Establishing a complete list of available names for a taxon often requires an enormous amount of time but it is the keystone of any taxonomic revision, because otherwise it would be impossible to address the nomenclatural status of available names and to determine how many new species names are needed.

All available type specimens were re-examined beyond the taxon of interest (Peronia) because species names often are incorrectly classified when they are first created. For instance, Onchidium durum Labbé, 1934a was originally created for slugs with a smooth notum, but the types of O. durum clearly bear dorsal gills. Ignoring O. durum because it was created for slugs with a smooth notum would have led to an incomplete list of available Peronia species names. Several species names had to be transferred to Peronia, because they refer to slugs with dorsal gills, regardless of whether species were originally described with dorsal gills or not. When type specimens are not located, one needs to go through original species descriptions very carefully, and still beyond the taxon of interest. Reciprocally, not all species names ever classified in Peronia belong to Peronia: for instance, several specific names originally combined with Peronia refer to Onchidella species. Finally, many species names of doubtful application need to be commented upon.

In total, 51 species-group names had to be considered for the revision of Peronia. Of these, only 31 are available Peronia species names (Table 1). Indeed, ten of those 51 names are not classified in Peronia: eight names refer to Onchidella species, one to a Wallaconchis species, and one to a Marmaronchis species. And, ten other names are nomina dubia as they refer to species which may belong to any onchidiid genus or which may not even belong to an onchidiid genus.

Many subsequent incorrect spellings are encountered in the onchidiid literature. The subsequent incorrect spelling of a name is not available and, to our knowledge, no subsequent incorrect spelling is in prevailing usage (ICZN 1999: Article 33.3). Subsequent incorrect spellings of specific names are corrected throughout the present monograph. When a spelling mistake is quite big, it is pointed out, such as, for instance, when JE Gray (1850: 117) erroneously used Peronia tongensis instead of Peronia tongana, and when Mörch (1872b: 325) erroneously used Peronia vermiculata instead of Peronia verruculata. In addition, many Peronia species were originally classified in Buchannan’s (1800) Onchidium, for which some authors (e.g., Plate 1893; Hoffmann 1928; Labbé, 1934a) used the unjustified emendation Oncidium, which is systematically corrected as Onchidium.

AM Australian Museum, Sydney, New South Wales, Australia

ANSP Academy of Natural Sciences, Drexel University, Philadelphia, Pennsylvania, USA

BNHS Bombay Natural History Society, Mumbai, India

BPBM Bernice Pauahi Bishop Museum, Honolulu, Hawaii, USA

CASIZ California Academy of Sciences, San Francisco, California, USA

ITBZC Institute of Tropical Biology, Zoology Collection, Vietnam Academy of Science and Technology, Ho Chi Minh City, Vietnam

MNHN Muséum national d’Histoire naturelle, Paris, France

MTQ Museum of Tropical Queensland, Townsville, Queensland, Australia

NHMD Zoological Museum, Natural History Museum of Denmark, University of Copenhagen, Denmark

NHMUK Natural History Museum, London, United Kingdom

NMSA KwaZulu-Natal Museum, Pietermaritzburg, KwaZulu-Natal, South Africa

NSMT National Museum of Nature and Science, Tokyo, Japan

NTM Museum and Art Gallery Northern Territory, Darwin, Northern Territory, Australia

PNM National Museum of the Philippines, Manila, Philippines

RBINS Royal Belgian Institute of Natural Sciences, Brussels, Belgium

SMF Senckenberg Forschungsinstitut und Naturmuseum, Frankfurt am Main, Germany

SMNH Swedish Museum of Natural History, Stockholm, Sweden

UF University of Florida, Gainesville, USA

UMIZ Universitas Malikussaleh, North Aceh, Sumatra, Indonesia

USMMC Universiti Sains Malaysia, Mollusk Collection, Penang, Malaysia

WAM Western Australian Museum, Perth, Western Australia, Australia

ZMB Museum für Naturkunde, Berlin, Germany

ZMH Zoologisches Museum, Hamburg, Germany

ZSM Zoologische Staatssammlung München, Munich, Germany

ZRC Zoological Reference Collection, Lee Kong Chian Natural History Museum, National University of Singapore

Our molecular analyses and anatomical species descriptions are based on a data set of 179 individuals specifically gathered for the present study. Of those 179 individuals, 112 were collected by the Dayrat lab, of which 91 were deposited in countries or states of origin (Australia, Hawaii, India, Indonesia, Japan, Malaysia, Philippines, Singapore, Vietnam) and 21 (from Madagascar and Mauritius) were deposited at the MNHN. Of those 179 individuals, 36 were collected during MNHN expeditions organized by Philippe Bouchet (Madagascar, Mozambique, New Caledonia, Papua New Guinea, and Vanuatu) and are all preserved at the MNHN; the specimens from New Caledonia were collected by Adam Bourke. Of those 179 individuals, 12 were collected by several collaborators: Sadar Aslam collected three specimens from Pakistan deposited at the MNHN; Clay Carlson collected two specimens from Guam deposited at the CAS and the MNHN; Owen Griffiths collected one specimen from Mauritius deposited at the MNHN; Shau Hwai (Aileen) Tan collected two specimens from Malaysia deposited at the USM; and Tomoyuki Nakano collected four specimens from Japan deposited at the NSMT. And, finally, 19 were found in museum collections: four (AM), four (NHMUK), two (NMSA), and nine (UF).

