u003c/p\u003e","description":"","filename":"Video2wholenervesegmented.mov","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/fd19c4912c4e6d541d6093f9.mov"},{"id":94860858,"identity":"938394a1-f77e-4924-ac4d-0af4a7cff142","added_by":"auto","created_at":"2025-10-31 13:02:43","extension":"avi","order_by":3,"title":"","display":"","copyAsset":false,"role":"supplement","size":104339224,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSupplementary video 3\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTentacle animation. Animation of an oral tentacle with segmentation of nerve tissue (green) and vascularization (red). The rendering and segmentation were performed with VG Studio Max (Version 2022.4, 64-bit).\u003c/p\u003e","description":"","filename":"Video3OralTentacleSegmentation.avi","url":"https://assets-eu.researchsquare.com/files/rs-6701261/v1/ecda88b6f34a09f1f79ae85d.avi"}],"financialInterests":"There is \u003cb\u003eNO\u003c/b\u003e Competing Interest.","formattedTitle":"New insights into unique anatomical structures of the ascidian Halocynthia papillosa obtained by multimodal imaging","fulltext":[{"header":"Introduction","content":"\u003cp\u003eThe ascidian \u003cem\u003eHalocynthia papillosa\u003c/em\u003e (Linnaeus, 1767) is a solitary and sessile benthic filter feeder found in the Mediterranean Sea and along the Portuguese coast of the Northeast Atlantic\u003csup\u003e1\u003c/sup\u003e. Ascidians have become significant model organisms in recent years, primarily due to phylogenetic analyses that identify them as the closest relatives of vertebrates, forming a sister group within chordates\u003csup\u003e2,3\u003c/sup\u003e. Their classification as an evolutionary link between chordates and invertebrates has made them valuable subjects for various biological studies\u003csup\u003e4\u003c/sup\u003e. Ascidians are particularly interesting in ecological and developmental research, as well as in pharmaceutical studies and drug discovery. Numerous bioactive compounds and symbiotic interactions with bacteria in ascidian tissues contribute to their highly developed immune systems\u003csup\u003e5,6\u003c/sup\u003e. Ascidians play a crucial role in benthic ecosystems, particularly in the vertical transport of organic material\u003csup\u003e7\u003c/sup\u003e. Following filtration, ascidians assimilate essential nutrients while expelling waste as excretory pellets through the atrial siphon. These pellets subsequently settle on the seafloor, serving as a food source for scavengers and deposit feeders\u003csup\u003e8\u003c/sup\u003e. Therefore, ascidians, which significantly contribute to reef biomass, facilitate the aggregation of suspended organic matter, enriching nutrient cycles and supporting reef ecosystem stability. Their role in these processes highlights their ecological importance in marine environments.\u003c/p\u003e \u003cp\u003eDespite its high abundance in the Mediterranean Sea and the accessibility of its habitat in shallow coastal waters, \u003cem\u003eH. papillosa\u003c/em\u003e has not been widely adopted as an experimental model organism. However, research on the anatomy and biology of this species was conducted from the early 1970s through the 2000s\u003csup\u003e9\u0026ndash;13\u003c/sup\u003e. These studies primarily focused on histochemical and microscopic examinations of the tunic. In contrast, extensively studied ascidian model organisms such as \u003cem\u003eCiona intestinalis\u003c/em\u003e have been the subject of numerous investigations covering nearly all physiological functions, including the nervous system and embryonic development\u003csup\u003e14\u0026ndash;17\u003c/sup\u003e. Comparative analyses of the tunic across different ascidian species and detailed examinations of their neural structures reveal that while the fundamental anatomical features are conserved, they show significant variations in morphology and developmental patterns\u003csup\u003e18\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eComprehensive microscopic analyses have characterized the structural organization of ascidian tunics, identifying distinct layers. The outermost region, known as the outer cuticle, contains species-specific scales, while the inner layer, called the fundamental layer, consists of the primary structural tissue\u003csup\u003e9\u003c/sup\u003e. Molecular studies have identified the main components of the \u003cem\u003eH. papillosa\u003c/em\u003e tunic as sulfated acids, mucopolysaccharides, tunicin, and proteins\u003csup\u003e10\u003c/sup\u003e. The fundamental layer can be divided into distinct sub-layers. While most research has focused on developmental stages, investigations have been limited to individuals 30 days post-morphogenesis, with no comprehensive assessments of adult specimens\u003csup\u003e11\u0026ndash;13\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eEven less is known about the neural system in \u003cem\u003eH. papillosa\u003c/em\u003e, however, it has been well studied in other ascidian species\u003csup\u003e18\u003c/sup\u003e. Originating from the tentacles, the subcoronal nerve (SCN) extends into the velar sphincter (VS), which encircles the entire lumen of the oral siphon. Nerves extending from the VS into the tentacles form a plexus where motor and sensory components are integrated and indistinguishable. Only a few afferents, with nuclei located in the central ganglion (CG), are distinguishable\u003csup\u003e19\u003c/sup\u003e. The ascending nerves of the coronal organ connect directly to the CG via a nerve bundle, forming a reflex loop with the velar sphincter and squirt muscles\u003csup\u003e20\u003c/sup\u003e. A study by Braun and Stach (2019) provides a comprehensive reference for an essential overview of the CG\u003csup\u003e18\u003c/sup\u003e. Their investigation, which examined 18 ascidian species using microscopic data, enabled the reconstruction of three-dimensional models, revealing variations in the number and organization of brain nerves. However, the study did not include any \u003cem\u003eHalocynthia\u003c/em\u003e species.\u003c/p\u003e \u003cp\u003eThe present study aimed to investigate various sections of the \u003cem\u003eH. papillosa\u003c/em\u003e tunic and the neural structures thought to be linked to sound perception and general neural processing. To thoroughly analyze tissue diversity, we employed a combination of microscopic techniques, molecular approaches, and high-throughput tomography X-ray imaging (HiTT) for three-dimensional reconstructions. Initial examinations used low-magnification microscopy, including light microscopy and Thunder microscopy, to provide an overview of neuronal structures and selected body regions, create true-color representations, and visualize autofluorescence \u003cem\u003ein vivo\u003c/em\u003e. Subsequently, high-resolution imaging techniques, such as confocal microscopy, were applied to further investigate fine structural details. Autofluorescence imaging was used to generate three-dimensional maximum intensity projections (MIP) of thick tunic sections. In addition to the molecular characterization of tunic components using Fourier-transform infrared spectroscopy (FTIR), high-resolution three-dimensional imaging was conducted using the advanced HiTT technology from EMBL at DESY in Hamburg. The integration of these multimodal imaging methods offers a detailed and multidimensional perspective on neuronal structures and tunic architecture, providing novel insights into their organization and composition.