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Article

New Clavelina (Ascidiacea) Species from the Bahamas †

by
Xavier Turon
1 and
Susanna López-Legentil
2,*
1
Department of Marine Ecology, Centre d’Estudis Avançats de Blanes (CEAB-CSIC), Accés Cala Sant Francesc 14, 17300 Blanes, Spain
2
Department of Biology and Marine Biology, Center for Marine Science, University of North Carolina Wilmington, 5600 Marvin K. Moss Lane, Wilmington, NC 28409, USA
*
Author to whom correspondence should be addressed.
†
zoobank: urn:lsid:zoobank.org:pub:3A916520-9058-42F5-89B0-B76D2C1DF491, urn:lsid:zoobank.org:act:42CD0FE3-B7AF-4D99-A45A-5090A3B120B9; urn:lsid:zoobank.org:act:CD8E1435-9ACD-438B-A9AB-B4C798EBB730; urn:lsid:zoobank.org:act:88FBC552-FA0B-4FB1-9271-1FC5DD2DE4A9.
Taxonomy 2024, 4(3), 661-679; https://doi.org/10.3390/taxonomy4030034
Submission received: 1 August 2024 / Revised: 29 August 2024 / Accepted: 30 August 2024 / Published: 3 September 2024
Browse Figures
Versions Notes

Abstract

:
The ascidian fauna of the Bahamas remains grossly understudied. Here, we examined specimens of the genus Clavelina collected from four Bahamian islands using morphological observations and genetic barcoding. Only three species of Clavelina have been previously reported in the Caribbean: C. picta, C. oblonga, and C. puertosecensis. Here, we report C. picta and three species new to science: C. rochae, C. pawliki, and C. erwinorum. C. picta and C. pawliki were found in the northernmost island surveyed, while C. rochae and C. erwinorum were particularly prevalent on the southeastern Bahamian islands. A complete review of genetic barcoding data and morphological characters of accepted Clavelina species was performed. The unique combination of in vivo coloration, morphological characteristics, haplotypes, and species distribution supported the establishment of three new species, significantly adding to the diversity of the genus Clavelina in the Caribbean.

1. Introduction

The Bahamas islands are located within the Lucayan Archipelago in the Atlantic Ocean, although they are commonly considered part of the Caribbean. The archipelago comprises nearly 700 islands and cays, most of which are uninhabited [1]. The islands are crossed by the Gulf Stream and Antilles Current, two major warm ocean currents (see [2]). The Gulf Stream originates in the Gulf of Mexico and circulates between Florida and the northern Bahamian islands. The Antilles Current flows westward across the Atlantic from Africa and passes the outer Bahamas. The Antilles current moves northward during the summer, causing higher temperatures in the Northern Bahamas [3]. During the winter months, the current shifts southward, warming the southern islands [3]. Both currents influence the marine communities around the islands, with most of the current biota in the Bahamas being of Caribbean origin [4].
The ascidian (Chordata; Tunicata) fauna of the Bahamas is poorly known, with only a handful of studies reporting a few species [5]. Wahl [6] characterized epibionts on 15 species, seven of which were only identified at the genus level (Didemnum). Later, López-Legentil et al. [7] described the cyanobacterial diversity in multiple specimens from three Bahamian didemnids. Sealey and Black [8] recently listed seven ascidian species from the Exuma Cays Land and Sea Park (the first marine reserve in the Bahamas). Besides these studies, a few others have occasionally utilized an ascidian species collected from the Bahamas to address specific research topics (e.g., [9], chemical defenses).
The genus Clavelina (Aplousobranchia; Clavelinidae) currently comprises 46 species and is globally distributed [10]. In the Caribbean, three species of Clavelina have been observed: C. oblonga Herdman, 1880; C. puertosecensis Millar and Goodbody, 1974; and C. picta (Verrill, 1990) [5,11,12,13,14,15]. To our knowledge, only C. picta has been reported in the Bahamas. Svane and Young’s [16] study on the ascidian larvae included the first photograph of a colony, and the species was later reported by Wahl [6] and Sealey and Black [8].
Clavelina is one of the few ascidian genera where species can be identified in living animals using their external characteristics, such as zooid arrangement and color patterns [17,18]. After fixation, colors usually fade away, and internal zooid characteristics (often hindered by contraction and preservation) must be examined. Thus, combining in vivo images with detailed descriptions of zooid morphological characters is mandatory for taxonomic work in this group. Recently, DNA barcoding of eukaryotes and free access to genetic databases have facilitated the identification of species by a broader range of researchers. Here, we combined field observations, morphological analyses, and DNA barcoding to identify four species of Clavelina from the Bahamas, three of which are new to science.

2. Materials and Methods

2.1. Samples

Samples of Clavelina species were photographed and collected from Sweeting’s Cay (Grand Bahama), Stirrup Cay, Plana Cay, and San Salvador by SCUBA diving in 2008 and 2010 (Table 1, Figure S1) aboard the Research Vessel Seward Johnson in 2008 and the Research Vessel Walton Smith in 2010. In situ pictures of Clavelina species were also taken in the Bahamas in 2007 and were used to inform each species’ distribution (Table 1). In 2008 and 2010, after pictures were taken in situ, colonies were collected and brought aboard, where one piece of each colony was preserved in absolute ethanol and another piece was anesthetized by cold exposure in a freezer after the animal was relaxed [19]. Note that in some cases, relaxation after collection was not achieved. When water was mostly frozen, the animals were killed by adding a small volume of formaldehyde. They were then preserved in 4% formaldehyde for morphological observation and storage. At least ten zooids per colony were extracted from the tunic and examined for morphological characteristics. Dissections were performed with micro-scissors, and, when necessary, structures were stained with Masson’s hemalum to increase contrast. Samples were identified based on specialized literature (see Discussion). Voucher specimens were deposited at the Biological Collection Facility of the Center for Advanced Studies of Blanes (CEAB, CSIC).

2.2. DNA Extraction, Amplification, and Sequencing

Ethanol-preserved zooids were separated from the tunic under a stereomicroscope and stored in absolute ethanol at −20 °C until processed. Total DNA was extracted using the DNeasy Blood and Tissue kit (Qiagen). The primer set LCO1490 and HCO2198 [20] was used to amplify a fragment of the Cytochrome c Oxidase subunit I (COI) mitochondrial gene. Total PCR volume was 25 μL, including 1 μL of each primer (10 μM), 10 nmol of each dNTP, 1x Reaction Buffer (Ecogen), 5 units of BIOTAQTM polymerase (Ecogen), and 10 to 20 μg/mL DNA. Reactions were carried out in a GenAmp thermocycler (Applied Biosystems), with the following conditions: 5 min at 94 °C, followed by 35 cycles of 30 s at 94 °C, 30 s at 45 °C, and 1.5 min at 72 °C, and a final extension time of 10 min at 72 °C. PCR products were purified and sequenced at Macrogen, Inc. All sequences have been deposited in GenBank.

2.3. Phylogenetic Analyses

Raw sequence data were processed in Geneious v.R11 [21] and aligned using Clustal X [22] with a gap opening penalty of 28 and a gap extension penalty of 4. All available sequences for Clavelina and some sequences from the closely related genus Pycnoclavella were retrieved from GenBank to build the phylogenetic trees. The polycitorid species Cystodytes dellechiajei (GenBank acc. num. AY523063) was used as an outgroup taxon. Neighbor-joining (NJ) and maximum likelihood (ML) analyses were conducted in MEGA v. 10 [23,24]. The Tamura–Nei nucleotide substitution model was used for NJ analyses, and data were re-sampled using 10,000 bootstrap replicates [25]. For ML analyses, we used the GTR + I + G [26] model with substitution rates varying among sites according to an invariant and gamma distribution. We performed 1000 bootstrap replicates.

