Put a tiger in your tank: the polyclad flatworm Maritigrella crozieri as a proposed model for evo-devo
© Lapraz et al.; licensee BioMed Central Ltd. 2013
Received: 7 June 2013
Accepted: 14 August 2013
Published: 9 October 2013
Polyclad flatworms are an early branching clade within the rhabditophoran Platyhelminthes. They provide an interesting system with which to explore the evolution of development within Platyhelminthes and amongst Spiralia (Lophotrochozoa). Unlike most other flatworms, polyclads undergo spiral cleavage (similar to that seen in some other spiralian taxa), they are the only free-living flatworms where development via a larval stage occurs, and they are the only flatworms in which embryos can be reared outside of their protective egg case, enabling embryonic manipulations. Past work has focused on comparing early cleavage patterns and larval anatomy between polyclads and other spiralians. We have selected Maritigrella crozieri, the tiger flatworm, as a suitable polyclad species for developmental studies, because it is abundant and large in size compared to other species. These characteristics have facilitated the generation of a transcriptome from embryonic and larval material and are enabling us to develop methods for gene expression analysis and immunofluorescence techniques. Here we give an overview of M. crozieri and its development, we highlight the advantages and current limitations of this animal as a potential evo-devo model and discuss current lines of research.
KeywordsEvolutionary and developmental biology Larvae Neuropeptides Planarians Polyclad flatworms Regeneration Spiralians Stem cells Transcriptome Turbellarians
Platyhelminthes, or flatworms, are a group of soft-bodied, usually hermaphroditic invertebrates especially renowned for their neoblast stem cell system and their pronounced ability to regenerate [1, 2]. They are members of the Spiralia, and, with recent evidence supporting the exclusion of the Acoelomorpha and Xenoturbellida (see, for example, ), they comprise two monophyletic groups: the Catenulida and the Rhabditophora [4, 5], with the latter including the Polycladida.
The Polycladida are among the earliest branching Platyhelminthes [6–9]. They sport a highly branched gut (the name-giving feature of the Polycladida), and they have retained certain developmental characteristics common to other spiralian taxa, such as endolecithal eggs and (at least partially) spiral cleavage [10–12]. Embryonic development features a prominent mesentoblast and epibolic gastrulation. Whereas all other free-living flatworms are direct developers, both direct and indirect development is known in polyclads. Indirect development typically features a spherical, usually eight-lobed, three-eyed Müller’s larva. Other larval forms include a four-lobed, two-eyed Goette’s larva and a dorsoventrally flattened, eight-lobed, twelve-eyed Kato’s larva.
Developmental types in polyclads
Intracapsular Müller’s larva 
Intracapsular/reduced Müller’s larva 
Reduced Müller’s larva 
Direct development 
Goette’s larva 
Direct development 
Direct development 
Direct development [B Egger, unpublished observations]
Direct development 
Stylochus (Imogine) aomori
Goette’s larva 
Stylochus (Imogine) mcgrathi
Goette’s larva 
Stylochus (Imogine) mediterraneus
Goette’s larva 
Stylochus (Imogine) uniporus
Goette’s larva 
Stylochus (Imogine) zebra
Stylochus (Stylochus) flevensis
Goette’s larva 
Stylochus (Stylochus) frontalis
Direct development 
Stylochus (Stylochus) neapolitanus
Direct development 
Stylochus (Stylochus) pilidium
Goette’s larva 
Müller’s larva 
Adult polyclads are extremely dorsoventrally flattened, with body lengths typically in the centimetre range. They are subdivided into two groups characterized by the presence (Cotylea) or absence (Acotylea) of a cup-shaped ventral sucker located along the midline of the body and posterior to the genital openings [21, 36]. Although larval and juvenile stages may be found swimming in the water column, adults are usually found on the substrate . Polyclads are capable of regeneration, but that does not include the regeneration of the head, with the exception of the acotylean Cestoplana, in which pieces cut just posterior of the brain are able to regenerate the brain .
Herein we highlight the polyclad Maritigrella crozieri as a candidate model for evo-devo. We review past work done on this species, give a preview of new experimental data and discuss the most relevant scientific topics and future directions.
Maritigrella crozieri, a polyclad species for evo-devo
Despite the Lophotrochozoa being one of the three major branches of bilaterians, very few representatives of the Lophotrochozoa are found amongst classical experimental models. Emerging new model molluscs, annelids and platyhelminths have been pushed forward recently to fill this taxonomic gap . Among these, the principal platyhelminth models are triclads, which, despite being valuable models for stem cell and regeneration biology, have a derived mode of development, making comparative studies difficult (; see also review in ).
We have chosen M. crozieri as a suitable polyclad representative for evolutionary and developmental studies. The major advantages of this species are its ease of collection, its readily observable spiral cleavage, its biphasic life cycle with an eight-lobed Müller’s larva and its large size with many eggs, which can be obtained and raised without eggshells.
Description of Maritigrella crozieri
Additional file 1: Sperm transfer of adult Maritigrella crozieri. (AVI 1 MB)
On the dorsal surface, it has an irregular pattern of black stripes on a white to orange background that inspired the name for the genus  (Figures 1A through 1C, 2B and 2D through 2G). The eyes are located in two broad wedges, merging dorsal to the brain (cerebral eyes) and at both the ventral and dorsal bases of the tentacles (tentacular eyes, Figure 1C and 1D). We have counted more than 90 cerebral eyes in an individual (Figure 1C), which is slightly higher than the number (about 70) reported by Newman et al. . The animal’s maximum reported length was up to 30 mm  and 31.3 ± 2.7 mm (n = 20) . We have since found several significantly larger individuals, with the greatest measured length being 56 mm (Figure 1B).