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003eImaging techniques at various scales were utilized to enhance understanding of the anatomical structures of the solitary ascidian \u003cem\u003eHalocynthia papillosa\u003c/em\u003e. The imaging results presented here range from the level of the whole animal (3 x 6 cm) to the details of neural structures (200\u0026ndash;400 \u0026micro;m).\u003c/p\u003e \u003cp\u003eMagnetic resonance imaging (MRI) data was collected from a single individual ascidian. To achieve detailed visualization of internal structures, high-resolution 2D images (100x100 \u0026micro;m in-plane) were acquired in both transverse (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA-B) and sagittal (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC-D) orientations. Additionally, isotropic imaging was conducted at a lower resolution (250x250x250 \u0026micro;m) for three-dimensional renderings of prominent structures (supplementary video 1). The strong contrast in the T2-weighted images allowed for a clear differentiation of internal structures between the zooid and the surrounding tunic. Within the zooid, the pharyngeal basket (PB) was visible in both the sagittal and transverse images (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB-C). In the transverse view, the arrangement of PB structures at a 90\u0026deg; angle became clear. The endostyle extending from posterior to anterior was highly distinguishable against its surroundings (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA-C). Furthermore, the stomach and intestine, with the latter stretching to the atrial siphon, were identified (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eD). With additional contrast adjustments, neural structures like the dorsal tubercle (DT) also became apparent, allowing for measurements of its dimensions. The DT measured 1.51 mm in width and 1.29 mm in length as determined from the transverse images (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFor a general overview of the tunic, we imaged slices from five different individual ascidians using light microscopy. Light microscopy revealed the tunic structure in the cuticular zone (CZ), which contains the cuticular shed (CS) and the underlying fiber bundles (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB), the median zone (MZ) that houses numerous tunicin layers with varying densities of embedded color pigments (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB-C), the peri-epidermal zone (PE) characterized by a high density of nuclei and vacuolar cells\u003csup\u003e9\u003c/sup\u003e, and the epithelium (E), a thin layer that separates the zooid from the tunic itself (rose line in Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). The subcuticular zone (SC) could not be visualized using the magnification applied with light microscopy and is therefore only presented schematically (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). The integration of various imaging techniques (x-ray, light microscopy, DAPI staining) enabled the creation of a comprehensive scheme summarizing all the information (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). The appearance of the CS and MZ varied among samples from different individuals and across different sections of the same individual. In tunic sections that had not been exposed to sunlight (left or right side of the animal, depending on its orientation in the reef, and animals collected from caves or beneath overhangs), the MZ appeared pale white with barely distinguishable tunicin layers (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB), while in sections with intense light exposure, the MZ displayed an orange to deep red hue (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC). Overall, the MZ exhibited a color gradient from pale to dark as light exposure increased. When the red color pigments were visible in the tunicin layers, a gradient (decreasing from outside to inside across tunicin layer groups) and a sub-gradient (increasing from outside to inside within each tunicin layer group) were observed (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe CS appeared more consistent among slices from different individuals and various slices of the same individual in terms of color and overall shape. The scales were light orange, regardless of the animal's location and coloration (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB-C). Higher-magnification detailed images of the tunic, taken with a Thunder microscope \u003cem\u003ein vivo\u003c/em\u003e (top view) and a Confocal microscope (in slice), showed strong autofluorescence exclusively in the CS (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD-L), but not in the underlying tissues (Fig. F\u0026amp;I). The fluorescent pattern of the CS allowed for comparisons between the tunic of relaxed individuals (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD-F) and contracted individuals (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG-I). At low magnification, dark, non-fluorescent spaces between the CS were evident in relaxed individuals (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD). In contracted individuals, no spaces were observed, and the fluorescence covered the entire tunic surface (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG). At higher magnification, a superposition of neighboring CS became clear (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eH).\u003c/p\u003e \u003cp\u003eThe superposition of different CS was not uniform, with some neighboring CS overlapping more than others (white arrow in Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eH). This variation was also visible in the slices (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eF\u0026amp;I). The different imaging angles and magnifications revealed a superposition of the CS during contraction, with some CS being subordinate to others. Various sectional planes of the CS could be depicted (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eF\u0026amp;I) and illustrated in a scheme (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eI'). Higher magnification images of the CS structure were obtained through confocal microscopy (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eJ-L). The CS structure appeared conical and tapered, with small tips aligned with the main axial tip on the surface (white arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eJ-K). In the medial direction, rounded, bubble-like structures beneath the CS were visible, oriented in alignment with the overall orientation of the cone (white arrows, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eJ). The subcuticular structure and fluorescence resembled the outer surface of the cone. A channel was visible, narrowing towards the tip. Higher magnification (63x) revealed fluorescent particles arranged like a string toward the tip (white arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eL). The presence of fiber bundles had been established in earlier investigations, with the nuclei situated below the spine\u003csup\u003e10\u003c/sup\u003e. DAPI staining highlighted the cell nuclei and their distribution in the tunicin layers and around the CS. Nuclei were absent in the CS and within the CS channel (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA). They were regularly distributed in the tunicin layers and occurred in greater density around the CS.