3. Results

3.1. Morphology and Species Descriptions

3.1.1. Notes on Morphology

We provide notes on some morphological aspects of clavelinids (which also apply to other aplousobranch ascidians) as, in our experience, they are often misunderstood and inadequately described or represented in specimens’ descriptions. One of these aspects is the twisting of the abdomen. In a typical clavelinid, the body is divided into a thorax, containing the branchial sac, and an abdomen, which houses the gut and the gonads. The dorsoventral orientation of the zooids (and hence the right–left axis) is determined by the thorax, with the endostyle marking the ventral region and the atrial siphon the dorsal one.
The esophagus opens near the endostyle at the narrow posterior end of the branchial sac, runs downward, and ends in a bulbous stomach. From the stomach, the intestine runs to the bottom of the abdomen, turns 180 degrees, and passes anteriorly to open at an anus placed dorsally in the lower part of the peribranchial cavity. In this configuration, which we call the straight abdomen (Figure 1A), the stomach is located ventrally. However, in many zooids, the abdomen twists clockwise (as seen from the thorax), causing the stomach to occupy a dorsal position (henceforth twisted abdomen, Figure 1B). Some aplousobranch species consistently have a twisted or a straight abdomen, so this character is likely genetically fixed within species (e.g., in many didemnids). In other aplousobranchs, such as clavelinids, the abdomen position is variable even within the same colony. We have observed twisted abdomens in well-relaxed clavelinid zooids and straight abdomens in strongly contracted zooids, so we do not believe that different abdomen orientations result solely from contraction. Abdomen orientation affects the distribution of all organs within it and should always be mentioned in species descriptions and figures. For instance, gonads in clavelinids lie on the right-hand side of the gut loop, but if the abdomen is twisted, the gonads lie on the left side. Likewise, the two main fiber bundles that run along the abdomen end in terminal ampullae on the left side of straight abdomens but on the right side of twisted ones.
A second morphological character of taxonomic value in clavelinids, often overlooked or misreported, is the configuration of the thoracic muscles. They comprise muscular bands (hereafter fibers) of varying thickness depending on their position, orientation, and contraction degree. The arrangement of these fibers affects how the thorax contracts. The most typical pattern features longitudinally oriented fibers (Figure 2A), with some or all of these fibers extending into the abdomen and forming two bundles that end in terminal ampullae in the posterior abdomen. Some of these fibers originate at the anterior region of the endostyle, others close to the oral siphon, and others dorsally around the neural ganglion and the atrial siphon. Formulae have been proposed to describe the origin of these muscles [27,28]. However, in some species, the fibers are oriented transversely (Figure 2B), usually starting at the endostyle and running towards the dorsal branchial side. Some fibers may end there or become inconspicuous, while others merge and move posteriorly close to the mid-dorsal line before entering the abdomen. Other species have a pattern where the more ventral fibers run obliquely from the oral siphon to the endostyle (Figure 2C), where they may become indistinct or merge into a few fibers that run posteriorly along both sides of the endostyle. In the species reported here, we have instances of all these configurations.

3.1.2. Clavelina picta (Verrill 1900)

References and synonymy:
Diazona picta [29], p. 591; Rhodozona picta [30], p. 335; Clavelina picta Berrill [31], p. 83; Clavelina picta [5], p. 138.
Collection codes: CEAB.ASC.001_A, CEAB.ASC.001_B, and CEAB.ASC.001_C.
GenBank accession number: PP891404.
Colonies can comprise tens of zooids (Figure 3A,B). Colonies are formed by a basal tunic from which digitations with groups of 4–6 zooids emerge. The abdomens are usually embedded in a common tunic, with the thoraces free and enveloped by their own tunic. The color of the colonies is distinctive, with an uninterrupted red-carmine ring that surrounds the oral aperture and runs down ventrally, following the endostyle (Figure 3A,B). The whitish neural ganglion is visible just outside the red band. There is a variable amount of scattered pigment of the same reddish color, particularly over the dorsal part of the thoraces, which otherwise have a whitish tinge (Figure 3B). White specks can make the oral siphon appear eight-lobed, but the margins of both siphons are smooth. The dorsal rim of the oral siphon is slightly elevated. A shallow dorsal infolding is visible on relaxed zooids along the length of the branchial sac. The visible part of the abdominal area is reddish. The colonies often intermingle with Clavelina pawliki (see below; Figure 3B).
The zooids (Figure 4A-C) are up to 20 mm long (measured from the topmost of the tunic). The tunic is cartilaginous in preserved material, while a soft inner layer surrounds the zooids in some colonies. The thorax has smooth-rimmed apertures, and the atrial siphon is subterminal and close to the oral siphon (Figure 4B). The thoracic musculature (Figure 4D–G) is formed by 4–5 fibers that run from the oral siphon towards the endostyle, 4–6 fibers that run from the oral area downwards, and 3–4 fibers that run from the dorsal area downwards. These fibers can be subdivided and anastomosed. The pattern of thoracic musculature corresponds to the ventral type in Figure 2C. The fibers that run towards the endostyle do not end there but instead run posteriorly and parallel to the endostyle before entering the abdomen (Figure 4E). In straight abdomens, a right and a left bundle of fibers enter the abdomen and run posteriorly, ending on the left side of the digestive loop and forming two papillae posterior to the stomach. The fibers on the right side cross ventrally to the other side (Figure 4G) and end in the papilla closer to the stomach, while the fibers on the left run down vertically and end in the papilla closer to the intestine. In twisted ones, the sides are interchanged.
There are ca. 20 simple tentacles arranged in several rings, with the longer ones occupying a posterior position. The biggest tentacle lies just above the neural gland and its oblique or vertical slit aperture. There are often copepods in the branchial sac. Depending on the size of the zooid, there are 16 to 24 rows of stigmata with up to 50 stigmata per half row (Figure 4B,H). The transverse vessels form a lamina between rows that originates elongated papillae at the dorsal midline (Figure 4H).
Abdomens can be straight, twisted, or half-twisted, even within the same colony. The stomach is quadrangular or elongated. Stomachs have marked ridges or only one marked fold (the typhlosole), depending on their fullness level (Figure 4B). The stomach lies some distance away from the bottom of the abdomen and is continued by a gut that loops and turns upward without distinct constrictions or divisions (Figure 4B). The anus opens in the lower part of the peribranchial cavity at the same level as the last rows of stigmata.
Gonads are found only in some examined zooids and lie on the right side (if the abdomen is straight) between the stomach and the lower gut loop. They are formed by a cluster of ova and smaller, whitish male follicles (Figure 4C). Embryos and larvae (up to 17) are brooded on the right side of the peribranchial cavity. The mature larvae measure up to 1 mm in trunk length, have separated ocellus and statocyte in the sensory vesicle, and have an anterior ventral stalk from which three adhesive papillae emerge: two dorsal and one ventral (Figure 4I). The coiled tail is lodged between the dorsal and ventral papillae.