Collecting expeditions of the Dayrat lab were led by Benoît Dayrat in the Andaman Islands (India), West Bengal (India), Peninsular Malaysia, the Philippines, Singapore, New South Wales (Australia), and Northern territory (Australia), by Tricia Goulding in Queensland (Australia), Mauritius, Madagascar, Vietnam, and western India, by Munawar Khalil in Indonesia, and by Rebecca Cumming in Japan. Sites were accessed by car or by boat. Although each site was explored for an average of two hours, the exact time spent at each site also depended on the time of the low tide, the weather conditions, etc. Photographs were taken to document the kind of habitat being visited as well as the diverse microhabitats where specimens were collected. Specimens were individually numbered and photographed in their respective habitat. At each site, as much diversity as possible was sampled: even specimens that looked similar were individually numbered so that the presence of cryptic diversity could be tested. Importantly, a piece of tissue was cut for all specimens individually numbered for DNA extraction and the remainder of each specimen was relaxed (using magnesium chloride) and fixed (using 10% formalin or 70% ethanol) for comparative anatomy.

Specimens included in molecular analyses. DNA extraction numbers unique to each individual are indicated in phylogenetic trees as well as in lists of material examined and in figure captions (numbers are between brackets). Our molecular data set includes 190 Peronia individuals, only eleven of which correspond to COI sequences obtained from GenBank or BOLD (Table 2). Anatomical descriptions are based on those 179 Peronia individuals for which sequences were generated for the present study as well as the available type material for existing names (see below).

All DNA sequences for the eleven outgroups are from our previous studies (Dayrat et al. 2016, 2017, 2018, 2019a, b, c, d; Dayrat and Goulding 2017; Goulding et al. 2018a, b, c), with the exception of the nuclear sequences for Laspionchis boucheti, which are new (Table 2). Most Peronia mitochondrial and nuclear sequences in our molecular data set are new (Table 2). New mitochondrial COI and 16S sequences are provided for 169 individuals, and COI and 16S sequences for ten specimens are from a previous study (Dayrat et al. 2011; see below). In addition, all COI sequences in GenBank and BOLD closely related to Peronia sequences in our dataset were examined, and COI sequences for eleven individuals were selected to be included in phylogenetic analyses (Table 2; see below). All nuclear 28S and ITS2 sequences are new except for two individuals which have been used as outgroups in several of our previous studies: [696-2] from Okinawa and [706] from Hawaii (Table 2).

COI sequences publicly available. Eleven COI sequences obtained from GenBank (10) and BOLD (1) were added to our own data set (179 COI sequences) for a total of 190 sequences. Four COI sequences are from China (Sun et al. 2014), two from Singapore (Chang et al. 2018), two from Japan (Takagi et al. 2019), one from the Persian Gulf (unpublished), one from Gujarat, western India (unpublished), and one from Iran (Maniei et al. 2020a). All those sequences were merely referred to as Peronia sp., except for the specimens from China (referred to as Peronia verruculata), the specimen from western India (referred to as Onchidium verruculatum), and the specimen from Iran (recently described as P. persiae, a name regarded here as a synonym of P. verruculata, unit #4). Correct identifications are provided here for all those sequences (Table 2). Note that in the case of duplicate sequences available in GenBank, only one representative was selected. So, for instance, Chang et al. (2018) published many Peronia COI sequences that cluster in two mitochondrial units which they refer to as “Singapore clade” and “Peronia sp. 2 clade.” One sequence for their “Singapore clade” and one sequence for their “Peronia sp. 2 clade” are included here, which is enough to demonstrate that their two units correspond to our two mitochondrial units #1 and #3 of Peronia verruculata. Also, all COI sequences by Maniei et al. (2020a) for P. persiae cluster together within the unit #4 of P. verruculata, so only one of those individuals is included in our analyses: one individual is enough to demonstrate that P. persiae is a junior synonym of P. verruculata. A 16S sequence is available for the individual from Iran (Maniei et al. 2020a); no 16S sequences are available for any of the other COI sequences from GenBank and BOLD, so gaps were inserted in the mitochondrial concatenated alignment.