The male genital opening is anterior to the female opening, with the sucker being situated behind them (, Figure 1E). Whereas Hyman  stated that “there is no stylet” in later works the stylet (the sclerotised tip of the male copulatory organ) was found to be present and to be about 130 μm long . Insemination has been reported to occur hypodermically by stabbing . In our observations, animals were not stabbing each other, but rather gently depositing sperm on the dorsal surface of their partners with their stylet (Figure 2E through 2G; see also Additional file 1). Histological sections of the spermatophores on or in the epidermis are required for ascertaining dermal impregnation instead of hypodermic insemination .
Maritigrella crozieri collection and prey preference
The type locality of M. crozieri is Bermuda [41, 42], and animals have subsequently been collected off mainland Florida and the Florida Keys [12, 13, 43, 47]. All our sampling efforts were concentrated on the Florida Keys, and we found animals (from west to east) on Sugarloaf Key, Cudjoe Key, Summerland Key, Ramrod Key, Big Pine Key, No Name Key, Long Key and Upper Matecumbe Key (Figure 2A). While most animals could be found on the ascidian Ecteinascidia turbinata (Figure 2B through 2D), specimens of M. crozieri were occasionally encountered on mangrove roots without Ecteinascidia or on the shallow ocean floor.
M. crozieri shows a strong preference for the orange ascidian E. turbinata as a food source [41, 43]. We occasionally found animals with a purple gut instead of an orange gut, indicating other prey items. Crozier  identified Ascidia curvata and Phallusia nigra (formerly Ascidia atra) as other possible prey for M. crozieri and speculated that the food specificity of adult specimens of M. crozieri may be an acquired taste of the juvenile, depending on which ascidian species the juvenile had settled. Newman et al. , on the other hand, claim that M. crozieri feeds exclusively on E. turbinata.
Additional file 2: Adult Maritigrella crozieri gliding under the water’s surface and then dropping to the bottom of the tank. (AVI 10 MB)
Laboratory cultures and development
Specimens of M. crozieri were able to survive up to 143 days without feeding, shrinking significantly during this time of starvation (Figure 1B, inset) at room temperature in 3.5% artificial seawater (hw-Meersalz professional and hw-Marinemix professional; Wiegandt GmbH, Krefeld, Germany).
The egg-laying period in the laboratory lasts for more than 3 months (107 days), showing that animals under ongoing starvation are still able to produce and lay eggs. The great majority of eggs are laid during the first 2 months of captivity. Gravid M. crozieri lay eggs directly on the tunic of E. turbinata in the wild and on the side of their container, or even under the water surface, when in captivity. Each embryo is contained within a thick spherical capsule, which collectively form a compact monolayer (egg plate) of 50 to 1,000 capsules. Importantly for experimental manipulation of the embryos, puncturing the paired uteri of gravid adults with a dissecting needle releases viable fertilised eggs devoid of their capsules (see ). If raised in a gelatin-coated petri dish in filtered seawater treated with antibiotics, these naked eggs develop in the same way as their encapsulated counterparts .
Genome and transcriptome
The haploid genome size (1C) of M. crozieri is estimated to be about 2.5 Gb (2,511.2 ± 35.8 Mb; n = 3) by flow cytometry (J Spencer Johnston, personal communication), providing the first genome size information for a polyclad flatworm. Diploid genome sizes of 38 free-living flatworms were shown to have a considerable range between about 0.1 and 40 Gb, with an average of about 5 Gb , which corresponds very closely to M. crozieri’s diploid genome size.
General properties of the Maritigrella crozieri transcriptome assembly produced using the Trinity assembly tool for de novo reconstruction of transcriptome sequences from RNA-seq data
Number of contigs
Total size of contigs
Number of contigs >500 nucleotides
Number of contigs >1,000
Number of contigs >10,000
Number of contigs >100,000
Mean contig size
Median contig size
Regeneration and stem cells
The regenerative capacity of some flatworms has been known for two centuries  and is reliant on the neoblast stem cell system. Some flatworms can readily regenerate a head when cut into 100 pieces  or regenerate to a complete organism from only 1,500 cells . However, not all flatworms are able to regenerate all missing body parts or even to regenerate at all . The “polyclad rule for regeneration” was postulated as the ability to regenerate all parts of the body but the brain . This is true for all polyclads studied so far, with the notable exception of Cestoplana, which is able to regenerate its head from the posterior fragment when amputated just posterior to the brain . Although more than 10 acotylean polyclads have been observed to regenerate , only a single cotylean species, Thysanozoon brocchii, has been the subject of published regeneration studies [58, 59]. Herein we can report that M. crozieri (n = 3) is able to regenerate both laterally and posteriorly at room temperature (Figure 3), although regeneration in small animals that have not fed for weeks or months is slow and extensive regeneration studies are still outstanding.
Neoblasts are totipotent stem cells and the only proliferating cells in adult rhabditophoran flatworms [60, 61]. In the Rhabditophora, neoblasts are located exclusively in the mesenchymal space and at the base of the gastrodermis, but they are conspicuously absent in the epidermis . Migration of stem cells into the epidermis has been demonstrated for juvenile polyclads [62, 63]. Interestingly, in late embryos of Notocomplana humilis and Cycloporus japonicus, and even in the Müller’s larvae of the latter, mitoses were detected in the epidermis . In M. crozieri larvae, the first Müller’s larvae with successful bromodeoxyuridine (BrdU) labelling following the protocol given by Egger et al.  using 20-h BrdU incubation time, we could not detect proliferating cells (S-phase or mitotic) within the epidermis (see Figure 7E).
Regeneration in flatworms and stem cell research on them so far have been focused mainly on triclads and Macrostomum, but they are interesting topics to study in polyclads. In particular, the absence (as in triclads ) or presence (as in Macrostomum ) of proliferating cells within the regeneration blastema will help determine the ancestral mode of tissue repair in flatworms. Large polyclads such as M. crozieri are amenable to microsurgery, thereby allowing us to explore more precisely the limits of their regenerative capacity.