3.1.3. Clavelina rochae sp. nov.

Collection codes: holotype: CEAB.ASC.009_A; paratypes: CEAB.ASC.009_B, CEAB.ASC.009_C, CEAB.ASC.009_D, and CEAB.ASC.009_E.
GenBank accession numbers: PP891405-6.
Colonies are often formed by only a few scattered zooids (Figure 3C,D). Zooids are mostly independent, sometimes linked by stolons and sometimes with the posterior part of the abdomens embedded in a common tunic (Figure 5A). The transparent tunic has a faint purple ring around the oral siphon (Figure 3C). In some instances, the pigment continues downwards over the endostyle and the dorsal line of the branchial sac. The branchial sac has a dorsal furrow. The thorax has an electric blue tinge of variable intensity, sometimes forming a secondary band posterior to the purple ring (Figure 3D). Other times, the purple ring was indistinct, and the blue hue was the primary color of the thoraces. The oral aperture is large; its diameter is close to the whole width of the zooids (Figure 3D).
Zooids (Figure 5B–D) are up to 23 mm long (measured with the tunic). The tunic is soft and sometimes has an internal softer layer that is hard to remove from the zooids. The thoracic musculature (Figure 5C) consists of oblique and vertical fibers: 2–5 going from the oral siphon towards the endostyle, 3–10 originating around the oral siphon area and running down the branchial sac, and 0–3 originating between the neural gland and the atrial siphon and running downwards vertically. The muscular arrangement corresponds to the ventral type in Figure 2C. The fibers reaching the endostyle bend posteriorly and run parallel to both sides of the endostyle before entering the abdomen and ending in two terminal ampullae on the lower gut loop’s left side (in straight abdomens). There are 12–14 tentacles in the oral siphon, the largest one placed over the neural gland and its vertical aperture slit. Fourteen to 19 stigmata rows are observed, with over 50 stigmata per half-row (Figure 5E). An elevated lamina between rows continues dorsally onto the wide, pointed dorsal languets.
There are both straight and twisted abdomens (Figure 5B) within the same colony. The stomach is quadrangular, globular, or elongated, often with conspicuous ridges and a marked typhlosole. The post-stomach has no constrictions, and the gut turns and runs anteriorly (Figure 5D). The anus opens at the posterior end of the peribranchial cavity. The gonads have a few (7–10) ova and numerous male follicles to the right of the posterior part of the gut loop in straight abdomens. Embryos and larvae (up to 16) are brooded on the right-hand side of the peribranchial cavity (Figure 5C). The mature larvae are ca. 1 mm in trunk length, have separate ocellus and statocyte in the sensory vesicle, and a rare arrangement of the adhesive papillae: the two dorsal papillae arise from an anterior outgrowth, while the ventral papilla originates from a separated ventral stalk (Figure 5F). The coiled tail of the unhatched larva is lodged between the dorsal and the ventral papillae.
Etymology: The species name is dedicated to Rosana Moreira da Rocha for her numerous and key contributions to ascidian taxonomy and systematics.

3.1.4. Clavelina pawliki sp. nov.

Collection codes: holotype: CEAB.ASC:004_A; paratype: CEAB.ASC.004_B.
GenBank accession numbers: PP891402-3.
Colonies of this species can reach tens of zooids that are mostly independent of each other and are united by stolons or a basal layer of tunic (Figure 6A). It often forms colonies intermingled with Clavelina picta and sometimes the phlebobranch Ecteinascidia turbinata (Figure 3B,F). The zooids are slightly longer and slenderer than those of C. picta. The transparent tunic has a well-defined black or dark blue ring around the oral siphon (Figure 3E,F). This ring is interrupted dorsally at the neural ganglion and ventrally at the anterior tip of the endostyle (Figure 3F). The same dark pigment is more or less scattered over the posterior part of the thorax and may be abundant over the abdomens. Occasionally, the pigment forms a fine strip over the mid-dorsal line of the thorax and/or a double strip at each side of the endostyle. White pigment accumulates over the two siphonal apertures, on a line between both, and over the endostyle (Figure 3F). In relaxed zooids, a deep furrow is apparent over the dorsal midline of the branchial sac. Dark blue vesicles are scattered throughout the basal layer of the tunic and the stolons.
The zooids (Figure 6B) are up to 23 mm long (measured with the tunic). The tunic is firm. The apertures are smooth. The oral siphon is somewhat elevated in its dorsal rim, while the posterior rim of the atrial siphon has a small protruding lobe, often with white pigment. The deep-blue color around the oral siphon becomes a black ring in preserved animals (Figure 6B). The thoracic musculature is of the transverse type (Figure 2B), with the anterior-most five to seven bands of fibers originating around the siphons and running towards the dorsal midline. Moreover, 14–15 bands originate at the endostyle and cross towards the dorsal zone (Figure 6C). The endostylar side of these fibers is often bifurcated (Figure 6D). Dorsally, some fiber bands become indistinct, while others bend posteriorly, forming tracks that run parallel to both sides of the dorsal midline before entering the abdomen (Figure 6E). The transverse orientation of the muscle bands causes the thorax to become narrow and elongated in contracted zooids.
A membrane with a festooned rim links nine to 12 tentacles. The aperture of the neural gland is round or oval. The branchial sac has 20–21 rows of ca. 50 stigmata (per half-row). There are elevated transverse laminae between rows, dorsally forming wide and pointed dorsal languets (Figure 6F).
The abdomens examined for this species are primarily straight, but some twisted abdomens are also observed. The internal structures are hard to observe due to the amount of pigment. The stomach is elongated and quadrangular or irregular in section, with a typhlosole and a few ridges (likely due to contraction). There are no constrictions differentiating sections in the gut loop. The anus opens at the level of the last 2–3 rows of stigmata. Some zooids have incipient or well-developed gonads on the right side of the gut loop in straight abdomens. The gonads have a few rounded ova and many male follicles. Embryos and larvae are brooded on the posterior-right part of the peribranchial cavity (Figure 6B). When there are just a few embryos, they form a single row, and if many are present, they form an irregular clump (up to 13 embryos observed). The mature larvae are ca. 1 mm in trunk length, with an ocellus, a statocyte, and a well-marked oral siphon (Figure 6G). A peculiar feature of these larvae is a plate-like anterior outgrowth joined to the trunk by a ventral stalk. The three papillae, two dorsal and one ventral, arise from this plate. In preserved material, patches of white pigment are observed on the larval trunk, anterior process, and tail.
Etymology: The species is named in recognition of Joseph R. Pawlik, who coordinated and secured funding for the three expeditions to the Bahamas that led to this study and for his contributions to ascidian chemical ecology.