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFor the structural tunic analysis of the three-dimensional HiTT X-ray images, we selected a sectional plane in the middle of the tunic, approximately equidistant from the epithelial tissue and the spines. In this top view of the tunic, a parallel arrangement of lanes, 50\u0026ndash;150 \u0026micro;m wide, was visible (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB-C). These lanes exhibited a crescent-shaped structure and spiraled in some areas. Neighboring lanes were aligned in the same direction, occasionally interspersed with white (i.e., very X-ray-dense) structures that appeared as tiny dots in the sectional image (white arrowhead, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB). The spiralization observed in the top view (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE-F) and the side views of similar positions (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD) suggests that the tunicin forms a cone-shaped depression (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA) from which the fiber bundles (white arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB\u0026amp;E) and the CS emerge. The three-dimensional HiTT images supported this assumption. Even within the spiralized tunicin, the crescent-shaped structures remained intact. The dense, white dot-like structures, presumed to be the fiber bundles, were located precisely in the center of the spiralized tunicin (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE-F). In the side view, the less complex layered structure of the tunicin was visible (arrows, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD). This simple layered structure faded at the elevation of the CS (arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD).\u003c/p\u003e \u003cp\u003eIn addition to the anatomical and structural investigations, Fourier transform infrared spectroscopy (FTIR) was conducted on the tunic slices. The analysis included 10 measurement points evenly distributed over a 10 \u0026micro;m thick tunic slice. Measurement points 1\u0026ndash;8 were located in the MZ of the tunic, measurement point 9 in the PE, and measurement point 0 in the SC and CZ (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003e). Absorbance peaks were found at 1060 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, 1035 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, 1336/7 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, 1430 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, and 985 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, and were identified as cellulose-like or tunicin complex\u003csup\u003e21\u003c/sup\u003e. A protein compound was detected between 1500 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e and 1700 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e (absorbance peak at 1547 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e corresponding to amide band II absorption, and 1650 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e corresponding to amide I absorption). The second most prominent peak complex (3200 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e \u0026ndash; 3500 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e) exhibited numerous OH-bonds\u003csup\u003e21\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe central nerve (red box, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA) was identified as a continuous cord that bifurcates dichotomously twice before each siphon (white arrowhead and black outlined arrowhead, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eB) and encircles the siphon like a ring. From this surrounding ring, individual strands branched off multiple times, innervating the individual tentacles. A row of adjacent muscle strands was arranged parallel to the nerve (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC-D, darker blue), while an outer layer of muscle strands was found perpendicular to both the nerve and the inner strands (lighter blue). These muscle strands formed bundles (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC-D), which were also visible in the light microscope images (black arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eB). Using a stitching protocol developed by EMBL Hamburg\u003csup\u003e2\u003c/sup\u003e, the entire data set (total length 8492.25 \u0026micro;m) acquired by HiTT in various orientations could be visualized in a three-dimensional reconstruction (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eE-F). The dorsal strand plexus (DSP) could also be visualized and was found partially running parallel to the nerve while also wrapping around it until it lost contact with the nerve and disappeared from the field of view before the nerves divided dichotomously near the atrial siphon (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eE-F). Towards the oral siphon, the DSP appeared to be covered by muscles (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eE). At this point, the DSP seemed to emerge from the nerve, and together they appeared to form cavities (black arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eG). The nerve did not exhibit any apparent thickening or structural changes at any site that could be identified as the CG (supplementary video 2).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe dorsal tubercle (DT) of \u003cem\u003eH. papillosa\u003c/em\u003e was examined using various imaging techniques (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). It was located above the nerve cord, just anterior to the oral siphon (OS), and became visible under a light microscope following dissection (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB). In the low-magnification image, both siphons (OS and AS, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB) and the entire central nerve cord (black arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB), including the initial dichotomous branching of the nerve near the atrial siphon, were identifiable (white arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB). Adjacent to the oral siphon, the DT obscured the branching of the central nerve in the light microscopic view. The DT had a diameter ranging from 2 to 5 mm (depending on the animal and measurement direction) and featured a ciliated funnel that appears horseshoe-shaped with elevated horns within a jelly-like shell (7 H\u0026ndash;I), as observed and investigated previously in \u003cem\u003eMicrocosmus bitunicatus\u003c/em\u003e and \u003cem\u003eHalocynthia roretzi\u003c/em\u003e\u003csup\u003e18,22\u003c/sup\u003e. The elevation of the horns became apparent in the lateral view (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eG-H). Light microscopy displayed the transparent tissue along with a yellow-to-orange hue of the ciliated funnel (black arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eI). The HiTT acquisitions of the DT were conducted on tissue that underwent a dehydration series with ethanol to enhance contrast during scanning. Consequently, the DT appeared shrunken in the HiTT images (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eC-F). Three-dimensional rendering visualized the DT's isolation from the surrounding tissue (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eD), the rear view free of surrounding tissue (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eE), and a segmentation of the central nerve cord beneath the DT (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eE). This segmentation revealed the dichotomous nerve division located directly beneath the DT (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eE-F).