Vouchers used in Dayrat et al. (2011). Ten of our Peronia specimens were tentatively identified by Dayrat et al. (2011) at a time when nothing was known about the onchidiid species diversity in general and most especially in the genus Peronia. Most of those ten specimens were merely referred to with numbers (e.g., Peronia sp. 1). In order to avoid any confusion, those specimens are all included here so that correct species names are provided (Table 2). The specimen [443] (CASIZ 180486) identified as Peronia peronii from Guam really belongs to P. peronii. The specimen [696-2] (UF 352288) identified as Peronia cf. verruculata from Okinawa belongs to the new species P. okinawensis. The specimen [706] (UF 303653) identified as Peronia sp. 1 from Hawaii belongs to P. platei. The specimen [734] (AM C.459511) identified as Peronia sp. 3 from Queensland, Australia, belongs to the new species P. sydneyensis. Two specimens belong to P. madagascariensis: [735] (NHMUK 20060414) identified as Peronia cf. peronii from Mozambique, and [703] (UF 332088) identified as Peronia sp. 2 from Oman. Four specimens belong to P. verruculata: [712] (UF 368518) identified as Scaphis sp. from Cebu, Philippines, [730] (NHMUK 20080190) identified as Peronia sp. 4 from Mozambique, [733] (NHMUK 20060257) identified as Peronia sp. 5 from Mozambique, and [731] (NHMUK 20050628) identified as Peronia sp. 6 from Sulawesi, Indonesia.

Types of existing species-group names. All type specimens available for all onchidiid species-group names have been examined in context of the revision of the entire family. Comments on many onchidiid types can be found in our previous revisions (Dayrat et al. 2016, 2017, 2018, 2019a, b, c, d; Dayrat and Goulding 2017; Goulding et al. 2018a, b, c). In total, 118 type specimens (holotypes, lectotypes, paralectotypes, syntypes, etc.) are commented on here for the first time. Fifteen of those 118 type specimens are commented on in the general discussion because they are types of nomina dubia which may or may not refer to Peronia slugs. All the other (103) types are commented on in species descriptions because they are the types of 25 species-group names which must be classified in Peronia and which are not nomina dubia (Table 1).There are only two Peronia species names for which types could not be located: Scaphis lata Labbé, 1934a, and Paraperonia jousseaumei Labbé, 1934a. Finally, 14 lectotypes are designated here in order to clarify the application of 14 species names, usually because syntypes belong to different species or come from very distant localities.

Many type specimens were not labeled as types and were found within the general collections. In most cases, it was easy to determine that specimens were types because the information on the labels would match perfectly to that of the original descriptions. However, finding Labbé’s types was challenging, with the exception of the holotype, by monotypy, of Onchidium astridae Labbé, 1934b, preserved in Bruxelles (RBINS I.G.9223/MT.3822): it was not marked as a holotype, but the name Onchidium astridae is on the label, and the locality and collector information is matching.

The types of all the other Peronia species (and one subspecies) described by Labbé are preserved at the MNHN (the monograph in which those new taxa were described was almost exclusively based on material from the MNHN). The major issue with this material is that Labbé did not write any of his new species names on any of the labels. To be fair and fully accurate, there are actually three jars for which a specific name was written in pencil and in tiny letters on labels: one jar contains the type material of Onchidium durum (MNHN-IM-2000-33698), and two other jars contain part of the type material of Paraperonia gondwanae (MNHN-IM-2000-33683, MNHN-IM-2000-33688). Eleven years ago, Dayrat (2009) considered that identifying the types of Labbé’s onchidiid species names in the MNHN collection would be too risky (because specimens could be erroneously interpreted as types). However, after Virginie Héros (who is in charge of the Mollusk type collection at the MNHN) correctly remarked that it should still be possible to find some of Labbé’s types, an excel file was generated including all the old onchidiid material preserved at the MNHN and all the material cited in Labbé’s (1934a) monograph. By comparing various information (localities, names of the collectors, collecting dates, specimen sizes), it then became clear that many specimens could be identified as types with great confidence, even though they were not labeled as types and Labbé’s species names were not indicated on the labels.

For instance, originally, no jar clearly labeled as the type material of Scaphis carbonaria was found at the MNHN. However, of the old jars found at the MNHN with specimens from New Caledonia, only one matches perfectly the information provided in Labbé’s original description of S. carbonaria: an individual collected in 1880 by Réveillère (with an identification as Peronia). Other jars with one or more specimens from New Caledonia were collected by Fisher in 1878 and by François in 1894. Therefore, it is extremely likely that the specimen collected by Réveillère in 1880 is the holotype, by monotypy, of Scaphis carbonaria (MNHN-IM-2000-33708). In many cases, however, identifying the types happened to be much more challenging because there were several jars with the same locality, the same collector, and the same collecting date. In order to avoid any future confusion, Labbé’s types are commented on in great detail in species descriptions. There are only two of Labbé’s species for which no type material could be confidently traced back at the MNHN: Scaphis lata Labbé, 1934a, and Paraperonia jousseaumei Labbé, 1934a.

Finally, the type material of Peronia persiae, recently described by Maniei et al. (2020a), was not borrowed for examination. Regardless, there is no doubt that P. persiae is a junior synonym of both P. verruculata (Cuvier, 1830) and P. gondwanae (Labbé, 1934a) (Table 1), because all the COI and 16S sequences published for P. persiae cluster within the mitochondrial unit #4 of P. verruculata.