Hypotheses on phylogenetic relationships and character evolution
Among the metazoan phyla, many diverse groups have a biphasic life cycle in which they pass through a pelagic larval stage that gives rise to a benthic adult animal through metamorphosis. The presence of such larvae is patchily distributed, however, raising the question of their evolutionary origin. The biphasic life cycle could be an ancient characteristic of animals, homologous in those groups in which it is found and repeatedly lost in those groups that lack it; or it may have evolved repeatedly via convergent evolution as a similar adaptation to some consistent selective forces. This dilemma has not been resolved, and the origin of larvae remains the subject of intense debate (for review, see the opposing views of Raff  and Nielsen ).
The phylogenetic position and features of polyclad flatworms make them a valuable model with which to gain insight into the evolution of larval forms within the Spiralia. Although it has been argued that the existence of larvae in a single order of free-living platyhelminths points to its independent evolution in polyclads rather than repeated loss in most other flatworm clades , a basally branching position of the polyclads within platyhelminths allows it to be parsimoniously considered as a primitive character [8, 69]. Polyclad flatworms can exhibit a range of developmental modes even within the same genus (Table 1): direct development with benthic juvenile worms, intermediate development with ciliated larvae metamorphosing within their eggshells and indirect development with pelagic swimming larvae that metamorphose postembryonically. These larvae have been classified into three main types: Kato’s larval features resemble those of a modified juvenile, whereas ciliary bands and cephalic ganglia found in Goette’s and Müller’s larvae have been thought by some to resemble those of the nemertean pilidium larva . Others have suggested that the polyclad larva may be homologous to the trochophore larva .
The practical advantages and pioneering works on M. crozieri’s larva, together with the growing access to large-scale molecular, imaging and phylogenetic tools, will help to elucidate the evolution of larval forms and features. Evidence of homology of Müller’s larval characteristics with those of other spiralian larvae would hint at indirect development as the primitive platyhelminth and spiralian condition. On the other hand, the nonhomology of larval characteristics would suggest that similar larval forms can evolve repeatedly and independently.
Evolution of the gut
Development of a through gut in metazoans has been a critical innovation that provides efficient food-processing. Despite this importance, its evolution is still surprisingly enigmatic. Within Eumetazoa, blind guts without a separate anus are found in ctenophores and cnidarians, whereas most bilaterians typically develop a through gut with two distinct openings. Some notable exceptions exist within bilaterians, however, as some species develop only a blind gut. This is the case for Xenacoelomorpha, Ophiuroidea, some Brachiopoda, some Rotifera, some Gnathostomulida and almost all Platyhelminthes . Two scenarios can be envisioned to explain the presence of a blind gut in those animals: Either they retained a characteristic that was present in their stem group, or they secondarily lost an anus that was already present in their stem group.
The nested position of Platyhelminthes within spiralians (for example, ) suggests that a secondary loss of the anus in flatworms is the most parsimonious explanation. In support of this view, it has been suggested that a divergent developmental program could account for the absence of a through gut in Platyhelminthes. The most obvious example is found in polyclads whereby fourth-quartet macromeres (4A–4D), which give rise to endodermal structures in other spiralians, degenerate [10, 11, 71, 73]. Also, many of the genes found expressed along the anteroposterior axis of bilaterian guts are not expressed in the gut or were lost in triclads . Only one of the three ParaHox genes, xlox, is found in Schmidtea mediterranea and Schmidtea polychroa genomes, whereas gsx and cdx, as well as the T-box-containing gene brachyury, are missing . The extent of such a loss is less pronounced in the Macrostomida  and in the polyclad Discocelis tigrina, which are more basally branched within the Platyhelminthes, indicating that these genes were secondarily lost in triclads and cannot be related to the lack of an anus in almost all Platyhelminthes.
Whilst our attempts to identify in the transcriptome of M. crozieri homologs of brachyury and xlox have been unsuccessful so far, we found homologs of gsx and cdx (Additional file 5: Figure S1, and Additional file 6). Saló and colleagues  reported the absence of a gsx homolog but the presence of an xlox homolog in the polyclad Discocelis tigrina, but its sequence has never been published and therefore could not be compared with the sequences from our transcriptome. It will be important in future studies to determine in M. crozieri’s embryos and juveniles the presence and the expression of these as well as other gut-related genes to better understand the evolutionary origin of the Platyhelminthes’ blind gut.
Evolution of the nervous system and phototaxis
Many aspects of nervous system evolution remain elusive. One of them is the transition from the diffuse nervous system found in cnidarians to the centralised one found in most bilaterians [75, 76]. Although some studies suggest that centralisation and complex patterning of the nervous system in adult animals predates the protostome–deuterostome split [77, 78], others favour the idea that the urbilaterian possessed a far less complex nervous system [76, 79].
The relatively simple nervous system of invertebrate ciliated larvae has been proposed to be informative regarding the evolution of the central nervous system , possibly recapitulating a transitional form en route to a complex adult nervous system. Conserved gene expression and immunoreactivity in the neurogenic region of these larvae [81–84] may hint at a common evolutionary history. However, the disappearance of the larval apical organ and ciliary band nerves during metamorphosis [80, 85], and also the small number of markers or phyla investigated, make the evolutionary significance of those comparisons difficult to determine .
Whilst precise descriptions of the nervous system of Macrostomum lignano[2, 87] and Schmidtea mediterranea have already provided landmarks for juvenile and adult platyhelminth neuroanatomy, very little is known concerning platyhelminth larvae. In this respect, future studies on the nervous system of larval M. crozieri can provide an additional data set for determining the evolutionary significance of the nervous system of ciliated larvae.
Recent work on Platynereis dumerilii suggested that the phototactic behaviour of their larvae represent a paradigm for the evolution of the nervous system . Phototaxis in P. dumerilii larvae relies on the presence of pigmented photoreceptors that connect directly to a ciliated locomotor cell of the prototrochal ring. In sponges and cnidarians, phototaxis relies on single cells, which have both light-detecting and ciliary locomotory functions, whereas several specialised cooperating cells are found in P. dumerilii. The latter could represent an early step in an evolutionary complexification of neural circuitry and visual systems .