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eAnother area of focus was the oral tentacles. The oral tentacles of \u003cem\u003eH. papillosa\u003c/em\u003e were arranged in a ring within the oral siphon, oriented directly into the inflowing water stream through the siphon (OS, Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eA). Each tentacle featured smaller sub-tentacles emerging from its main structure (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eA\u0026rsquo; \u0026amp; B-C). These sub-tentacles were attached to the lower (posterior) end of the main tentacles (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eB-C). The tentacles appeared rounder toward the outer side of the oral siphon (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eB) and flatter toward the inner side of the zooid (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eC). Generally, the tentacles tapered in the medial direction (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eB-C). Segmentation of the HiTT images revealed nervous structures (green) and blood vessels (red) within the tentacles (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eC-E). Blood vessels extended posteriorly inside the tentacle, branching into each sub-tentacle (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eC). When the surrounding tissue was removed in the 3-dimensional projection, the segmented blood vessels (red) and nerves (green) became visible (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eD-E). In partial isolation, where only half of the tissue was removed, additional details about the branching of nerves and blood vessels in the sub-tentacles became evident (black and black outlined arrowheads, Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eD). Many smaller vessels were present inside the tentacles but were not included in the visualization. For clarity, only the larger vessels (first and second order) were segmented and displayed (supplementary video 3).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eThis study elucidates the anatomical features of the ascidian \u003cem\u003eHalocynthia papillosa\u003c/em\u003e through comprehensive whole-animal imaging techniques. It provides an in-depth examination of neural structures, oral tentacles, and tunic sections, achieved through three-dimensional visualization and segmentation of High-Throughput Tomography images, magnetic resonance imaging, and confocal microscopy. For the first time, magnetic resonance imaging successfully visualized an entire ascidian, including its fragile endostyle and pharyngeal basket, within a fully liquid-submerged specimen. Additionally, we have precisely identified the autofluorescent regions within the cuticle sheds of the tunic for the first time in both this species and ascidians in general. Structural analyses utilizing Fourier-transform infrared spectroscopy delineated variations in compositional characteristics across the different layers of the tunic.\u003c/p\u003e \u003cp\u003eUnlike in most other ascidian species, the central nerve of \u003cem\u003eH. papillosa\u003c/em\u003e did not show any visible thickening that would allow locating the cerebral ganglion (CG). Braun and Stach reconstructed the central nervous system of 14 different ascidian species\u003csup\u003e18\u003c/sup\u003e. All four species investigated that belong to the same order as \u003cem\u003eH. papillosa\u003c/em\u003e (Stolidobranchia) showed distinct thickening of the nerve at the CG. For the genus \u003cem\u003eHalocynthia\u003c/em\u003e, only one study (on \u003cem\u003eH. roretzi)\u003c/em\u003e has published images of the CG, and neither has found any thickening in the central nerve\u003csup\u003e23\u003c/sup\u003e. In the respective study, the authors refer to the entire nerve between the two ends of dichotomous branching as the central ganglion of \u003cem\u003eH. roretzi\u003c/em\u003e\u003csup\u003e23\u003c/sup\u003e. In \u003cem\u003eH. roretzi\u003c/em\u003e, this section was reported to be 10 mm long\u003csup\u003e23\u003c/sup\u003e. In the present study with \u003cem\u003eH. papillosa\u003c/em\u003e, we measured the nerve section between the two dichotomous branches to be 65 mm long, which is ~\u0026thinsp;10 times larger than the known sizes of cerebral ganglia in solitary ascidians\u003csup\u003e18\u003c/sup\u003e. We, therefore, suggest that the cerebral ganglion only comprises a subsection of the nerve between the dichotomous branches and is visually not distinct. Further investigations, including staining methods such as Nissl staining, MAP2, BP102, and Pan-Nav1, should clarify this assumption and are planned for future studies. Additional \u003cem\u003ein vivo\u003c/em\u003e Ca\u003csup\u003e2+\u003c/sup\u003e signal imaging could clarify the axonal transport in behavioral stress experiments. The CG is possibly located near or at the anterior dichotomous branching underneath the dorsal tubercle (DT). As in other Stolidobranchia, the DT is also often located close to the CG\u003csup\u003e18\u003c/sup\u003e. Furthermore, dichotomous branching has not been observed other than polytomous branching in Stolidobranchia\u003csup\u003e18\u003c/sup\u003e, but it is present multiple times in \u003cem\u003eH. papillosa.\u003c/em\u003e Emerging from the first dichotomous branch from the CG, the branching continues until the more peripheral nerves merge into the circular oral or atrial siphon nerve. Additionally, the current data uniquely reveals the three-dimensional internal characteristics of the nerve, as the cerebral nerve forms cavities at the emerging region with the dorsal strand plexus (DST), which may have formed during the development of the cerebral nerve.\u003c/p\u003e \u003cp\u003eThe anatomy and function of ascidian oral tentacles were first described in \u003cem\u003eBotryllus schlosseri\u003c/em\u003e\u003csup\u003e\u003cem\u003e24\u003c/em\u003e\u003c/sup\u003e and later detailed in the model organisms \u003cem\u003eCorella inflata, Styela plicata\u003c/em\u003e, and \u003cem\u003eCiona robusta\u003c/em\u003e. This study is the first to image and characterize the oral tentacles of \u003cem\u003eH. papillosa\u003c/em\u003e. Unlike the oral tentacles of most other species besides \u003cem\u003eMolgula socialis\u003c/em\u003e\u003csup\u003e25\u003c/sup\u003e, we found that \u003cem\u003eH. papillosa\u003c/em\u003e has branched oral tentacles. The tentacles were divided into three orders of sub-tentacles toward the posterior end, whereas \u003cem\u003eMolgula socialis\u003c/em\u003e has four sub-orders\u003csup\u003e25\u003c/sup\u003e. Furthermore, the tentacles were rounded toward the superior direction, which we suggest contributes to lower resistance against the incoming water stream and allows the coronal organ to be present as a fringe on the inferior margin of the tentacles across all suborders. The findings regarding sub-tentacles appear inconsistent within the order of Stolidobranchia, as the family of \u003cem\u003eStylidae\u003c/em\u003e exhibits different shapes, suborders, abundance, and arrangement of the tentacles\u003csup\u003e24\u003c/sup\u003e. The dual abundance observed in different families within the order of Stolidobranchia leads us to hypothesize a secondary loss of sub-tentacles within this order. Further investigations of families within Stolidobranchia should be conducted to confirm this hypothesis.