Additional material examined (historical museum collections). In addition to the 189 specimens included in the molecular analyses (not including the eleven outgroups) and the 118 type specimens of existing nominal species, 297 old specimens were obtained from museum collections from which no DNA could be extracted. Those specimens correspond to a total of 60 jars. One jar contains 161 specimens. All other jars contain fewer than 15 specimens. These old museum specimens are not included in the anatomical species descriptions, except for the description of Peronia verruculata from the Red Sea. Instead, these additional specimens are commented on in the species remarks. The additional specimens were especially useful to provide geographic records from places which could not be visited, such as the Chagos Archipelago, Nicobar Islands, Persian Gulf, and Socotra. Identifying Peronia species using only anatomical traits is challenging but possible (see below). Finally, some of the historical specimens from museum collections were studied by previous authors, and their re-examination allowed us to confirm or reject many identifications from the literature.

Size (length/width) is indicated in millimeters (mm) for each specimen. Both the external morphology and the internal anatomy were studied. All anatomical observations were made under a dissecting microscope and drawn with a camera lucida. Radulae and male reproductive organs were prepared for scanning electron microscopy (Zeiss SIGMA Field Emission Scanning Electron Microscopy). Radulae were cleaned in 10% NaOH for a week, rinsed in distilled water, briefly cleaned in an ultrasonic water bath (less than a minute), sputter-coated with gold-palladium and examined by SEM. Soft parts (penis, accessory penial gland, etc.) were dehydrated in ethanol and critical point dried before coating.

Anatomical species descriptions are based on those 179 Peronia individuals for which sequences were generated for the present study as well as on the available type material for species with existing names (see below). To avoid unnecessary repetition, the description of anatomical features that are virtually identical between Peronia species (e.g., nervous system, heart, and stomach) is not repeated for each species. However, all the characters that are useful for species comparison (e.g., intestinal loops and male apparatus) are described for every species. Special attention has been given to illustrating the holotype and the type locality of each new species.

Species are being described following a phylogenetic order. The detailed description of Peronia verruculata is based on the mitochondrial unit #1, by far the most widespread (from Peninsular Malaysia to the West Pacific) and most abundant (55 specimens in our study), but variations in the other units are precisely reported and figure captions indicate the unit to which each illustrated individual belongs.

In onchidiids, types of intestinal loops are defined based on the pattern of the intestine on the dorsal aspect of the digestive gland (with the digestive gland still in place). Plate (1893) first distinguished four types of intestinal loops (types I to IV) and Labbé (1934a) later added a type V. Only the types I and V are found in Peronia. Hoffmann (1928: 51, pl. 3, fig. 11) noted before Labbé that intestinal loops of type V differ from other types and he referred to them as type Ia. Labbé’s terminology (type V) is preferred because past authors have adopted it and because a type V is very different from a type I. The different types of intestinal loops and their individual variation are best revealed by coloring sections of the intestine differently (Dayrat et al. 2019b, c, d): a clockwise intestinal loop is colored in blue, a counterclockwise intestinal loop is colored in yellow, and a transitional loop between them is colored in green (Fig. 1).

The intestine first appears dorsally on the right side. In intestinal loops of type I, the intestine starts by forming a clockwise (blue) loop which does not make a complete circle. As a result, the transitional (green) loop is oriented to the right (Fig. 1A–F). In two species with intestinal loops of type I (P. okinawensis and P. peronii), the transitional loop is oriented between 12 and 3 o’clock (Fig. 1D–F). In the three other species with intestinal loops of type I (P. sydneyensis, P. verruculata, and P. willani), the transitional loop is oriented between 3 and 6 o’clock (Fig. 1A–C). In intestinal loops of type V, the intestine starts by forming immediately a counterclockwise (yellow) loop. In intestinal loops of type V, the counterclockwise loop is oriented between 10 and 11 o’clock (Fig. 1G–I). Four Peronia species are characterized by intestinal loops of type V: P. griffithsi, P. madagascariensis, P. platei, and P. setoensis.

Figure 1. 