M. crozieri larvae provide direct access to further testing of this hypothesis. In an assay similar to the one previously used for P. dumerilii, the polyclad larvae behaved positively phototactic (Figure 6M, Additional file 3 and Additional file 4; see also ), and they possess, like larvae of Platynereis dumerilii, rhabdomeric pigmented eyespots that develop closely associated with the bilateral cephalic ganglia , allowing close comparison between larvae of these two species.
The nervous system of M. crozieri larvae has been investigated using standard neuronal markers . Expansion of this work with additional specific markers, such as neuropeptides, is warranted. Neuropeptides have an early evolutionary origin  and have been shown to be implicated in the control of swimming and settlement behaviour [90, 91]. The rich repertoire of neuropeptides found in Platynereis dumerilii, together with a microscopic registration technique [92, 93], have proven to be powerful tools to characterise and map individual neurons in a whole larva.
M. crozieri is especially amenable to similar approaches. The recently established transcriptome of M. crozieri allowed us to identify a number of conserved neuropeptide motifs, such as an AVRLIRLamide and a GVWSNDPWamide. Antibodies directed against the mature form of the neuropeptide show that distinct subsets of cells in M. crozieri are immunoreactive to these antibodies (Figure 7F and 7G; for a suitable staining protocol, see ). Determination of the extent of homology and specificity in development and nervous topology between M. crozieri and other spiralians will have repercussions on our understanding of neuronal evolution. The increased availability of neuropeptidomes in other larvae, such as the sea urchin [94, 95], and two recent global analyses of neuropeptide evolution in Metazoa [96, 97] should provide a solid comparison framework for all bilaterians.
M. crozieri’s large size at maturity facilitates the extraction of hundreds of naked embryos from the adult and also makes the collection of individuals from the field easier. Adult worms can be kept in the laboratory without food for a considerable time and still produce eggs. The larvae can be kept alive in the laboratory for weeks, although raising them to metamorphosis has not been achieved to date.
An embryonic and larval transcriptome has been sequenced and assembled and is currently being analysed and complemented by full-genome sequencing. These resources are facilitating obtaining genes of interest for in situ probe synthesis, among others. We are in the process of developing a protocol for whole-mount in situ hybridization for M. crozieri. Whole-mount immunofluorescent staining works well  (Figure 7E through 7G), and we have produced polyclonal antibodies against some neuropeptides identified in the transcriptome (Figure 7F and 7G).
We thank Kevin Olsen, Andrew Gillis, Fraser Simpson and Lena Egger for help with collecting and maintaining specimens, and we thank Lena Egger also for providing Figure 1A. J Spencer Johnston kindly provided the genome size data. Dave Vaughan, Nick Disalvo, Robert Etti and Mark Knowles from Mote Marine Laboratory are gratefully acknowledged for general support at the station. We thank the two anonymous reviewers for careful review of the manuscript and valuable comments. This work was supported by a Leverhulme Trust grant (F/07 134/DA) and a Biotechnology and Biological Sciences Research Council grant (BB/H006966/1) (to MJT), as well as by a Sparkling Science grant from the Austrian Ministry of Science and Research (SPA/02-81) (to BE). MJT is supported by a Royal Society Wolfson Research Merit Award.
- Rieger RM, Tyler S, Smith JPS, Rieger GE: Platyhelminthes: Turbellaria. Microscopic Anatomy of Invertebrates: Volume 3. Platyhelminthes Nemertinea. Edited by: Harrison FW, Bogitsh BJ. 1991, New York: Wiley-Liss, 7-140.Google Scholar
- Egger B, Gschwentner R, Rieger R: Free-living flatworms under the knife: past and present. Dev Genes Evol. 2007, 217: 89-104. 10.1007/s00427-006-0120-5.PubMed CentralView ArticlePubMedGoogle Scholar
- Philippe H, Brinkmann H, Copley RR, Moroz LL, Nakano H, Poustka AJ, Wallberg A, Peterson KJ, Telford MJ: Acoelomorph flatworms are deuterostomes related to Xenoturbella. Nature. 2011, 470: 255-258. 10.1038/nature09676.PubMed CentralView ArticlePubMedGoogle Scholar
- Larsson K, Jondelius U: Phylogeny of Catenulida and support for Platyhelminthes. Org Divers Evol. 2008, 8: 378-387. 10.1016/j.ode.2008.09.002.View ArticleGoogle Scholar
- Giribet G: Assembling the lophotrochozoan (=spiralian) tree of life. Philos Trans R Soc Lond B Biol Sci. 2008, 363: 1513-1522. 10.1098/rstb.2007.2241.PubMed CentralView ArticlePubMedGoogle Scholar
- Ehlers U: Das phylogenetische System der Plathelminthes. 1985, Stuttgart, Germany: Gustav Fischer VerlagGoogle Scholar
- Littlewood DTJ: Platyhelminth systematics and the emergence of new characters. Parasite. 2008, 15: 333-341. 10.1051/parasite/2008153333.View ArticlePubMedGoogle Scholar