\u003c/p\u003e \u003cp\u003eThrough the segmentation of inner tentacular structures, we first outline a vascular network within the tentacles. To date, various papers have discussed the occurrence and characteristics of blood or hemolymph systems within tunicates. Consequently, we strongly endorse the concept of a closed or semi-closed vascular system based on the finely distributed capillary system of the tentacles, in alignment with the findings described in Konrad 2016\u003csup\u003e26\u003c/sup\u003e. The cross-sectional elliptical capillaries vary in diameter (height/width) from 79.0 \u0026micro;m/68.1 \u0026micro;m (\u0026plusmn;\u0026thinsp;8.40 \u0026micro;m/\u0026plusmn;9.65 \u0026micro;m) in the first-order tentacle to 33.3 \u0026micro;m/34.0 \u0026micro;m (\u0026plusmn;\u0026thinsp;2.83 \u0026micro;m/\u0026plusmn;4.23 \u0026micro;m) in the second order. Vessels of the third order were not measurable. The blood supply is generated via a single intratentacular vessel located inferiorly, which subdivides into one vessel per sub-tentacle (second order), alternating to the right and left. A similar single subdivision on alternating sides into the sub-tentacles of the third order was also observed.\u003c/p\u003e \u003cp\u003eIn contrast, an innervating structure was found on the superior side of the tentacle. Similarly, in the vessel distribution, the nerve formed sub-structures to the corresponding sub-tentacles, establishing the innervation of the coronal organ located on the inferior edge of the tentacles. Smaller capillaries emerging between the main branches and the sub-tentacles supply the nerve. The diameters of the first-order nerves are comparable in width, measuring 62.4 \u0026micro;m (\u0026plusmn;\u0026thinsp;11.60 \u0026micro;m) and height, 82.53 \u0026micro;m (\u0026plusmn;\u0026thinsp;8.45 \u0026micro;m), to the respective blood vessels of a similar order.\u003c/p\u003e \u003cp\u003eThe tunic of ascidians shows considerable variation across different orders, suborders, and families of ascidians. For example, in \u003cem\u003eCiona intestinalis\u003c/em\u003e, the tunic consists of a gelatinous and thin tissue\u003csup\u003e27\u003c/sup\u003e, whereas in \u003cem\u003eHalocynthia spp.\u003c/em\u003e, it has a leatherier consistency\u003csup\u003e28\u003c/sup\u003e. Notably, even within the same genus, such as \u003cem\u003eHalocynthia spp.\u003c/em\u003e, distinct differences in the tunic\u0026rsquo;s structure have been observed\u003csup\u003e29\u003c/sup\u003e. The cuticular sheds found in the tunic of \u003cem\u003eH. papillosa\u003c/em\u003e feature spikes that are absent in the closely related species \u003cem\u003eH. roretzi\u003c/em\u003e. When the animal contracts, these cuticular sheds form a strong armor that conceals the softer underlying tissues. Furthermore, our findings reveal, for the first time, a significant presence of autofluorescent tissue in solitary tunicates within the cuticular sheds (CS). This autofluorescence is highly concentrated in the cuticular sheds of the tunic, effectively obscuring all non-fluorescent parts of the tissue while the animal is contracted (see Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG, H).\u003c/p\u003e \u003cp\u003eStudies comparing tunics of different species have been limited, as the tunics of the well-known model organisms within the ascidians do not appear to contain any unique properties. In \u003cem\u003eH. papillosa\u003c/em\u003e, the first investigations of the tunic were carried out in the 1980s\u003csup\u003e10,11\u003c/sup\u003e, while the most recent findings on the tunic structure of \u003cem\u003eH. papillosa\u003c/em\u003e date back to the 1990s\u003csup\u003e12,13,30\u003c/sup\u003e. Thus, they no longer represent the current state of research and technology. In the present study, anatomical examinations were performed to analyze the structure and alignment of tunic fibers, along with a detailed investigation of the CS. The findings revealed that the autofluorescent CS, with an average diameter of approximately 500 \u0026micro;m in the animal's relaxed state, contains small gaps of up to 50 \u0026micro;m that do not exhibit autofluorescence. In the contracted state, however, these spaces are closed by an overlay of the CS, which completely conceals the deeper soft tissue and results in a seemingly uninterrupted fluorescent image of the outer tunic. In the approximately 300 \u0026micro;m-long conical cuticular spines, cavities were visible beneath the fluorescent cuticle layer. These cavities contained a chain of fluorescent particles arranged like beads on a string, which are transported up and/or down the spine\u0026rsquo;s center by fiber bundles. Similar observations have been reported in the embryogenesis of the same species, where granules were found along fiber bundles in the developing tunic two days after morphogenesis\u003csup\u003e13\u003c/sup\u003e. Unfortunately, the static image could not reveal the transport direction and requires further investigation.\u003c/p\u003e \u003cp\u003eUsing HiTT imaging combined with DAPI staining, we could also show that tunicin forms spiralized fibers to create a support structure for the cuticular sheds. This spiraling was visible only within the subcuticular zone (SC), which lies directly beneath the CS and originates from the parallel-arranged tunicin fibers. We suggest that this configuration allows the fiber bundles to extend through the center of the spiral, reaching the tip of the CS. The stained DAPI nuclei of the tunicin fibers also illustrated the dense spiral layering of the fibers in the SC area.\u003c/p\u003e \u003cp\u003eAdditional FTIR investigations were conducted on a thin slice of tunic. Previous studies on tunic slices of the closely related species \u003cem\u003eH. roretzi\u003c/em\u003e and \u003cem\u003eC. intestinalis\u003c/em\u003e showed similar results regarding tunicin detection in the central layers of the tunic\u003csup\u003e29,31\u003c/sup\u003e. Minor differences in the overall appearance of the FTIR output between \u003cem\u003eH. roretzi\u003c/em\u003e and the current study likely stemmed from slightly different scanner settings. Nevertheless, the current spikes in the range of 1060 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e to 1035 cm\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e are consistent with cellulose determinations previously published\u003csup\u003e21,29\u003c/sup\u003e. As anticipated from past light microscopy studies, the outer and inner layers exhibit distinct differences in the tunicin/cellulose spectrogram and show complex superimpositions of other tissue types over the tunicin that are not easily identifiable.\u003c/p\u003e \u003cp\u003eThe findings of this study provide a comprehensive anatomical examination of the tissues that make up the tunic, oral tentacles, and selected central nervous system structures in ascidians. A detailed analysis of these tissues is essential for understanding the ecological significance of ascidians, which have long been recognized for their role in environmental studies due to their species diversity and widespread presence across various marine ecosystems\u003csup\u003e17,32,33\u003c/sup\u003e. Solitary ascidians, in particular, offer valuable insights into the effects of anthropogenic stressors, such as noise pollution\u003csup\u003e34,35\u003c/sup\u003e, rising temperatures\u003csup\u003e36\u003c/sup\u003e, and variations in water quality\u003csup\u003e37,38\u003c/sup\u003e. Previous research on \u003cem\u003eH. papillosa\u003c/em\u003e has established a correlation between elevated temperatures and stress responses, as evidenced by changes in heat shock protein expression and behavioral modifications\u003csup\u003e36\u003c/sup\u003e. Ascidians have also been acknowledged as essential bioindicators in environmental monitoring\u003csup\u003e38\u003c/sup\u003e. Since ascidians are filter feeders, their physiological condition, settlement behavior, and survival rates in coastal environments can serve as indicators of water quality, particularly in relation to pollution levels and nutrient availability.