Intestinal types found in Peronia species. A clockwise intestinal loop is colored in blue, a counterclockwise intestinal loop is colored in yellow, and a transitional loop between them is colored in green. The big red arrow indicates the orientation of the transitional loop (A–F) or counterclockwise loop (G–I), and the small black arrows indicates the direction of the intestinal transport A type I, with a transitional loop oriented at 3 o’clock, P. sydneyensis, Australia, New South Wales, [1517] (AM C.468915.001) B type I, with a transitional loop oriented at 6 o’clock, P. verruculata, lectotype of P. anomala, Red Sea (MNHN-IM-2000-33678) C type I, with a transitional loop oriented between 4 and 5 o’clock, P. verruculata, lectotype, Red Sea (MNHN-IM-2000-22941) D type I, with a transitional loop oriented at 3 o’clock, P. peronii, paralectotype of Onchidium peronii, Timor (MNHN-IM-2000- 22938) E type I, with a transitional loop oriented at 1 o’clock, P. peronii, lectotype of Paraperonia fidjiensis, Fiji (MNHN-IM-2000-33692) F type I, with a transitional loop oriented between 1 and 2 o’clock, P. okinawensis, holotype, Japan, Okinawa, [696-4 H] (UF 352288) G type V, with a counterclockwise loop oriented between 10 and 11 o’clock, P. madagascariensis, Madagascar, [5501] (MNHN-IM-2009-16392) H type V, with a counterclockwise loop oriented at 10 o’clock, P. platei, lectotype, French Polynesia (SMNH-Type-7537) I type V, with a counterclockwise loop oriented at 11 o’clock, P. griffithsi, holotype, Mauritius, [3157 H] (MNHN-IM-2000-35265). Scale bars: 2 mm (A, B), 5 mm (C–G), 3 mm (H, I).

DNA was extracted using a phenol-chloroform extraction protocol with cetyltrimethyl-ammonium bromide (CTAB). The mitochondrial cytochrome c oxidase I region (COI) and 16S region were amplified using the following universal primers (all 5’-3’): LCO1490 GGT CAA CAA ATC ATA AAG ATA TTG G, and HCO2198 TAA ACT TCA GGG TGA CCA AAR AAY CA (Folmer et al. 1994), 16Sar-L CGC CTG TTT ATC AAA AAC AT (Palumbi 1996), and the modified Palumbi primer 16S 972R CCG GTC TGA ACT CAG ATC ATG T (Dayrat et al. 2011). The nuclear ITS2 and 28S regions were amplified with the following primers: LSU-1 CTA GCT GCG AGA ATT AAT GTG A, and LSU-3 ACT TTC CCT CAC GGT ACT TG (Wade and Mordan 2000), 28SC1 ACC CGC TGA ATT TAA GCA T (Hassouna et al. 1984), and 28SD3 GAC GAT CGA TTT GCA CGT CA (Vonnemann et al. 2005). The 25 μl PCRs for COI and 16S contained 15.8 μl of water, 2.5 μl of 10× PCR Buffer, 1.5 μl of 25 mM MgCl₂, 0.5 μl of each 10 μM primer, 2 μl of dNTP Mixture, 0.2 μl (1 unit) of TaKaRa Taq (Code No. R001A), 1 μl of 20 ng/μl template DNA, and 1 μl of 100× BSA (Bovine Serum Albumin). The PCRs for ITS2 used the reagents in the same amounts as COI and 16S, except that water was reduced to 14.8 μl and the amount of 100× BSA was increased to 2 μl. The PCRs for 28S included 14.8 μl of water, 2.5 μl of 10× PCR Buffer, 0.5 μl of each 10 μM primer, 1 μl of dNTP Mixture, 5 μl of Q solution (which includes MgCl₂) and 0.5 μl of 20 ng/μl template DNA. The thermoprofile used for COI and 16S was: 5 minutes at 94 °C; 30 cycles of 40 seconds at 94 °C, 1 minute at 46 °C, and 1 minute at 72 °C; and a final extension of 10 minutes at 72 °C. The ITS2 thermoprofile was: 1 minute at 96 °C; 35 cycles of 30 seconds at 94 °C, 30 seconds at 50 °C, and 1 minute at 72 °C; and a final extension of 10 minutes at 72 °C. The 28S thermoprofile was: 4 minutes at 94 °C; 38 cycles of 50 seconds at 94 °C, 1 minute at 52 °C, and 2 minutes 30 seconds at 72 °C; and a final extension of 10 minutes at 72 °C. The PCR products were cleaned with ExoSAP-IT (Affymetrix, Santa Clara, CA, USA) prior to sequencing. Untrimmed sequenced fragments represented approximately 680 bp for COI, 530 bp for 16S, 740 bp for ITS2, and 1030 bp for 28S.

Chromatograms were consulted to resolve rare ambiguous base calls. DNA sequences were aligned using Clustal W in MEGA 7 (Kumar et al. 2016). Eleven onchidiid species outside Peronia were selected as outgroups from our previous studies (Dayrat et al. 2011, 2016, 2017, 2018, 2019a, b, c, d; Dayrat and Goulding 2017; Goulding et al. 2018a, b, c): Alionchis jailoloensis Goulding & Dayrat in Goulding et al. 2018a, Laspionchis boucheti Dayrat & Goulding in Dayrat et al. 2019b; Marmaronchis vaigiensis (Quoy & Gaimard, 1825), Melayonchis eloisae Dayrat in Dayrat et al. 2017, Onchidella celtica (Cuvier in Audouin and Milne-Edwards 1832), Onchidina australis (Semper, 1880), Onchidium typhae Buchannan, 1800, Paromoionchis tumidus (Semper, 1880), Peronina tenera (Stoliczka, 1869), Platevindex luteus (Semper, 1880), and Wallaconchis sinanui Goulding & Dayrat in Goulding et al. 2018b. All new DNA sequences were deposited in GenBank and vouchers deposited in museum collections (Table 2). The ends of each alignment were trimmed. Alignments of mitochondrial (COI and 16S) sequences and nuclear (ITS2 and 28S) sequences were concatenated separately in order to test whether these two data sets support the same relationships. The concatenated mitochondrial alignment included 1014 nucleotide positions: 614 (COI) and 400 (16S). The concatenated ITS2 and 28S alignment included 1544 nucleotide positions: 535 (ITS2) and 1009 (28S). The haplotype ITS2 alignment (in which identical sequences were grouped into a single haplotype sequence) included 740 nucleotide positions.