- Martín-Durán JM, Egger B: Developmental diversity in free-living flatworms. EvoDevo. 2012, 3: 7-10.1186/2041-9139-3-7.PubMed CentralView ArticlePubMedGoogle Scholar
- Lockyer AE, Olson PD, Littlewood DTJ: Utility of complete large and small subunit rRNA genes in resolving the phylogeny of the Neodermata (Platyhelminthes): implications and a review of the cercomer theory. Biol J Linn Soc Lond. 2003, 78: 155-171. 10.1046/j.1095-8312.2003.00141.x.View ArticleGoogle Scholar
- Surface FM: The early development of a polyclad, Planocera Inquilina Wh. Proc Acad Nat Sci Phila. 1907, 59: 514-559.Google Scholar
- Boyer BC, Henry JQ, Martindale MQ: Dual origins of mesoderm in a basal spiralian: cell lineage analyses in the polyclad turbellarian Hoploplana inquilina. Dev Biol. 1996, 179: 329-338. 10.1006/dbio.1996.0264.View ArticlePubMedGoogle Scholar
- Rawlinson KA: Embryonic and post-embryonic development of the polyclad flatworm Maritigrella crozieri: implications for the evolution of spiralian life history traits. Front Zool. 2010, 7: 12-10.1186/1742-9994-7-12.PubMed CentralView ArticlePubMedGoogle Scholar
- Johnson KB, Forward RB: Larval photoresponses of the polyclad flatworm Maritigrella crozieri (Platyhelminthes, Polycladida) (Hyman). J Exp Mar Biol Ecol. 2003, 282: 103-112. 10.1016/S0022-0981(02)00448-3.View ArticleGoogle Scholar
- Scarpa J, Weis B, Ruppert E, Frick J, Ford A, Wright A: Direct evidence for planktotrophy in Müller’s larva of the tiger flatworm, Pseudoceros crozieri [abstract]. Am Zool. 1996, 36: 107-Google Scholar
- Rawlinson KA, Stella JS: Discovery of the corallivorous polyclad flatworm, Amakusaplana acroporae, on the Great Barrier Reef, Australia: the first report from the wild. PLoS One. 2012, 7: e42240-10.1371/journal.pone.0042240.PubMed CentralView ArticlePubMedGoogle Scholar
- Bolaños DM: Comparative embryology and muscle development of polyclad flatworms (Platyhelminthes-Rhabditophorans). PhD thesis. 2008, University of New Hampshire, Department of ZoologyGoogle Scholar
- Selenka E: Zoologische Studien: 2 Zur Entwickelungsgeschichte der Seeplanarien. 1881, Leipzig, Germany: W. EngelmannGoogle Scholar
- Anderson D: The embryonic and larval development of the turbellarian Notoplana australis (Schmarda, 1859) (Polycladida : Leptoplanidae). Mar Freshw Res. 1977, 28: 303-310. 10.1071/MF9770303.View ArticleGoogle Scholar
- Kato K: On the development of some Japanese polyclads. Jpn J Zool. 1940, 8: 537-573.Google Scholar
- Wheeler W: Planocera inquilina, a polyclad inhabiting the branchial chamber of Syncotypus canaliculatus. Gill J Morphol. 1894, 9: 195-201. 10.1002/jmor.1050090203.View ArticleGoogle Scholar
- Lang A: Die Polycladen (Seeplanarien) des Golfes von Neapel und der angrenzenden Meeresabschnitte: eine Monographie. 1884, Leipzig, Germany: Verlag Wilhelm EngelmannGoogle Scholar
- Claparède É: Glanures zootomiques parmi les Annélides de Port Vendres (Pyrénées Orientales). Mémoires de la Société de Physique et d’Histoire Naturelle de Genève. 1864, 17: 463-600.Google Scholar
- Remane A: Die Polycladen der Kieler Förde. Schriften Naturwiss Ver Schleswig-Holstein. 1929, 19: 73-79.Google Scholar
- Younossi-Hartenstein A, Hartenstein V: The embryonic development of the polyclad flatworm Imogine mcgrathi. Dev Genes Evol. 2000, 210: 383-398. 10.1007/s004270000086.View ArticlePubMedGoogle Scholar
- Galleni L: Polyclads of the Tuscan coasts. II. Stylochus alexandrinus Steinböck and Stylochus mediterraneus n. sp. from the rocky shores near Pisa and Livorno. Boll Zool. 1976, 43: 15-25. 10.1080/11250007609434882.View ArticleGoogle Scholar
- Lytwyn MW, McDermott JJ: Incidence, reproduction and feeding of Stylochus zebra, a polyclad turbellarian symbiont of hermit crabs. Mar Biol. 1976, 38: 365-372. 10.1007/BF00391376.View ArticleGoogle Scholar
- Rawlinson KA, Bolaños DM, Liana MK, Litvaitis MK: Reproduction, development and parental care in two direct-developing flatworms (Platyhelminthes : Polycladida : Acotylea). J Nat Hist. 2008, 42: 2173-2192. 10.1080/00222930802262758.View ArticleGoogle Scholar
- Hofker DJ: Faunistische Beobachtungen in der Zuidersee während der Trockenlegung. Z Für Morphol Ökologie Tiere. 1930, 18: 189-216. 10.1007/BF00419209.View ArticleGoogle Scholar
- Pearse AS, Wharton GW: The oyster “leech”, Stylochus inimicus Palombi, associated with oysters on the coasts of Florida. Ecol Monogr. 1938, 8: 605-656. 10.2307/1943085.View ArticleGoogle Scholar
- Girard C: Researches upon Nemerteans and Planarians. I, Embryonic Development of Planocera Elliptica. 1854, Philadelphia: Merrihew and ThompsonGoogle Scholar
- Provenzano AJ: Effects of the flatworm Stylochus ellipticus (Girard) on oyster spat in two saltwater ponds in Massachusetts. Proc Natl Shellfish Assoc. 1959, 50: 83-88.Google Scholar
- Müller J: Über verschiedene Formen von Seethieren. Arch Anat Physiol Wiss Med. 1854, 69-98.Google Scholar