\u003c/p\u003e \u003cp\u003eThe present study highlights two structural adaptations that may enhance the ability to perceive sound or vibrations. Given the increasing anthropogenic noise pollution in the oceans\u003csup\u003e39\u003c/sup\u003e, especially in coastal regions and harbors where these organisms are predominantly found, there is an urgent need for comprehensive research using indicator species in this area.\u003csup\u003e40\u003c/sup\u003e Consequently, examining the anatomical features and sound perception capabilities of ascidians alongside the traditional indicator species (mammals\u003csup\u003e41\u0026ndash;44\u003c/sup\u003e, fish\u003csup\u003e45\u003c/sup\u003e, selected invertebrates\u003csup\u003e35,40\u003c/sup\u003e) is crucial for a broad view of the oceans' changing environments.\u003c/p\u003e \u003cp\u003eAdditionally, our findings indicate the necessity of conducting investigations not solely on selected model organisms\u003csup\u003e\u003cem\u003e17\u003c/em\u003e\u003c/sup\u003e. While established model organisms like \u003cem\u003eC. intestinalis\u003c/em\u003e offer significant advantages\u0026mdash;attributed to their known whole genome sequencing data, the availability of genetically modified strains, and the remarkable transparency of their tunic\u003csup\u003e46\u003c/sup\u003e\u0026mdash; it is essential to acknowledge that even closely related species within the same phylogenetic family may display considerable anatomical variations, leading to functional and behavioral differences.\u003c/p\u003e \u003cp\u003eAnatomical investigations and an enhanced understanding of the cerebral system in various ascidians are also greatly significant. Previous studies reveal findings of rare chemical compounds and secondary metabolites unique to ascidians, many of which demonstrate toxic properties\u003csup\u003e47,48\u003c/sup\u003e or possess anti-tumor potential, making them highly valuable for medical research and drug development\u003csup\u003e49,50\u003c/sup\u003e. Beyond biomedical research, evolutionary interest in ascidians stems from their larvae, which serve as an optimal model for studying early chordate development. Their developmental features closely resemble those of vertebrates while retaining the genomic simplicity of invertebrates\u003csup\u003e51\u003c/sup\u003e, providing insights into the ancestral origins of chordates\u003csup\u003e11,52\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eIn addition to the general importance of understanding the differences in anatomical structures among various ascidian species, as previously explained, it is crucial to recognize the role of \u003cem\u003eH. papillosa\u003c/em\u003e as a species endemic to the Mediterranean Sea and Eastern Atlantic\u003csup\u003e53\u003c/sup\u003e. Despite its endemism, it is widely distributed and can be regarded as one of the most significant solitary ascidians in the Mediterranean Sea in terms of biomass. It holds economic value due to its popularity among scuba divers and plays a vital ecological role in benthic communities, particularly within coralligenous assemblages\u003csup\u003e54\u003c/sup\u003e. As a non-selective filter feeder, \u003cem\u003eH. papillosa\u003c/em\u003e consumes a broad range of microorganisms, including heterotrophic bacteria, phytoplankton, and ciliates, effectively regulating phytoplankton levels and contributing to water clarity\u003csup\u003e55\u003c/sup\u003e. Its filtration activity also establishes it as an important carbon sink, helping to mitigate eutrophication in marine ecosystems. Due to its sensitivity to environmental disturbances, such as sediment resuspension caused by diving activities, \u003cem\u003eH. papillosa\u003c/em\u003e serves as a valuable bioindicator for assessing the health of the benthic zones\u003csup\u003e54\u003c/sup\u003e. A decline in its population or size may indicate ecological stress before such stress becomes apparent at higher levels of the food web.\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eThis study underscores the limitations of concentrating detailed anatomical investigations exclusively on model species within a taxon. To achieve a more comprehensive understanding, well-established imaging and histological techniques should also be utilized for species that, while not widely distributed, play essential roles within their ecosystems. Integrating anatomical and physiological data into extensive genetic databases (e.g., GenBank) has already gained acceptance as standard practice. Biological sciences should further enhance the connection between anatomical and physiological findings with existing nucleotide and protein databases to develop a more holistic understanding of species and their ecological interactions. This strategy will become increasingly important as environmental changes quicken, resulting in shifts in species distributions and range expansions.\u003c/p\u003e"},{"header":"Materials and Methods","content":"\u003cdiv id=\"Sec6\" class=\"Section2\"\u003e \u003ch2\u003eStudy organism\u003c/h2\u003e \u003cp\u003e \u003cem\u003eHalocynthia papillosa\u003c/em\u003e is a common solitary ascidian found in the Mediterranean Sea. According to the Society for Laboratory Animal Science (GV-SOLAS), no animal testing application was necessary since the animal used is an invertebrate (non-cephalopod). However, the experiments were conducted only by individuals trained and educated in animal experimentation (EU function A). All experiments were performed to the best of our knowledge and in a manner intended to minimize stress and suffering to the animals. All invasive procedures were carried out only after anesthesia.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eAnimal Transportation\u003c/h3\u003e\n\u003cp\u003eThe organisms were retrieved from the Adriatic Sea near Pula on August 21st, 2022, at a depth ranging from 10 to 25 meters through scuba diving. After consultations with the Croatian Ministry of Environmental and Nature Protection, it was determined that a sampling permit was unnecessary for ascidians at the designated collection site near Pula (N44.83°, E13.84°). The collection process was carried out with precision, using a knife to carefully detach the substrate to which the organisms were adhered, thereby minimizing any potential injury to the specimens. Subsequent transportation to the aquatic laboratory at Ruhr-University Bochum (RUB) was achieved using plastic containers fitted with individual ventilation via air stones and provisions for continuous cooling to maintain a temperature of 16°C.\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eAnimal care and husbandry\u003c/h2\u003e \u003cp\u003eAscidians were placed in a FLUVAL Flex aquarium with a capacity of 123 liters, equipped with a continuous water flow system driven by an EHEIM compactON 1000 pump (15W), which has a throughput capacity of 1000 liters per hour. This setup was enhanced by an external chilling unit (TECO TK150, maintaining a temperature of 16°C) to ensure optimal thermal conditions. A comprehensive filtration system included an active carbon filter (AquaMedic carbolit 4mm), a biological filter using plastic balls (AquaMedic miniballs), and a ceramic filter (AquaNova NCR-0.5). Feeding protocols consisted of six daily feeds, providing a total of 20 ml (~ 2 x 10\u003csup\u003e8\u003c/sup\u003e cells) of \u003cem\u003eNannochloropsis salina\u003c/em\u003e per ascidian each day. Clean artificial seawater with a salinity of 37 to 39 PSU was supplied through a 30% water exchange every week. The light cycle was set to 12h/12h (light/dark) and was provided by an LED lamp (Aquasky, 6500K, 21W) integrated into the aquarium.