Three independent sets of phylogenetic analyses were performed: 1) Maximum Likelihood and Bayesian analyses with concatenated mitochondrial COI and 16S sequences; 2) Maximum Parsimony analyses with concatenated nuclear ITS2 and 28S sequences; 3) Maximum Parsimony analyses with ITS2 haplotype sequences. Maximum Parsimony analyses were conducted in PAUP v 4.0 (Swofford 2002) with gaps coded as a fifth character state, and 100 bootstrap replicates conducted using a full heuristic search. Prior to Maximum Likelihood and Bayesian phylogenetic analyses, the best-fitting evolutionary model was selected for each locus separately using the Model Selection option from Topali v2.5 (Milne et al. 2004): a GTR + G model was independently selected for COI and 16S. Maximum Likelihood analyses were performed using PhyML (Guindon and Gascuel 2003) as implemented in Topali. Node support was evaluated using bootstrapping with 100 replicates. Bayesian analyses were performed using MrBayes v3.1.2 (Ronquist and Huelsenbeck 2003) as implemented in Topali, with five simultaneous runs of 1.5×10⁶ generations each, sample frequency of 100, and burn in of 25% (and posterior probabilities were also calculated). Topali did not detect any issue with respect to convergence. All analyses were run several times and yielded the same result.

In addition, genetic distances between COI sequences were calculated in MEGA 7 as uncorrected p-distances. COI sequences were also translated into amino acid sequences in MEGA using the invertebrate mitochondrial genetic code to check for the presence of stop codons (no stop codon was found).

The monophyly of Peronia is strongly supported in all analyses except in the mitochondrial ML analyses (bootstrap of 58), which confirms that all onchidiid slugs with dorsal gills belong to the same clade (Figs 2–4).

Figure 2. 

Phylogenetic relationships between Peronia species based on concatenated mitochondrial COI and 16S DNA sequences. Numbers by the branches are the bootstrap values (maximum likelihood analysis) and the posterior probabilities (Bayesian analysis). Only the values > 50% (ML) and > 0.9 (Bayesian) are indicated. Numbers for each individual correspond to unique identifiers for DNA extraction. Information on specimens can be found in the lists of material examined and in Table 2. The color used for each species or mitochondrial unit is the same as the color used in Figs 3–6.

Figure 2. 

Continued.

Figure 2. 

Continued.

Seven nodes of higher relationships among Peronia species are well supported. Supports are indicated here in parentheses in the following order: ML bootstrap in mitochondrial analysis, Bayesian posterior probability in mitochondrial analysis, bootstrap in ITS2 analysis, bootstrap in ITS2 and 28S analysis (bootstrap values below 50% and posterior probabilities below 0.90 are replaced by a dash). Most basally, Peronia is always split in clades A and B. Clade A is strongly supported (99, 1.0, 100, 100) and includes P. peronii and P. okinawensis. Clade B is also strongly supported (99, 1, 93, 99) and includes clade C and P. madagascariensis as its most basal species. Clade C, which is consistently recovered but moderately supported (-, -, 90, 87), includes clade D and P. platei as its most basal species. Clade D (98, 1, 99, 86) includes the three clades E, F, and G, of which the relationships are unresolved (Fig. 4) or incongruent (Figs 2, 3). Clade E (83, 0.9, 80, 54) includes P. griffithsi and P. setoensis. Clade F (100, 1.0, 75, 93) includes P. sydneyensis and P. willani. Clade G (97, 1, 71, 94) includes the five least-inclusive mitochondrial units of P. verruculata. The relationships of those five units are not resolved (all support values are very low).

Figure 3. 

Maximum parsimony consensus tree within Peronia based on ITS2 sequences (identical sequences are represented as a single haplotype sequence). Numbers by the branches are the bootstrap values. Only the values > 50% are indicated. Numbers for each individual correspond to unique identifiers for DNA extraction. Information on specimens can be found in the lists of material examined and in Table 2. The color used for each species or mitochondrial unit is the same as the color used in Figs 2, 4–6.