3.1.5. Clavelina erwinorum sp. nov.

Collection codes: holotype: CEAB.ASC.006_A; paratypes: CEAB.ASC.006_B and CEAB.ASC.006_C.
GenBank accession numbers: PP891398-401.
Colonies comprise mostly independent zooids (up to 20), with only the posterior part of the abdomen embedded in a common basal tunic (Figure 7A). The zooid coloration is distinctive (Figure 3G,H). Over an otherwise transparent tunic is a yellow ring of pigment around the oral siphon. The ring’s thickness is variable and is interrupted dorsally (at the neural ganglion area) and ventrally (at the tip of the endostyle). The ring is thickened dorsally, and in some cases, there is a second anterior partial ring in this area. Blue lines run down the endostyle and the mid-dorsal line of the thorax in all zooids. Another blue strip goes from the oral to the atrial siphon, enclosing the neural gland. This coloration pattern is complemented with specks of yellow pigment at both sides of the dorsal area of the thorax, sometimes inconspicuous (Figure 3G) but coalescing into two thick parallel bands in some colonies. However, the yellow ring surrounding the oral siphon and the fainter blue bands are always present. Some colonies of the species were previously observed and photographed by SLL in Key Largo, Florida (USA; Figure 3H). They have well-developed yellow patches in the thorax.
Zooids measure up to 14 mm in length (Figure 7B). The tunic is soft but consistent. The thoracic muscles are of the longitudinal type (Figure 2A), 3–6 originating at the endostyle, 3–6 at the oral siphon area, and 2–5 from the neural ganglion or the atrial siphon (Figure 7C,D). The oral siphon has 15–20 tentacles of different sizes. The aperture of the neural gland is a vertical slit or an oblique oval. There are 16–18 rows of stigmata, with up to 60 stigmata per half-row (Figure 7E).
Most zooids observed have twisted or half-twisted abdomens. The stomach is plicated or globular depending on the degree of contraction. A very short post-stomach gives rise to a gut that runs downwards before turning upwards. No clear divisions can be seen in the gut. The anus ends in the peribranchial cavity at the level of the last rows of stigmata. No developed gonads are present, but in some zooids, a sperm duct and incipient testicular lobes are observed on the gut loop’s left side (in twisted abdomens). One zooid has six embryos incubated in the posterior right side of the branchial sac, including a probably not fully developed larva (Figure 7F). The sensory vesicle cannot be observed, and the papillae arise from two ectodermal outgrowths, one ventral and one dorsal (the latter originating the two dorsal papillae). The larval tail runs between the ventral and dorsal papillae.
Etymology: The species’ name is in homage to Patrick M. Erwin for his pioneering research on the ascidian microbiome and Elenor Lopez Erwin for the countless hours spent listening to her parents talk about sea squirts and bacteria.

3.2. Phylogenetic Analyses

Partial COI sequences with a final alignment length of 591 bp were obtained for all four species: one for Clavelina picta, two for C. pawliki, four for C. erwinorum, and two for C. rochae (accession numbers PP891398-PP891406). The sequence obtained for C. picta formed a well-supported clade with a specimen of the same species collected in Cuba (bootstrap values >99%; Figure 8). C. picta was the sister group to a clade formed by the two C. rochae sequences (bootstrap values = 100%), and both C. picta and C. rochae formed a well-supported clade together (bootstrap values = 96%). The two sequences of C. pawliki were nearly identical (one-point mutation between them) and formed a strongly supported clade that was a sister group to a clade comprising Mediterranean sequences for C. lepadiformis, C. gemmae, and C. sabbadini. The clade formed by all sequences obtained for C. erwinorum was not clearly associated with any other Clavelina species, with C. arafurensis being the closest branching clade. The four sequences of C. erwinorum were slightly different, with a percent identity ranging from 96.1 to 97.5%. Phylogenetic analyses matched morphological observations and supported the taxonomic assignments made.

4. Discussion

The four species reported here can be easily distinguished in living colonies because of their distinctive color patterns. While the amount of pigment varies within species, the overall patterns are consistent. These morphotypes also correlate well with groups defined by COI sequence data, forming clusters that set these species apart from all previously barcoded Clavelina species. Colony structures ranged from completely embedded abdomens to species with free or almost free zooids, united only by a basal layer of tunic or stolons, in this order: C. picta, C. erwinorum, C. pawliki, and C. rochae. Note that this character can vary across colonies and even within the same colony, so several specimens must be observed.

4.1. Comparison of Caribbean Clavelina Species

In the Caribbean at large (including Bermudas), three species of Clavelina had been previously identified: C. oblonga, C. picta, and C. puertosecensis [5,12,13,32,33]. A fourth species, C. brasiliensis, had been described from the northern Brazilian shore [34] and was later identified from the French Guiana [35]. Table 2 summarizes the main characteristics of these four species and the three new species described here.
The species C. oblonga was described from the Bermudas, is the most widely reported species in the Western North Atlantic [13], and has been introduced in many tropical and temperate areas [36,37]. It is particularly abundant in mangrove roots, harbors, and protected shallow habitats [38,39]. Although we did not observe C. oblonga during our surveys, we did not visit the habitats where the species is commonly found and cannot rule out its presence in the Bahamas. C. oblonga forms large colonies with easily distinguishable whitish zooids, unlike any species reported here. C. puertosecensis is characterized by zooids with a uniform purple or deep blue coloration [13,40]. C. brasiliensis has much larger zooids (up to 7.5 cm) than those reported here and features a distinct larval type with anterior swelling [34].
Only Clavelina picta had previously been reported in the Bahamas [6], and the colonies observed here match previous descriptions, both externally and internally ([30] as Rhodozona picta; [5,31]; Table 2). To our knowledge, no recent description of this species has been published, and the distribution of the muscular bands in the thoraces (with the anterior ones reaching ventrally towards the endostyle and running posteriorly alongside it) has never been reported before. C. rochae shares some morphological features with C. picta (Table 2), such as the arrangement of the muscular bands and the general zooid morphology. However, it differs by having colonies with sparse, mostly free zooids rather than having embedded abdomens. The color pattern also differs, with electric blue hues of variable intensity in C. rochae and the reddish band around the oral siphon thinner and wider in diameter than in C. picta. Accordingly, the zooids in C. rochae do not taper appreciably at the anterior end but have a wide oral opening and are “jar-shaped”, while those of C. picta are more “bottle-shaped”. The larvae are also different, with an anterior outgrowth holding the two dorsal papillae in C. rochae and three papillae arising from a single anterior stalk in C. picta (as described in [31]). However, the larvae may appear similar when not fully mature. Genetically, the new species is also well differentiated from C. picta (87.5–87.9% identity) but forms a well-supported clade with it, indicating that both species are closely related.
Among the Clavelina species reported herein, only C. pawliki resembles morphologically another Caribbean species not observed in the Bahamas, C. puertosecensis (Table 2). Both species have a similar pattern of thoracic musculature and colony structure [12]; however, the coloration is markedly different, and the larvae of C. puertosecensis have a wide ampullar anterior process instead of the plate-like process of C. pawliki. The infolding of the dorsal wall of the branchial sac in C. puertosecensis, highlighted as unusual by Millar and Goodbody [12] but not reported by Monniot [32], was present in all species examined here. The infolding was only visible in vivo and was lost in preserved zooids due to contraction.
Table 2. Main morphological and genetic characteristics of the seven species of Clavelina present in the Central West Atlantic, including the three new species described here. Colony: Colony structure; Color: Main color features; Max. length: Maximal zooid length; #stigmata: Number of stigmata rows; #stigmata/half row: Number of stigmata per half row; Musc. bands: Arrangement of muscular bands; Abdomen: Arrangement of abdomen; Larva: Larval features; Closest clade: Closest clade resulting from the COI phylogenetic analysis; nd: No data available.
Table 2. Main morphological and genetic characteristics of the seven species of Clavelina present in the Central West Atlantic, including the three new species described here. Colony: Colony structure; Color: Main color features; Max. length: Maximal zooid length; #stigmata: Number of stigmata rows; #stigmata/half row: Number of stigmata per half row; Musc. bands: Arrangement of muscular bands; Abdomen: Arrangement of abdomen; Larva: Larval features; Closest clade: Closest clade resulting from the COI phylogenetic analysis; nd: No data available.
Clavelina SpeciesColonyColorMax. Length#Stigmata#Stigmata/Half Row aMusc. Bands bAbdomenLarvaClosest CladeReferences
C. brasiliensisZooids freend80 mm17–20ndVentralndAnterior lobed swellingnd[34,35]
C. erwinorumZooids mostly free, with stolons or basal tunicYellow ring around oral siphon, thickened dorsally14 mm16–18up to 60LongitudinalMostly twistedOne ventral and one dorsal anterior outgrowth bearing papillaeC. arafurensisThis article
C. oblongaAbdomens embedded, thoraces freeScattered whitish spots in the thorax25 mmca. 2050–60LongitudinalMostly twistedSingle antero-ventral stalk bearing the three papillaeC. moluccensis[5,31,37]
C. pawlikiZooids mostly free, with stolons or basal tunicDark blue ring around oral siphon23 mm20–21up to 50TransverseMostly straightPlate-like anterior outgrowth bearing the three papillaeC. sabbadini/gemmae/lepadiformisThis article
C. pictaAbdomens embedded, thoraces freeRed ring around oral siphon and red band over endostyle20 mm16–24up to 50VentralStraight and twistedSingle antero-ventral stalk bearing the three papillaeC. rochaeThis article
C. puertosecensisZooids free, joined by stolonsPurplish blue color with yellow pigment around the siphons20 mm12–1850–70TransverseMostly twistedStout anterior process bearing the papillaend[12,32]
C. rochaeZooids mostly free, with stolons or basal tunicPurple ring around oral siphon, electric blue tinge23 mm14–1950+VentralStraight and twistedAnterior outgrowth bears dorsal papillae, ventral papilla in separate stalkC. pictaThis article
(a) Measured in the middle of the thorax. (b) Longitudinal, transverse, and ventral refer to the disposition seen in Figure 2A, Figure 2B, and Figure 2C, respectively.