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eLight microscopy\u003c/h3\u003e\n\u003cp\u003eA Leica EZ4 was used for light microscopic examinations. The selected lens configuration permits magnifications ranging from 8x to 35x, enabling various illumination techniques, including transmitted light, top light, and side light. This flexibility provided a thorough visualization of the diverse tissue specimens. The light microscope primarily captured consecutive images depicting tissues and dissections.\u003c/p\u003e\n\u003ch3\u003eMRI\u003c/h3\u003e\n\u003cp\u003eMagnetic resonance imaging (MRI) was conducted \u003cem\u003eex vivo\u003c/em\u003e using a 9.4 T horizontal small animal scanner (\u003cem\u003eBioSpec® 94/20 USR\u003c/em\u003e, Bruker BioSpin GmbH \u0026amp; Co. KG, Germany), equipped with the \u003cem\u003eB-GA12S HP\u003c/em\u003e gradient system (Bruker BioSpin GmbH \u0026amp; Co. KG, Germany) at the Leibnitz Institute for Neurobiology in Magdeburg, Germany. A \u003csub\u003e1\u003c/sub\u003eH transmit-receive volume coil with an inner diameter of 40 mm (Bruker BioSpin GmbH \u0026amp; Co. KG, Germany) was utilized for all measurements. The 4% paraformaldehyde (PFA) pre-fixed \u003cem\u003eH. papillosa\u003c/em\u003e was incubated in Fomblin® (a hydrogen-free, high-performance precision mechanics pump oil) for two hours to enhance the contrast between the specimen and the surrounding fluid. Based on preliminary experimental testing (not shown here), T2-weighted multi-slice multi-echo (MSME) sequences were found to be the most effective method for visualizing the ascidian.\u003c/p\u003e \u003cp\u003eBefore the anatomical scans were conducted, a B\u003csub\u003e0\u003c/sub\u003e-map was acquired (TR 15 ms, four avg.; image matrix 64’, FoV 40x40x40 mm\u003csup\u003e3\u003c/sup\u003e). The following imaging parameters were used: Sequence type: MSME sequence; TR: 13921.8 ms; TE: 16.1/80.5/187.7 ms; Echo averages: 2/10/10; transverse imaging (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA/B) with 25 contiguous slices of 0.5 mm; sagittal imaging (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC/D) with 55 contiguous slices of 0.5 mm; Image size: 3.2x3.8 cm²; Matrix size: 320x380 (resolution: 100x100 µm²); Scan time for transverse images: 0h33m44s; Scan time for sagittal images: 1h28m10s. Isotropic images for 3D visualization (supplemental video 1) were acquired with the same sequence and parameters, but with a matrix size of 320x380 (resolution: 250 x 250 x 250 µm³) and a scan time of 20 h 41 min. Contrast adjustments of the images were performed using ImageJ, and measurements of dimensions were obtained with Osirix®. 3D visualization and explosion videos were produced with VGStudio Max (2024.4).\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eHiTT\u003c/h2\u003e \u003cp\u003eHigh-throughput tomography (HiTT) was conducted at the European Molecular Biology Laboratory (EMBL) beamline P14, utilizing the Petra III storage ring at the Deutsches Elektronen-Synchrotron (DESY) in Hamburg\u003csup\u003e56\u003c/sup\u003e. Given the constraints imposed by a field of view (FOV) limited to a width of 1.3 mm, the dissection of the ascidian was undertaken to isolate specific sections of interest. These included the oral tentacles, the complete nerve between the two siphons, a 1 mm punch biopsy of tunic tissue near the oral siphon, and muscle strands between the siphons. The isolated segments were subsequently fixed in 4% PFA overnight, followed by a series of increasing alcohol concentrations according to the protocol established by Zhanmu et al. (2020)\u003csup\u003e57\u003c/sup\u003e to achieve a terminal concentration of 99.8% ethanol, thereby enhancing image contrast in the HiTT data. After preparation, the samples were carefully placed into a 200 µm pipette tip, sealed at the end, fixed to a magnetic goniometer base (MiTeGen Type B5), and covered with the corresponding magnetic lid. The samples were loaded into SPINE pucks and then placed into a sealed sample dewar maintained at room temperature within the beamline hutch. A computer-controlled robotic arm (MARVIN\u003csup\u003e58\u003c/sup\u003e) was used to mount the samples onto the diffractometer. The optimal scanning position was precisely determined using an integrated light microscope within the diffractometer setup (Arinax MD3). The X-ray energy at beamline P14 is adjustable within the range of 7–30 keV, with our measurements conducted at 12.7 keV. A 10-fold magnifying objective was used for each scan, resulting in an effective voxel size of 650 nm. Samples were imaged at four different propagation distances of 73 mm, 77 mm, 83 mm, and 92 mm. For each distance, a total of 1810 projection images were obtained over an 181° rotation angle, along with 100 additional flat-field frames\u003csup\u003e56\u003c/sup\u003e. The total acquisition time for all four distances was 136 seconds. Following the acquisition phase, reconstruction was completed fully automatically within one minute using the in-house TOMO-CTF software package. Phase retrieval was performed using the contrast-transfer function approach, employing a beta-delta ratio of 0.1. In the case of larger samples, a tiled acquisition of multiple adjacent data sets was performed. Reconstructed volumes were stitched together later using NRStitcher\u003csup\u003e59\u003c/sup\u003e. Data analysis, including segmentation and 3D visualization, was conducted using VG Studio Max (Version 2022.4, 64-bit) following the methodologies outlined in Albers et al. (2024)\u003csup\u003e56\u003c/sup\u003e.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eConfocal microscopy\u003c/h2\u003e \u003cp\u003eConfocal microscopy was performed at Ruhr University Bochum (RUB) using a Leica TCS SP5 system. For this analysis, the tunic tissue specimens were fixed in a 4% paraformaldehyde (PFA) solution and subsequently embedded in Tissue-Tec®. Tissue slicing for visualizing cuticular sheds was carried out with a CM 3050 S microtome, producing sections with a thickness of 50 µm. The imaging process involved positioning the tissue slices, which were affixed to super frost sample holders and covered with a cover slip, in an inverted orientation within the microscope. A 40x objective lens was utilized for imaging CS sections, while a 63x oil immersion lens provided enhanced magnification. To generate three-dimensional maximum intensity projections of the spines with significant depth and minimal background noise, virtual slice stacks comprising up to 101 optical planes were recorded. Additional CS sections were imaged in a non-fixed state to further confirm the absence of autofluorescence induced by PFA fixation. All acquired images relied solely on the tissue's inherent autofluorescence without introducing any artificial fluorescence. The GFP laser used for imaging was a DPSS 561.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eThunder\u003c/h2\u003e \u003cp\u003eThunder imaging was conducted at Ruhr-University Bochum (RUB) using a Leica microscope (model M205 FCA) equipped with a DFC9000 GT camera. This advanced microscopy system offers a magnification range from 0.75x to 16.0x, facilitating extensive imaging applications. Thunder imaging techniques were employed to create whole-animal images and perform initial assessments of auto-fluorescence within the CS at low magnification in the living specimen (\u003cem\u003ein vivo\u003c/em\u003e). Throughout the study, the straightforward microscope design proved most suitable for two-dimensional imaging with DAPI staining, as light and filters can be easily adjusted. Additionally, conventional mounting fluids were replaced with Roti®-Mount FluorCare DAPI solution (Roth) for tissue sample preparation, allowing for image acquisition with a high signal-to-noise ratio.