The monophyly of each species recognized here is strongly supported in all analyses, except for the special case of P. sydneyensis (see below, species delineation). Within four species, some least-inclusive units are supported by the mitochondrial markers but not by comparative anatomy and nuclear markers (Figs 2–4): two units within P. peronii (one unit from Mauritius and the other from the West Pacific); two units within P. platei (one unit from Hawaii and the other from Papua New Guinea); two units within P. griffithsi (one unit from Mauritius and the other from Kei Islands and Papua New Guinea); and three units of P. verruculata from South-East Asia and the West Pacific (units #1, #2, and #3). Two least-inclusive mitochondrial units within P. verruculata from the western Indian Ocean (units #4 and #5) are also monophyletic in nuclear analyses (Figs 2–4) but are anatomically cryptic (see below). Note that populations of P. verruculata from the Red Sea are not represented in molecular analyses (see below, species delineation).

In mitochondrial analyses (Fig. 2), P. sydneyensis and P. willani form together the strongly supported clade E and the monophyly of each species is also strongly supported. In nuclear analyses (Figs 3, 4), they also form a strongly supported clade but P. sydneyensis is paraphyletic with respect to P. willani. Both species are close geographically (P. sydneyensis is distributed in New South Wales, Queensland and New Caledonia, and P. willani is distributed in the Northern Territory) and may be the result of a recent divergence. The paraphyly in nuclear analyses most likely is the result of incomplete lineage sorting (see below, species delineation).

Figure 4. 

Maximum parsimony consensus tree within Peronia based on concatenated nuclear ITS2 and 28S sequences. Numbers by the branches are the bootstrap values. Only the values > 50% are indicated. Numbers for each individual correspond to unique identifiers for DNA extraction. Information on specimens can be found in the lists of material examined and in Table 2. The color used for each species or mitochondrial unit is the same as the color used in Figs 2, 3, 5, 6.

Pairwise genetic distances were calculated for a total of 13 units (Fig. 5, Table 3): the five mitochondrial units within P. verruculata as well as the eight other species. A barcode gap is found in all cases, apart from the mitochondrial unit #1 of P. verruculata.

Figure 5. 

Diagram to help visualize the data on pairwise genetic distances between COI sequences within and between species and mitochondrial units (P. verruculata) in Peronia (see Table 3). Ranges of minimum to maximum distances are indicated (in percentages). For instance, within P. willani, individual sequences are between 0 and 1.9% divergent; individual sequences between P. willani and the other species or units are minimally 4.3% and maximally 16.8% divergent. The colors are the same as those used in Figs 2–4, 6.

All Peronia slugs are characterized by dorsal gills which are not found in other onchidiids. They are also all characterized by a unique combination of internal traits: they are the only onchidiid slugs with intestinal loops of type I or V, an accessory penial gland, and no rectal gland. The fact that any slug with this combination of traits belongs to a Peronia species is helpful to identify specimens with dorsal gills retracted inside the notum.

There are no external differences between Peronia species. In the field, it is not possible to reliably identify any of them, especially because sympatric species are often found together at the exact same sites. Individuals of very large size (longer than 100 mm) are only found in P. peronii, but smaller individuals are impossible to distinguish externally from other species. Also, tall papillae over the entire notum seem to be mostly found in P. peronii and P. madagascariensis, but that may be due to the fact that slugs of both species are the largest, and it remains difficult to define exactly what a tall papilla is because papilla size is highly variable.

Internal differences help identify some species reliably, but not all (Table 4). Internal differences are almost exclusively based on combinations of traits because no Peronia species is characterized by any unique, distinctive feature, except for P. peronii (characterized by a spine of the accessory penial gland longer than 3 mm) and P. sydneyensis (characterized by strong protuberances on the spine of the accessory penial gland), and it remains difficult to identify Peronia species anatomically. For instance, where they overlap geographically (Queensland and New Caledonia), P. verruculata and P. sydneyensis can only be distinguished based on the length of the spine of the accessory penial gland, the presence of strong protuberances near the tip of the spine of the accessory penial gland, and the length of the penial hooks, which are all traits that are hardly accessible to a non-expert. However, only two Peronia species are cryptic externally and internally: P. setoensis and P. platei, which are not sister taxa (Figs 2–4) and do not overlap geographically, at least based upon current data (Fig. 6). Finally, the mitochondrial units of P. verruculata cannot be reliably distinguished anatomically.

Figure 6. 

Geographical distribution of the Peronia species A distribution of all Peronia species except for P. peronii B distribution of P. peronii. The colors are the same as those used in Figs 2–5. Colored areas correspond to hypothetical geographical ranges based on confirmed records only. Distinct colors are used for each unit of P. verruculata. The distribution of P. verruculata in the Indo-West Pacific is actually continuous. However, because it is unclear which units are present in regions from where we have no fresh material of P. verruculata (red areas), no unit of P. verruculata is shown there. For P. peronii, black dots correspond to material identified based on anatomical characters and blue dots correspond to material with DNA sequences. Details on species distribution can be found in each species description.