4.2. Comparison with Other Clavelina Species

Besides C. picta, C. puertosecensis, C. oblonga, and C. brasiliensis, 42 additional Clavelina species are listed in the World Ascidiacea Database [10]. The main morphological characteristics of these species were recently listed by Hasegawa and Kajihara [41]. We analyzed these descriptions and looked for shared features (e.g., colony structure, zooid size, number of stigmata rows) with the new species described herein. Once lookalike species were identified, we carefully checked their color patterns (when available) and main zooid and larvae characteristics. Some relevant observations are listed below.
Several species have yellow rings in the siphonal area reminiscent of the coloration of C. erwinorum. Among them, C. amplexa Kott, 2002 (described from Darwin, Australia) has a yellow patch at each side of the anterior part of the thorax but with a triangular shape [42], unlike the ring seen in C. erwinorum. C. amplexa also has far fewer stigmata in the branchial sac (ca. 20 per half-row). The Pacific species C. robusta Kott, 1990 has a yellowish or fluorescent green band around the oral siphon, but it is not ventrally interrupted as in C. erwinorum. C. robusta also has a band around the atrial siphon, and the overall color of the zooids is translucent-dark or bluish [28,43,44]. The zooids of C. robusta also have more muscular bands arising from the endostyle and fewer stigmata per half-row (20–24, [43]). C. cyclus Tokioka and Nishikawa, 1975 (described from Japan) also has a yellowish band around the oral siphon, but it is not interrupted dorsally, and the color pattern is different [45]. C. cyclus also has fewer stigmata per half-row in the branchial sac (up to 50; [17,46]).
A few other species of Clavelina have the transverse arrangement of muscle bands seen in C. pawliki and C. puertosecensis. C. coerulea Oka, 1934, has similar zooid morphology but differs in color pattern and lacks the plate-like anterior process in its larvae [17,47]. C. obesa Nishikawa and Tokioka, 1976 has oblique (rather than transverse) muscle bands, a different colony color, fewer muscles, and fewer stigmata rows than C. pawliki [17,43]. The Indo-Pacific C. arafurensis Tokioka, 1952, differs from C. pawliki in having almost completely embedded zooids, different color patterns, and a post-abdomen [44,48]. Another Indo-Pacific species, C. moluccensis (Sluiter, 1904), has a predominantly transverse muscle arrangement but with more muscle bands and a characteristic pattern of three blue spots between the siphons [49]. Unlike C. pawliki, C. moluccensis has a bulbous pre-stomach, and the intestine is divided into regions [28,38]. The Australian species C. nigra Kott, 1990, has transverse muscle bands, but they are more numerous, the zooids are uniformly dark, and the gut structure is also different.
Finally, the muscular arrangement found in C. picta and C. rochae, where the more ventral muscles originate from the oral siphon area and extend ventrally to the endostyle, has not been reported in other Clavelina species. This arrangement is more similar to that described in some Diazonidae species [28,44].
In conclusion, the combination of in vivo color patterns, morphological characters, and genetic data supports the establishment of the three newly described species, adding to the known variability of the genus Clavelina, particularly in Caribbean waters.

4.3. Geographic Distribution in the Bahamas

Regarding their distribution, C. picta and C. pawliki were observed in the northernmost surveyed location (Sweetings Cay, Grand Bahama), while C. rochae and C. erwinorum were observed on the southeastern Bahamian islands (Stirrup, West Plana, and San Salvador). This distribution appears to be determined by the short-lived nature of the ascidian larvae [16] and the two main currents in the region. The Gulf Stream originates in the Gulf of Mexico, flows through the strait of Florida and the northernmost Bahamian islands, and then continues along the eastern coastline of the United States. The strength and direction of this current likely prevent the larvae of C. picta and C. pawliki from reaching the southern Bahamian islands surveyed here. In contrast, the Antilles Current flows northward east of the Antilles and joins the Gulf Stream past the outer Bahamas. This current is unlikely to reach Grand Bahama, as the island is further west and is partially sheltered by Great Abaco on the east. However, although C. erwinorum was not observed in Sweetings Cay, it was observed in Key Largo, Florida. Thus, some degree of connectivity among populations must exist. Further sampling in Central America and the Gulf of Mexico could clarify whether the Caribbean current also plays a role in the species’ current distribution.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/taxonomy4030034/s1. Figure S1: Map of the Bahamas showing the geographic location of the four surveyed islands and cays (Sweetings, Stirrup, West Plana, and San Salvador).

Author Contributions

Conceptualization, X.T. and S.L.-L.; methodology, X.T. and S.L.-L.; formal analysis, X.T. and S.L.-L.; resources, X.T. and S.L.-L.; data curation, X.T. and S.L.-L.; writing and editing, X.T. and S.L.-L.; funding acquisition, X.T. and S.L.-L. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by a Systematics Research grant from the Linnean Society and the Systematics Association to SLL. Use of the UNOLS Research Vessel Walton Smith was supported by a grant from the US National Science Foundation (OCE-0550468 to Dr. Joseph R. Pawlik). Project MARGECH PID2020-118550RB from the Spanish Government (MICIU/AEI/10.13039/501100011033) provided partial funding.

Data Availability Statement

Voucher specimens were deposited at the Center for Advanced Studies of Blanes (CEAB, CSIC) Biological Collection Facility. DNA sequences are accessible through GenBank (accession numbers PP891398-PP891406).