- Gammoudi M, Noreña C, Tekaya S, Prantl V, Egger B: Insemination and embryonic development of some Mediterranean polyclad flatworms. Invertebr Reprod Dev. 2012, 56: 272-286. 10.1080/07924259.2011.611825.View ArticleGoogle Scholar
- Teshirogi W, Ishida S, Jatani K: On the early development of some Japanese polyclads. Rep Fukuara Mar Biol Lab. 1981, 2-31.Google Scholar
- Tang QY, Wang YJ, Wang XA: Early embryo and larva of Planocera reticulata: in vitro fertilization and SEM observation. Chin J Zool. 2011, 46: 66-71.Google Scholar
- Faubel A: The Polycladida, Turbellaria: proposal and establishment of a new system. Part I. The Acotylea. Mitteilungen Aus Dem Hambg Zool Mus Inst. 1983, 80: 17-121.Google Scholar
- Prudhoe S: A Monograph on Polyclad Turbellaria. 1985, London: British Museum (Natural History)Google Scholar
- Child CM: Studies on regulation. X. The positions and proportions of parts during regulation in Cestoplana in the absence of the cephalic ganglia. Arch Für Entwicklungsmechanik Org. 1905, 20: 157-186. 10.1007/BF02162809.View ArticleGoogle Scholar
- Tessmar-Raible K, Arendt D: Emerging systems: between vertebrates and arthropods, the Lophotrochozoa. Curr Opin Genet Dev. 2003, 13: 331-340. 10.1016/S0959-437X(03)00086-8.View ArticlePubMedGoogle Scholar
- Newmark PA, Sánchez Alvarado A: Not your father’s planarian: a classic model enters the era of functional genomics. Nat Rev Genet. 2002, 3: 210-219.View ArticlePubMedGoogle Scholar
- Crozier WJ: On the pigmentation of a polyclad. Proc Am Acad Arts Sci. 1917, 52: 725-730. 10.2307/20025707.View ArticleGoogle Scholar
- Hyman LH: Acoel and polyclad Turbellaria from Bermuda and the Sargassum. Bull Bingham Oceanogr Collect. 1939, 7:Art. 1: 1-26. 9 plates [Peabody Museum of Natural History, Yale University]Google Scholar
- Newman LJ, Norenburg JL, Reed S: Taxonomic and biological observations on the tiger flatworm, Maritigrella crozieri (Hyman, 1939), new combination (Platyhelminthes, Polycladida, Euryleptidae) from Florida waters. J Nat Hist. 2000, 34: 799-808. 10.1080/002229300299264.View ArticleGoogle Scholar
- Newman LJ, Cannon LRG: A new genus of euryleptid flatworm (Platyhelminthes, Polycladida) from the Indo-Pacific. J Nat Hist. 2000, 34: 191-205. 10.1080/002229300299606.View ArticleGoogle Scholar
- Bolaños DM, Quiroga SY, Litvaitis MK: Five new species of cotylean flatworms (Platyhelminthes: Polycladida) from the wider Caribbean. Zootaxa. 2007, 1650: 1-23.Google Scholar
- Litvaitis MK, Bolaños DM, Quiroga SY: When names are wrong and colours deceive: unravelling the Pseudoceros bicolor species complex (Turbellaria: Polycladida). J Nat Hist. 2010, 44: 829-845. 10.1080/00222930903537074.View ArticleGoogle Scholar
- Bolaños DM, Litvaitis MK: Embryonic muscle development in direct and indirect developing marine flatworms (Platyhelminthes, Polycladida). Evol Dev. 2009, 11: 290-301. 10.1111/j.1525-142X.2009.00331.x.View ArticlePubMedGoogle Scholar
- Boyer BC: Development of in vitro fertilized embryos of the polyclad flatworm, Hoploplana inquilina, following blastomere separation and deletion. Rouxs Arch Dev Biol. 1987, 196: 158-164. 10.1007/BF00376309.View ArticleGoogle Scholar
- Jékely G, Colombelli J, Hausen H, Guy K, Stelzer E, Nédélec F, Arendt D: Mechanism of phototaxis in marine zooplankton. Nature. 2008, 456: 395-399. 10.1038/nature07590.View ArticlePubMedGoogle Scholar
- Meijering E, Dzyubachyk O, Smal I: Methods for cell and particle tracking. Methods Enzymol. 2012, 504: 183-200.View ArticlePubMedGoogle Scholar
- Gregory TR, Hebert PD, Kolasa J: Evolutionary implications of the relationship between genome size and body size in flatworms and copepods. Heredity (Edinb). 2000, 84: 201-208. 10.1046/j.1365-2540.2000.00661.x.View ArticleGoogle Scholar
- Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, Adiconis X, Fan L, Raychowdhury R, Zeng Q, Chen Z, Mauceli E, Hacohen N, Gnirke A, Rhind N, di Palma F, Birren BW, Nusbaum C, Lindblad-Toh K, Friedman N, Regev A: Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat Biotechnol. 2011, 29: 644-652. 10.1038/nbt.1883.PubMed CentralView ArticlePubMedGoogle Scholar
- Bortoluzzi S, d’Alessi F, Romualdi C, Danieli GA: The human adult skeletal muscle transcriptional profile reconstructed by a novel computational approach. Genome Res. 2000, 10: 344-349. 10.1101/gr.10.3.344.PubMed CentralView ArticlePubMedGoogle Scholar
- Dalyell JG: Observations on Some Interesting Phenomena in Animal Physiology. 1814, Edinburgh, Archibald Constable: Exhibited by Several Species of Planariae. Illustrated by Coloured Figures of Living AnimalsGoogle Scholar
- Morgan TH: Experimental studies of the regeneration of Planaria maculata. Arch Für Entwicklungsmechanik Org. 1898, 7: 364-397. 10.1007/BF02161491.View ArticleGoogle Scholar
- Egger B, Ladurner P, Nimeth K, Gschwentner R, Rieger R: The regeneration capacity of the flatworm Macrostomum lignano––on repeated regeneration, rejuvenation, and the minimal size needed for regeneration. Dev Genes Evol. 2006, 216: 565-577. 10.1007/s00427-006-0069-4.PubMed CentralView ArticlePubMedGoogle Scholar