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eFTIR spectroscopy\u003c/h2\u003e \u003cp\u003eFourier-transform infrared spectroscopy was performed at the Chair of Applied Electrodynamics and Plasma Technology (AEPT) at Ruhr University Bochum, Germany, using a Bruker Hyperion 3000 equipped with an Infinity 1 video camera. The tunic was sliced immediately after anesthesia using a vibratome (Leica VT 1200) without any fixing detergents in salt water (38 PSU). The resulting slice thickness was calibrated to 10 µm. The specimens were promptly transferred to silicon wafers for drying following the slicing procedure. The samples underwent overnight incubation before measurement. In a nitrogen-precooled scanning environment, the scanning process utilized an LN-MCT-D316-025 detector with a total scanning duration of 14 minutes and 17 seconds. It included 32 sample scans and 32 background scans in the microscopic scanning position.\u003c/p\u003e \u003c/div\u003e "},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eData availability statement\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets generated during and/or analyzed during the current study are available from the corresponding author upon request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgments\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eHiTT data were collected on EMBL Beamline P14 on the Petra III synchrotron in Hamburg. We thank Diving Pula for support with logistics.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eL.H.: Hypothesis generation, conceptualization, literature search, methodology, data scanning, image analysis, data evaluation and analysis, drawing, manuscript writing. J.A.: Methodology, data scanning, image analysis, data evaluation, and analysis. A.M.: Methodology, data scanning, image analysis, data evaluation, and analysis. E.D.: Methodology, data scanning. I.S.: Methodology S.H.: Conceptualization, funding acquisition. J.G.: Hypothesis generation, conceptualization, methodology, data scanning, image analysis, manuscript writing. M.H.: Hypothesis generation, conceptualization, methodology, image analysis, data evaluation and analysis, manuscript writing.\u003c/p\u003e\n\u003cp\u003eAll authors reviewed the manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEthics declaration\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAccording to the Society for Laboratory Animal Science (GV-SOLAS), no animal testing application was needed as the used animal is an invertebrate (non-cephalopod).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAdditional Information\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eAhyong, S.\u003cem\u003e et al.\u003c/em\u003e (WoRMS Editorial Board, 2025).\u003c/li\u003e\n\u003cli\u003eDelsuc, F., Brinkmann, H., Chourrout, D. \u0026amp; Philippe, H. 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(European Molecular Biology Laboratory, EMBL, Hamburg Unit, JACoW, 2018).\u003c/li\u003e\n\u003cli\u003eMiettinen, A., Oikonomidis, I. V., Bonnin, A. \u0026amp; Stampanoni, M. NRStitcher: non-rigid stitching of terapixel-scale volumetric images. \u003cem\u003eBioinformatics\u003c/em\u003e \u003cstrong\u003e35\u003c/strong\u003e, 5290-5297 (2019). https://doi.org/10.1093/bioinformatics/btz423\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"nature-portfolio","isNatureJournal":true,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"","title":"Nature Portfolio","twitterHandle":"","acdcEnabled":false,"dfaEnabled":false,"editorialSystem":"ejp","reportingPortfolio":"","inReviewEnabled":true,"inReviewRevisionsEnabled":false},"keywords":"Neuroanatomy, MRI, HiTT, Synchrotron, Autofluorescence, H. papillosa","lastPublishedDoi":"10.21203/rs.3.rs-6701261/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6701261/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eCurrent understanding of the anatomical structures of ascidians remains limited, particularly in regionally confined but abundant species such as \u003cem\u003eHalocynthia papillosa\u003c/em\u003e from the Mediterranean Sea. This study employed multimodal imaging techniques, including Light, Thunder, and fluorescent confocal microscopy, to investigate selected neural structures and the tunic of \u003cem\u003eH. papillosa \u003c/em\u003eat various magnifications. We utilized advanced 3D imaging methods, such as Magnetic Resonance Imaging (MRI) and High-Throughput Tomography (HiTT) at a synchrotron beamline. Additionally, chemical analysis using Fourier-Transformation Infrared-Spectroscopy (FTIR) was conducted. Our 3D and isotropic high-resolution renderings revealed significant structural differences in this species compared to others, notably the absence of a cerebral thickening. HiTT imaging identified three distinct suborders of oral tentacles, each with its own innervation and blood supply. Using Thunder and fluorescent Confocal microscopy, we also documented autofluorescence in ascidian cuticular sheds for the first time \u003cem\u003ein vivo\u003c/em\u003e and \u003cem\u003eex vivo \u003c/em\u003ein greater detail. Furthermore, HiTT X-ray imaging of the tunic revealed a spiralized structure emerging from the tunicin layers, which could be verified in this species for the first time using FTIR. The high-resolution, state-of-the-art imaging techniques presented in this study establish a strong foundation for future studies on \u003cem\u003eH. papillosa \u003c/em\u003eand other solitary ascidians, and highlight the need to expand research beyond traditional model species.\u003c/p\u003e","manuscriptTitle":"New insights into unique anatomical structures of the ascidian Halocynthia papillosa obtained by multimodal imaging","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-10-31 13:02:37","doi":"10.21203/rs.3.rs-6701261/v1","editorialEvents":[],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"communications-biology","isNatureJournal":true,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"commsbio","sideBox":"Learn more about [Communications Biology](http://www.nature.com/commsbio/)","snPcode":"","submissionUrl":"","title":"Communications Biology","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"ejp","reportingPortfolio":"Communications Series","inReviewEnabled":true,"inReviewRevisionsEnabled":false}}],"origin":"","ownerIdentity":"1249d673-3733-48c5-8c5b-5a5566f24820","owner":[],"postedDate":"October 31st, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[{"id":49462528,"name":"Biological sciences/Zoology/Animal physiology"},{"id":49462529,"name":"Biological sciences/Biological techniques/High-throughput screening"},{"id":49462530,"name":"Biological sciences/Biological techniques/Imaging/3-D reconstruction"},{"id":49462531,"name":"Biological sciences/Biological techniques/Imaging/Fluorescence imaging"},{"id":49462532,"name":"Biological sciences/Biological techniques/Imaging/Magnetic resonance imaging"}],"tags":[],"updatedAt":"2026-04-23T07:22:48+00:00","versionOfRecord":{"articleIdentity":"rs-6701261","link":"https://doi.org/10.1038/s42003-026-10102-5","journal":{"identity":"communications-biology","isVorOnly":false,"title":"Communications Biology"},"publishedOn":"2026-04-22 04:00:00","publishedOnDateReadable":"April 22nd, 2026"},"versionCreatedAt":"2025-10-31 13:02:37","video":"","vorDoi":"10.1038/s42003-026-10102-5","vorDoiUrl":"https://doi.org/10.1038/s42003-026-10102-5","workflowStages":[]},"version":"v1","identity":"rs-6701261","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6701261","identity":"rs-6701261","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}