Acknowledgments

We thank Mari Carmen Pineda and Paula Anglada for their help with sequencing. Patrick M. Erwin (UNCW) participated in the 2010 expedition and helped with sample collection.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Harris, D.R.; Albury, E.P.; Saunders, G. The Bahamas. Encyclopedia Britannica. 2024. Available online: https://www.britannica.com/place/The-Bahamas (accessed on 5 June 2024).
  2. Sealey, N.E. Bahamian Landscapes. An introduction to the Geography of the Bahamas, 2nd ed.; Media Enterprises: Nassau, Bahamas, 1994; p. 128. [Google Scholar]
  3. Shaklee, R.V. Weather and Climate, San Salvador Island, Bahamas; The Bahamian Field Station Ltd.: San Salvador, Bahamas, 1996; p. 67. [Google Scholar]
  4. Buchan, K.C. The Bahamas. Mar. Pollut. Bull. 2000, 41, 94–111. [Google Scholar] [CrossRef]
  5. Van Name, W.G. The North and South American ascidians. Bull Am. Mus. Nat. Hist. 1945, 84, 1–476. [Google Scholar]
  6. Wahl, M. Bacterial epibiosis on Bahamian and Pacific ascidians. J. Exp. Mar. Biol. Ecol. 1995, 191, 239–255. [Google Scholar] [CrossRef]
  7. López-Legentil, S.; Song, B.; Bosch, M.; Pawlik, J.R.; Turon, X. Cyanobacterial diversity and a new Acaryochloris-like symbiont from Bahamian sea-squirts. PLoS ONE 2011, 6, e23938. [Google Scholar] [CrossRef]
  8. Sealey, K.S.; Black, K. Looking back to see forward: Species diversity changes since 1980 on Bahamian reefs through diver photography. Acad. Biol. 2023, 1, 1.21. [Google Scholar] [CrossRef]
  9. Vervoort, H.C.; Pawlik, J.R.; Fenical, W. Chemical defense of the Caribbean ascidian Didemnum conchyliatum. Mar. Ecol. Prog. Ser. 1998, 164, 221–228. [Google Scholar] [CrossRef]
  10. Shenkar, N.; Gittenberger, A.; Lambert, G.; Rius, M.; Moreira da Rocha, R.; Swalla, B.J.; Turon, X.; Ascidiacea World Database. Accessed through: World Register of Marine Species. 2024. Available online: https://www.marinespecies.org/ (accessed on 5 June 2024).
  11. Millar, R.H. Some ascidians from the Caribbean. Stud. Fauna Curaçao Other Caribb. Isl. 1962, 59, 61–77. [Google Scholar]
  12. Millar, R.H.; Goodbody, I. New species of ascidian from the West Indies. Stud. Fauna Curaçao Other Caribb. Isl. 1974, 45, 142–161. [Google Scholar]
  13. Goodbody, I. Diversity and distribution of ascidians (Tunicata) in the Pelican Cays, Belize. Atoll Res. Bull. 2000, 480, 330–333. [Google Scholar] [CrossRef]
  14. Goodbody, I. The ascidian fauna of Port Royal, Jamaica I. Harbor and mangrove dwelling species. Bull. Mar. Sci. 2003, 73, 457–476. [Google Scholar]
  15. Palomino-Alvarez, L.A.; Rocha, R.M.; Simões, N. Checklist of ascidians (Chordata, Tunicata) from the southern Gulf of Mexico. ZooKeys 2019, 832, 1–33. [Google Scholar] [CrossRef] [PubMed]
  16. Svane, I.B.; Young, C.M. The ecology and behavior of ascidian larvae. Oceanogr. Mar. Biol. Ann. Rev. 1989, 27, 45–90. [Google Scholar]
  17. Nishikawa, T.; Tokioka, T. Contributions to the Japanese ascidian fauna XXIX. Notes on some Clavelinids from the Japanese waters. Publ. Seto Mar. Biol. Lab. 1976, XXIII, 63–82. [Google Scholar] [CrossRef]
  18. Monniot, C.; Monniot, F.; Laboute, P. Coral Reef Ascidians of New Caledonia; Orstom Editions: Paris, France, 2001; pp. 156–161. [Google Scholar]
  19. Turon, X. Estudio de las Ascidias de las Costas de Cataluña e Islas Baleares. Ph.D. Thesis, University of Barcelona, Barcelona, Spain, 1987. [Google Scholar]
  20. Folmer, O.; Black, M.; Hoeh, W.; Lutz, R.; Vrijenhoek, R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 1994, 3, 294–299. [Google Scholar]
  21. Kearse, M.; Moir, R.; Wilson, A.; Stones-Havas, S.; Cheung, M.; Sturrock, S.; Buxton, S.; Cooper, A.; Markowitz, S.; Duran, C.; et al. Geneious Basic: An integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics 2012, 28, 1647–1649. [Google Scholar] [CrossRef]
  22. Thompson, J.D.; Gibson, T.J.; Plewniak, F.; Jeanmougin, F.; Higgins, D.G. The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997, 25, 4876–4882. [Google Scholar] [CrossRef]
  23. Kumar, S.; Stecher, G.; Li, M.; Knyaz, C.; Tamura, K. MEGA X: Molecular evolutionary genetics analysis across computing platforms. Mol. Biol. Evol. 2010, 35, 1547–1549. [Google Scholar] [CrossRef]
  24. Stecher, G.; Tamura, K.; Kumar, S. Molecular evolutionary genetics analysis (MEGA) for macOS. Mol. Biol. Evol. 2020, 37, 1237–1239. [Google Scholar] [CrossRef] [PubMed]
  25. Felsenstein, J. Phylogenies and the comparative method. Am. Nat. 1985, 125, 1–15. [Google Scholar] [CrossRef]
  26. Tavaré, S. Some probabilistic and statistical problems in the analysis of DNA sequences. In Some Mathematical Questions in Biology—DNA Sequence Analysis; Miura, R.M., Ed.; American Mathematics Society: Providence, RI, USA, 1986; pp. 57–86. [Google Scholar]
  27. Tokioka, T.; Nishikawa, T. Contributions to the Japanese ascidian fauna XXX. Further notes on Japanese Clavelinids. Publ. Seto Mar. Biol. Lab. 1976, XXIII, 341–350. [Google Scholar] [CrossRef]
  28. Kott, P. The Australian Ascidiacea. Part 2, Aplousobranchia (1); Memoirs of the Queensland Museum: South Brisbane, QLD, Australia, 1990; Volume 29, pp. 1–298. [Google Scholar]
  29. Verrill, A.E. Additions to the Tunicata and Molluscoidea of the Bermudas; Transactions of the Connecticut Academy of Arts and Sciences: New Haven, CT, USA, 1900; Volume 10, pp. 588–594. [Google Scholar]
  30. Van Name, W.G. The Ascidians of the Bermuda Islands; Transactions of the Connecticut Academy of Arts and Sciences: New Haven, CT, USA, 1902; Volume XI, pp. 325–411. [Google Scholar]
  31. Berrill, N.J. Ascidians of the Bermudas. Biol. Bull. 1932, 62, 77–88. [Google Scholar] [CrossRef]
  32. Monniot, F. Ascidies littorales de Guadeloupe. II. V. Polycitoridae. Bull. Mus. Natl. Hist. Nat. 1983, 4, 949–962. [Google Scholar]
  33. Goodbody, I. Ascidians from Caribbean shallow water localities. Stud. Fauna Curaçao Other Caribb. Isl. 1984, LXVII, 62–76. [Google Scholar]
  34. Millar, R.H. Ascidians (Tunicata: Ascidiacea) from the northern and north-eastern Brazilian shelf. J. Nat. Hist. 1977, 11, 169–223. [Google Scholar] [CrossRef]
  35. Monniot, F. Ascidians (Tunicata) of the French Guiana Expedition. Zootaxa 2016, 4114, 201–245. [Google Scholar] [CrossRef]
  36. Rocha, R.M.; Kremer, L.P.; Fehlauer-Ale, K.H. Lack of COI variation for Clavelina oblonga (Tunicata, Ascidiacea) in Brazil: Evidence for its human-mediated transportation? Aquat. Invasions 2012, 7, 419–424. [Google Scholar] [CrossRef]
  37. Ordóñez, V.; Pascual, M.; Fernández, M. Turo When invasion biology meets taxonomy: Clavelina oblonga (Ascidiacea) is an old invader in the Mediterranean Sea. Biol. Invasions 2016, 18, 1203–1215. [Google Scholar] [CrossRef]