- Olmsted JMD: The role of the nervous system in the regeneration of polyclad turbellaria. J Exp Zool. 1922, 36: 48-56. 10.1002/jez.1400360103.View ArticleGoogle Scholar
- Loeb J: Einleitung in die vergleichende Gehirnphysiologie und vergleichende Psychologie, mit besonderer Berücksichtigung der wirbellosen Thiere. 1899, Leipzig, Germany: J. A. BarthView ArticleGoogle Scholar
- Monti R: La Rigenerazione Nelle Planarie Marine. 1900, Milan: Milan R. Istituto LombardoGoogle Scholar
- Baguñà J, Romero R, Saló E, Collet J, Auladell C, Ribas M, Riutort M, García-Fernàndez J, Burgaya F, Bueno D: Growth, degrowth and regeneration as developmental phenomena in adult freshwater planarians. Exp Embryol Aquat Plants Anim. 1990, 195: 129-162. [NATO ASI Series A: Life Sciences. Edited by Marthy HJ.]View ArticleGoogle Scholar
- Wagner DE, Wang IE, Reddien PW: Clonogenic neoblasts are pluripotent adult stem cells that underlie planarian regeneration. Science. 2011, 332: 811-816. 10.1126/science.1203983.PubMed CentralView ArticlePubMedGoogle Scholar
- Egger B, Steinke D, Tarui H, De Mulder K, Arendt D, Borgonie G, Funayama N, Gschwentner R, Hartenstein V, Hobmayer B, Hooge M, Hrouda M, Ishida S, Kobayashi C, Kuales G, Nishimura O, Pfister D, Rieger R, Salvenmoser W, Smith J, Technau U, Tyler S, Agata K, Salzburger W, Ladurner P: To be or not to be a flatworm: the acoel controversy. PLoS One. 2009, 4: e5502-10.1371/journal.pone.0005502.PubMed CentralView ArticlePubMedGoogle Scholar
- Drobysheva IM, Mamkaev YV: On mitosis in embryos and larvae of polyclads (Platyhelminthes). Belg J Zool. 2001, 131: 65-66.Google Scholar
- Ladurner P, Egger B, Mulder K, Pfister D, Kuales G, Salvenmoser W, Schärer L: The stem cell system of the basal flatworm Macrostomum lignano. Stem Cells: From Hydra to Man. Edited by: Bosch TCG. 2008, Dordrecht: Springer Netherlands, 75-94.View ArticleGoogle Scholar
- Saló E, Baguñà J: Regeneration and pattern formation in planarians I. The pattern of mitosis in anterior and posterior regeneration in Dugesia (G) tigrina, and a new proposal for blastema formation. J Embryol Exp Morphol. 1984, 83: 63-80.PubMedGoogle Scholar
- Egger B, Gschwentner R, Hess MW, Nimeth KT, Adamski Z, Willems M, Rieger R, Salvenmoser W: The caudal regeneration blastema is an accumulation of rapidly proliferating stem cells in the flatworm Macrostomum lignano. BMC Dev Biol. 2009, 9: 41-10.1186/1471-213X-9-41.PubMed CentralView ArticlePubMedGoogle Scholar
- Raff RA: Origins of the other metazoan body plans: the evolution of larval forms. Philos Trans R Soc Lond B Biol Sci. 2008, 363: 1473-1479. 10.1098/rstb.2007.2237.PubMed CentralView ArticlePubMedGoogle Scholar
- Nielsen C: How did indirect development with planktotrophic larvae evolve?. Biol Bull. 2009, 216: 203-215.PubMedGoogle Scholar
- Littlewood DTJ, Rohde K, Clough KA: The interrelationships of all major groups of Platyhelminthes: phylogenetic evidence from morphology and molecules. Biol J Linn Soc Lond. 1999, 66: 75-114. 10.1111/j.1095-8312.1999.tb01918.x.View ArticleGoogle Scholar
- Jägersten G: Evolution of the Metazoan Life Cycle: A Comprehensive Theory. 1972, New York: Academic PressGoogle Scholar
- Nielsen C: Trochophora larvae: cell-lineages, ciliary bands and body regions. 2. Other groups and general discussion. J Exp Zool B Mol Dev Evol. 2005, 304B: 401-447. 10.1002/jez.b.21050.View ArticleGoogle Scholar
- Westheide W, Rieger R: Spezielle Zoologie. Teil 1: Einzeller und Wirbellose Tiere. 2. Auflage [Gebundene Ausgabe]. 2006, Spektrum Akademischer Verlag: Heidelberg, GermanyGoogle Scholar
- Martín-Durán JM, Romero R: Evolutionary implications of morphogenesis and molecular patterning of the blind gut in the planarian Schmidtea polychroa. Dev Biol. 2011, 352: 164-176. 10.1016/j.ydbio.2011.01.032.View ArticlePubMedGoogle Scholar
- Saló E, Tauler J, Jimenez E, Bayascas JR, Gonzalez-Linares J, Garcia-Fernàndez J, Baguñà J: Hox and ParaHox genes in flatworms: characterization and expression. Am Zool. 2001, 41: 652-663. 10.1668/0003-1569(2001)041[0652:HAPGIF]2.0.CO;2.Google Scholar
- Arendt D, Denes AS, Jékely G, Tessmar-Raible K: The evolution of nervous system centralization. Philos Trans R Soc Lond B Biol Sci. 2008, 363: 1523-1528. 10.1098/rstb.2007.2242.PubMed CentralView ArticlePubMedGoogle Scholar
- Northcutt RG: Evolution of centralized nervous systems: two schools of evolutionary thought. Proc Natl Acad Sci USA. 2012, 109 (Suppl 1): 10626-10633.PubMed CentralView ArticlePubMedGoogle Scholar
- Denes AS, Jékely G, Steinmetz PRH, Raible F, Snyman H, Prud’homme B, Ferrier DEK, Balavoine G, Arendt D: Molecular architecture of annelid nerve cord supports common origin of nervous system centralization in bilateria. Cell. 2007, 129: 277-288. 10.1016/j.cell.2007.02.040.View ArticlePubMedGoogle Scholar
- Nomaksteinsky M, Röttinger E, Dufour HD, Chettouh Z, Lowe CJ, Martindale MQ, Brunet JF: Centralization of the deuterostome nervous system predates chordates. Curr Biol. 2009, 19: 1264-1269. 10.1016/j.cub.2009.05.063.View ArticlePubMedGoogle Scholar