  38. Monniot, C. Les genres Archidistoma et Clavelina (Ascidiacea, Clavelinidae) dans le canal du Mozambique. Zoosystema 1997, 19, 193–209. [Google Scholar]
  39. Goodbody, I. The ascidian fauna of a Jamaican lagoon: Thirty years of change. Rev. Biol. Trop. 1993, 41, 35–38. [Google Scholar]
  40. Rocha, R.M.; Zanata, T.B.; Moreno, T.R. Keys for the identification of families and genera of Atlantic shallow water ascidians. Biota Netrop. 2012, 12, 269–303. [Google Scholar] [CrossRef]
  41. Hasegawa, N.; Kajihara, H. Graveyards of giant pandas at the bottom of the sea? A strange-looking new species of colonial ascidians in the genus Clavelina (Tunicata: Ascidiacea). Species Divers. 2004, 29, 53–64. [Google Scholar] [CrossRef]
  42. Kott, P. Ascidiacea (Tunicata) from Darwin, Northern Territory, Australia. In The Beagle, Records of the Museums and Art Galleires of the Northern Territory; Darwin, Northern Territory Museum of Arts and Sciences: Darwin, NT, Australia, 2002; Volume 18, pp. 19–55. [Google Scholar]
  43. Monniot, F.; Monniot, C. New collections of ascidians from the Western Pacific and Southeastern Asia. Micronesia 1996, 29, 133–279. [Google Scholar]
  44. Monniot, F.; Monniot, C. Ascidians from the tropical western Pacific. Zoosystema 2001, 23, 248–336. [Google Scholar]
  45. Tokioka, T.; Nishikawa, T. Contributions to the Japanese ascidian fauna XXVII. Some ascidians from Okinawa, with notes on a small collection from Hong Kong. Publ. Seto Mar. Biol. Lab. 1975, XXII, 323–341. [Google Scholar] [CrossRef]
  46. Nishikawa, T. Chordata. In Guide to Seashore Animals of Japan with Color Pictures and Keys; Nishimura, S., Ed.; Hoikusha: Osaka, Japan, 1995; Volume II, pp. 573–610. [Google Scholar]
  47. Oka, A. Uber Clavelina coerulea n.sp. die erste von Japan bekannt gewordene Clavelina. Proc. Imp. Acad. 1934, 10, 365–366. [Google Scholar] [CrossRef]
  48. Tokioka, T. The Fauna of Akkeshi Bay. 18. Ascidia; Publications from the Akkeshi Marine Biological Station: Hokkaido, Japan, 1952; Volume 1, pp. 1–24. [Google Scholar]
  49. Sluiter, C.P. Die Tunicaten der Siboga-Expedition. Part 1. Die socialen und holosomen Ascidien. Siboga Exped. 1904, 56a, 1–126. [Google Scholar]
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Figure 1. Schematic representation of a zooid with a straight abdomen (A) and a twisted abdomen (B). D: dorsal side; V: ventral side. Note that gonads lie on the right in (A) and the left in (B).
Figure 1. Schematic representation of a zooid with a straight abdomen (A) and a twisted abdomen (B). D: dorsal side; V: ventral side. Note that gonads lie on the right in (A) and the left in (B).
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Figure 2. Schemes of the main patterns of thoracic musculature observed. (A) The most common longitudinal pattern in Clavelina species is fibers that run posteriorly from the anterior end of the endostyle and the oral, neural, and atrial zones. (B) In the transverse pattern, fibers originate on the ventral side, running dorsally and then passing posteriorly. (C) In the ventral pattern, the ventral fibers closest to the endostyle run towards it and then enter the abdomen. Note that this figure is highly schematic. In the fixed zooids, fibers anastomose, bifurcate, and merge over their length.
Figure 2. Schemes of the main patterns of thoracic musculature observed. (A) The most common longitudinal pattern in Clavelina species is fibers that run posteriorly from the anterior end of the endostyle and the oral, neural, and atrial zones. (B) In the transverse pattern, fibers originate on the ventral side, running dorsally and then passing posteriorly. (C) In the ventral pattern, the ventral fibers closest to the endostyle run towards it and then enter the abdomen. Note that this figure is highly schematic. In the fixed zooids, fibers anastomose, bifurcate, and merge over their length.
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Figure 3. Images of Bahamian Clavelina species. (A) Colony of C. picta from Sweetings Cay photographed on 30 May 2008. (B) C. picta (asterisk), C. pawliki (triangle), and Ecteinascidia turbinata (arrow) growing together in Sweetings Cay, photo taken on 10 June 2008. (C,D) C. rochae from Plana Cay, photographed on 25 June 2007. (E) Colony of C. pawliki photographed at Sweetings Cay on 17 June 2007. (F) C. pawliki and E. turbinata growing together in Sweetings Cay, photographed on 10 June 2008. (G) A colony of C. erwinorum from San Salvador photographed on 5 June 2008. (H) C. erwinorum photographed in Key Largo, Florida (USA) on 14 November 2006. Scale bar 1 cm.
Figure 3. Images of Bahamian Clavelina species. (A) Colony of C. picta from Sweetings Cay photographed on 30 May 2008. (B) C. picta (asterisk), C. pawliki (triangle), and Ecteinascidia turbinata (arrow) growing together in Sweetings Cay, photo taken on 10 June 2008. (C,D) C. rochae from Plana Cay, photographed on 25 June 2007. (E) Colony of C. pawliki photographed at Sweetings Cay on 17 June 2007. (F) C. pawliki and E. turbinata growing together in Sweetings Cay, photographed on 10 June 2008. (G) A colony of C. erwinorum from San Salvador photographed on 5 June 2008. (H) C. erwinorum photographed in Key Largo, Florida (USA) on 14 November 2006. Scale bar 1 cm.
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Figure 4. Clavelina picta. (A) Image of two zooid clumps; (B) Zooid without gonads. Note the straight abdomen. (C) Abdomen with gonads. (D–F) Left, ventral, and dorsal views of the thoracic musculature of a zooid. The dark body at the base of the branchial sac is a copepod. (G) Ventral view of the anterior part of a straight abdomen of the same zooid than in D–F, showing muscles passing from right to left. (H) Dissected thorax (stained). (I) Larva. Scale bars: (A), 15 mm; (B,C), 2 mm; (D–G), 2 mm; (H), 2 mm; (I), 0.4 mm.
Figure 4. Clavelina picta. (A) Image of two zooid clumps; (B) Zooid without gonads. Note the straight abdomen. (C) Abdomen with gonads. (D–F) Left, ventral, and dorsal views of the thoracic musculature of a zooid. The dark body at the base of the branchial sac is a copepod. (G) Ventral view of the anterior part of a straight abdomen of the same zooid than in D–F, showing muscles passing from right to left. (H) Dissected thorax (stained). (I) Larva. Scale bars: (A), 15 mm; (B,C), 2 mm; (D–G), 2 mm; (H), 2 mm; (I), 0.4 mm.
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Figure 5. Clavelina rochae. (A) Image of two zooids with their tunic. (B) Zooids extracted from the tunic. Note the twisted abdomen in the left zooid and the straight abdomen in the right zooid. (C) Branchial sac with brooded embryos. (D) Abdomen with gonads. (E) Dissected thorax (stained). (F) Larva. Scale bars: (A), 5 mm; (B), 2 mm; (C,D), 1 mm; (E): 1 mm; (F), 0.25 mm.
Figure 5. Clavelina rochae. (A) Image of two zooids with their tunic. (B) Zooids extracted from the tunic. Note the twisted abdomen in the left zooid and the straight abdomen in the right zooid. (C) Branchial sac with brooded embryos. (D) Abdomen with gonads. (E) Dissected thorax (stained). (F) Larva. Scale bars: (A), 5 mm; (B), 2 mm; (C,D), 1 mm; (E): 1 mm; (F), 0.25 mm.
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Figure 6. Clavelina pawliki. (A) Image of the basal part of two abdomens united to the basal layer of tunic. (B) Zooid. Note the straight abdomen and some brooded embryos. (C–E) Thoracic musculature from right (C), ventral (D), and dorsolateral (E) views. (E) Image stained to show the finer bands. (F) Dissected thorax (stained). (G) Larva (stained). Note that the tail separation from the body is an artifact of manipulation. Scale bars: (A), 2 mm; (B), 2 mm; (C–E), 2 mm; (F), 1 mm; (G), 0.3 mm.
Figure 6. Clavelina pawliki. (A) Image of the basal part of two abdomens united to the basal layer of tunic. (B) Zooid. Note the straight abdomen and some brooded embryos. (C–E) Thoracic musculature from right (C), ventral (D), and dorsolateral (E) views. (E) Image stained to show the finer bands. (F) Dissected thorax (stained). (G) Larva (stained). Note that the tail separation from the body is an artifact of manipulation. Scale bars: (A), 2 mm; (B), 2 mm; (C–E), 2 mm; (F), 1 mm; (G), 0.3 mm.
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Figure 7. Clavelina erwinorum. (A) Image of a clump of zooids basally embedded in a common tunic. (B) Zooid. Note the twisted abdomen. (C,D) Thoracic musculature from ventral (C) and laterodorsal (D) views. (E) Thorax cut open; the right side is shown (stained). (F) Larva. Scale bars: (A), 5 mm; (B), 1 mm; (C,D), 1 mm; (E): 1 mm; (F), 0.4 mm.
Figure 7. Clavelina erwinorum. (A) Image of a clump of zooids basally embedded in a common tunic. (B) Zooid. Note the twisted abdomen. (C,D) Thoracic musculature from ventral (C) and laterodorsal (D) views. (E) Thorax cut open; the right side is shown (stained). (F) Larva. Scale bars: (A), 5 mm; (B), 1 mm; (C,D), 1 mm; (E): 1 mm; (F), 0.4 mm.
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Figure 8. Phylogeny of partial Cytochrome Oxidase I gene sequences from clavelinidae species. The phylogenetic position of species from this study is highlighted (bold lettering). Labels on terminal nodes of sequences indicate the ascidian species and GenBank accession numbers or sampling code. Collection countries are noted in parentheses. The tree topology was obtained from maximum likelihood (ML) analysis. Individual bootstrap values from neighbor-joining (NJ) are under the tree nodes and, for ML analyses, above tree nodes when support values are greater than 50%. The scale bar represents the number of substitutions per site.
Figure 8. Phylogeny of partial Cytochrome Oxidase I gene sequences from clavelinidae species. The phylogenetic position of species from this study is highlighted (bold lettering). Labels on terminal nodes of sequences indicate the ascidian species and GenBank accession numbers or sampling code. Collection countries are noted in parentheses. The tree topology was obtained from maximum likelihood (ML) analysis. Individual bootstrap values from neighbor-joining (NJ) are under the tree nodes and, for ML analyses, above tree nodes when support values are greater than 50%. The scale bar represents the number of substitutions per site.
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Table 1. Clavelina species observed in the Bahamas. Sampling date, location, GPS position, site depth, and species name. Analyzed samples for each species are underlined.
Table 1. Clavelina species observed in the Bahamas. Sampling date, location, GPS position, site depth, and species name. Analyzed samples for each species are underlined.
DateLocationGPSSite DepthClavelina pictaClavelina pawlikiClavelina erwinorumClavelina rochae
15 June 2007Sweetings Cay, Grand Bahama26°34.182′ N; 77°53.342′ W 16.1 mXX
17 June 2007Sweetings Cay, Grand Bahama26°34.182′ N; 77°53.342′ W 13.3 mXX
19 June 2007Sweetings Cay, Grand Bahama26°34.326′ N; 77°53.733′ W 19.8 mXX
20 June 2007Stirrup Cay25°49.600′ N; 77°53.970′ W 11.9 mX
25 June 2007Plana Cay22°36.450′ N; 73°37.566′ W 31.7 m X
27 June 2007Sweetings Cay, Grand Bahama26°34.182′ N; 77°53.342′ W 23.4 m X
28 June 2007Sweetings Cay, Grand Bahama26°34.182′ N; 77°53.342′ W 18.8 m X
29 May 2008Sweetings Cay, Grand Bahama26°38.585′ N; 77°57.734′ W13.8 mX
30 May 2008Sweetings Cay, Grand Bahama26°38.585′ N; 77°57.734′ W20.4 mXX
5 June 2008San Salvador24°03.647′ N; 74°32.699′ W28.1 m XX
San Salvador24°04.161′ N; 74°32.684′ W29.4 m X
6 June 2008San Salvador24°04.703′ N; 74°32.797′ W29.8 m XX
7 June 2008West Plana Cay22°36.263′ N; 73°37.658′ W35.9 m X
West Plana Cay22°35.834′ N; 73°37.764′ W25.6 m X
8 June 2008Sweetings Cay, Grand Bahama22°36.450′ N; 73°33.728′ W27.6 m X
10 June 2008Sweetings Cay, Grand Bahama26°34.070′ N; 77°53.206′ W12.2 mXX
1 July 2010Sweetings Cay, Grand Bahama26°33.559′ N; 77°53.070′ W15.8 mXX
2 July 2010Sweetings Cay, Grand Bahama26°34.073′ N; 77°53.048′ W10.2 mXX
4 July 2010Sweetings Cay, Grand Bahama26°33.693′ N; 77°53.084′ W15.8 mXX
5 July 2010Sweetings Cay, Grand Bahama26°33.693′ N; 77°53.084′ W14.8 m X
8 July 2010San Salvador24°03.515′ N; 74°32.474′ W30.2 m XX
9 July 2010San Salvador24°04.228′ N; 74°32.683′ W33.2 m XX
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MDPI and ACS Style

Turon, X.; López-Legentil, S. New Clavelina (Ascidiacea) Species from the Bahamas. Taxonomy 2024, 4, 661-679. https://doi.org/10.3390/taxonomy4030034

AMA Style

Turon X, López-Legentil S. New Clavelina (Ascidiacea) Species from the Bahamas. Taxonomy. 2024; 4(3):661-679. https://doi.org/10.3390/taxonomy4030034

Chicago/Turabian Style

Turon, Xavier, and Susanna López-Legentil. 2024. "New Clavelina (Ascidiacea) Species from the Bahamas" Taxonomy 4, no. 3: 661-679. https://doi.org/10.3390/taxonomy4030034

APA Style

Turon, X., & López-Legentil, S. (2024). New Clavelina (Ascidiacea) Species from the Bahamas. Taxonomy, 4(3), 661-679. https://doi.org/10.3390/taxonomy4030034

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