- Hejnol A, Martindale MQ: Acoel development supports a simple planula-like urbilaterian. Philos Trans R Soc Lond B Biol Sci. 2008, 363: 1493-1501. 10.1098/rstb.2007.2239.PubMed CentralView ArticlePubMedGoogle Scholar
- Nielsen C: Larval and adult brains. Evol Dev. 2005, 7: 483-489. 10.1111/j.1525-142X.2005.05051.x.View ArticlePubMedGoogle Scholar
- Santagata S, Resh C, Hejnol A, Martindale MQ, Passamaneck YJ: Development of the larval anterior neurogenic domains of Terebratalia transversa (Brachiopoda) provides insights into the diversification of larval apical organs and the spiralian nervous system. EvoDevo. 2012, 3: 3-10.1186/2041-9139-3-3.PubMed CentralView ArticlePubMedGoogle Scholar
- Sinigaglia C, Busengdal H, Leclère L, Technau U, Rentzsch F: The bilaterian head patterning gene six3/6 controls aboral domain development in a cnidarian. PLoS Biol. 2013, 11: e1001488-10.1371/journal.pbio.1001488.PubMed CentralView ArticlePubMedGoogle Scholar
- Hay-Schmidt A: The evolution of the serotonergic nervous system. Proc Biol Sci. 2000, 267: 1071-1079. 10.1098/rspb.2000.1111.PubMed CentralView ArticlePubMedGoogle Scholar
- Dunn EF, Moy VN, Angerer LM, Angerer RC, Morris RL, Peterson KJ: Molecular paleoecology: using gene regulatory analysis to address the origins of complex life cycles in the late Precambrian. Evol Dev. 2007, 9: 10-24. 10.1111/j.1525-142X.2006.00134.x.View ArticlePubMedGoogle Scholar
- Nielsen C: Animal Evolution: Interrelationships of the Living Phyla. 2001, Oxford: Oxford University Press, 2Google Scholar
- Holland ND: Early central nervous system evolution: an era of skin brains?. Nat Rev Neurosci. 2003, 4: 617-627. 10.1038/nrn1175.View ArticlePubMedGoogle Scholar
- Morris J, Cardona A, Del Mar De Miguel-Bonet M, Hartenstein V: Neurobiology of the basal platyhelminth Macrostomum lignano: map and digital 3D model of the juvenile brain neuropile. Dev Genes Evol. 2007, 217: 569-584. 10.1007/s00427-007-0166-z.View ArticlePubMedGoogle Scholar
- Cebrià F: Organization of the nervous system in the model planarian Schmidtea mediterranea: an immunocytochemical study. Neurosci Res. 2008, 61: 375-384. 10.1016/j.neures.2008.04.005.View ArticlePubMedGoogle Scholar
- Grimmelikhuijzen CJP, Hauser F: Mini-review: the evolution of neuropeptide signaling. Regul Pept. 2012, 177 (Suppl): S6-S9.View ArticlePubMedGoogle Scholar
- Conzelmann M, Offenburger SL, Asadulina A, Keller T, Münch TA, Jékely G: Neuropeptides regulate swimming depth of Platynereis larvae. Proc Natl Acad Sci USA. 2011, 108: E1174-E1183. 10.1073/pnas.1109085108.PubMed CentralView ArticlePubMedGoogle Scholar
- Conzelmann M, Williams EA, Tunaru S, Randel N, Shahidi R, Asadulina A, Berger J, Offermanns S, Jékely G: Conserved MIP receptor–ligand pair regulates Platynereis larval settlement. Proc Natl Acad Sci USA. 2013, 110: 8224-8229. 10.1073/pnas.1220285110.PubMed CentralView ArticlePubMedGoogle Scholar
- Tomer R, Denes AS, Tessmar-Raible K, Arendt D: Profiling by image registration reveals common origin of annelid mushroom bodies and vertebrate pallium. Cell. 2010, 142: 800-809. 10.1016/j.cell.2010.07.043.View ArticlePubMedGoogle Scholar
- Asadulina A, Panzera A, Verasztó C, Liebig C, Jékely G: Whole-body gene expression pattern registration in Platynereis larvae. EvoDevo. 2012, 3: 27-10.1186/2041-9139-3-27.PubMed CentralView ArticlePubMedGoogle Scholar
- Menschaert G, Vandekerckhove TTM, Baggerman G, Landuyt B, Sweedler JV, Schoofs L, Luyten W, Van Criekinge W: A hybrid, de novo based, genome-wide database search approach applied to the sea urchin neuropeptidome. J Proteome Res. 2010, 9: 990-996. 10.1021/pr900885k.PubMed CentralView ArticlePubMedGoogle Scholar
- Rowe ML, Elphick MR: The neuropeptide transcriptome of a model echinoderm, the sea urchin Strongylocentrotus purpuratus. Gen Comp Endocrinol. 2012, 179: 331-344. 10.1016/j.ygcen.2012.09.009.View ArticlePubMedGoogle Scholar
- Mirabeau O, Joly JS: Molecular evolution of peptidergic signaling systems in bilaterians. Proc Natl Acad Sci USA. 2013, 110: E2028-E2037. 10.1073/pnas.1219956110.PubMed CentralView ArticlePubMedGoogle Scholar
- Jékely G: Global view of the evolution and diversity of metazoan neuropeptide signaling. Proc Natl Acad Sci USA. 2013, 110: 8702-8707. 10.1073/pnas.1221833110.PubMed CentralView ArticlePubMedGoogle Scholar
- Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, Li W, Lopez R, McWilliam H, Remmert M, Söding J, Thompson JD, Higgins DG: Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol Syst Biol. 2011, 7: 539-PubMed CentralView ArticlePubMedGoogle Scholar
- Waterhouse AM, Procter JB, Martin DM, Clamp M, Barton GJ: Jalview version 2: a multiple sequence alignment editor and analysis workbench. Bioinformatics. 2009, 25: 1189-1191. 10.1093/bioinformatics/btp033.PubMed CentralView ArticlePubMedGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.