Types of intestinal loops are useful for the identification of Peronia species (Fig. 1): species are characterized by intestinal loops of type V (P. griffithsi, P. madagascariensis, P. platei, and P. setoensis), type I with a transitional loop oriented between 12 and 3 o’clock (P. okinawensis and P. peronii), or type I with a transitional loop oriented between 3 and 6 o’clock (P. sydneyensis, P. verruculata, P. willani). Exceptions exist but are remarkably rare: only one individual in P. sydneyensis was found with a transitional loop slightly outside the range of that species (at 2 o’clock). Types of intestinal loops, however, can only be used in combination with other traits for the purpose of species identification.

The insertion of the retractor muscle of the penis is not very useful in identification because it mostly matches the distribution of the respective intestinal loop types (Table 4). An insertion near the heart is only found in the two species with intestinal loops of type I and transitional loops oriented between 12 and 3 o’clock (P. peronii and P. okinawensis). Within each species, all individuals share the same insertion of the retractor muscle (either near the heart or at the posterior end of the visceral cavity). However, in P. griffithsi, which is widely distributed from the West Pacific to Mauritius, individuals are characterized by both insertions. In P. peronii, the retractor muscle can exceptionally be vestigial (with no clear insertion).

The length of the muscular sac of the accessory penial gland varies depending on the size of animals, but it is useful to help identify some species. Indeed, only two species (P. peronii and P. willani) are characterized by a muscular sac which is longer than 20 mm (Table 4). The length of the spine of the accessory penial gland is helpful to distinguish closely related species which, otherwise, are very similar anatomically: P. peronii (at least 3 mm) and P. okinawensis (less than 2.3 mm); P. setoensis (more than 0.9 mm) and P. griffithsi (less than 0.62 mm); and P. sydneyensis (less than 1 mm) and P. willani (more than 1.5 mm). The diameter of the spine at its base can be used in exactly the same way. The length of penial hooks also differs between species: the longest hooks are found in P. madagascariensis (up to 100 μm), the shortest in P. setoensis and P. griffithsi (less than 25 μm).

The delineation of Peronia species is straightforward. They are all supported by independent data sets: they are reciprocally monophyletic with both mitochondrial and nuclear markers, and their monophyly is strongly supported; they are all separated by a large barcode gap; and they each are characterized by a unique combination of anatomical traits (with the exception of P. setoensis and P. platei, which are cryptic). Only two species need special attention: P. sydneyensis and P. verruculata.

The paraphyly of P. sydneyensis with respect to P. willani in nuclear analyses most likely is the result of incomplete lineage sorting, because lineage sorting progresses more rapidly for mitochondrial alleles than for nuclear alleles (Funk and Omland 2003). Also, P. sydneyensis and P. willani species are clearly distinct anatomically: in particular, P. sydneyensis is characterized by unique, strong protuberances near the tip of the spine of the accessory penial gland. Therefore, P. sydneyensis and P. willani are regarded as two recent but well-delineated species.

Despite some genetic structure, Peronia verruculata is regarded as a single species for various reasons. In mitochondrial analyses, P. verruculata is split in five least-inclusive mitochondrial units of which the relationships are basically unresolved due to low support (Fig. 2). In nuclear analyses, the mitochondrial units #1, #2, and #3 are not monophyletic (Figs 3, 4). Therefore, they should not be recognized as distinct taxa. In nuclear analyses, units #4 and #5 are monophyletic (Figs 3, 4). However, the mitochondrial units #1, #2, and #3 do not form a monophyletic group with respect to units #4 and #5. Recognizing mitochondrial units #4 and #5 each as a separate taxon would mean that mitochondrial units #1, #2, and #3 would also have to be recognized as separate taxa, which is unwarranted for the reasons given above.

All mitochondrial units of P. verruculata are cryptic anatomically (their anatomical traits display overlapping variation) while P. verruculata is clearly distinct from other Peronia species (Table 4). Finally, it would seem premature to recognize units #4 and #5 as independent lineages because our geographical sampling of P. verruculata is not continuous (Fig. 6). Future samples from southern India (including Sri Lanka) or the Arabian Sea (the coast of Yemen, Oman, Somalia) might show that the individuals of the western mitochondrial units #4 and #5 can still interbreed, exactly like units #1, #2, and #3. We therefore refrain from naming those five mitochondrial units within P. verruculata. They are merely regarded as mitochondrial units that indicate some genetic structure, but the current data do not suggest that they should be recognized as distinct taxa. Note that taxon names are already available for the mitochondrial units #1, #4, and #5 of P. verruculata (Table 1).

Finally, note that Peronia verruculata was described from the Red Sea, from which no fresh material could be obtained. However, the specimens examined from the Red Sea are anatomically indistinguishable from the specimens of the five mitochondrial units of P. verruculata (Table 4). Therefore, at this stage, there is no reason to think that the populations from the Red Sea belong to a distinct species and that the name P. verruculata cannot apply to the whole species from the Red Sea and South Africa all the way to the West Pacific (Japan, New Caledonia, and Queensland). At any rate, there are plenty of available names that can be used in the future if it were to be demonstrated that the populations from the Red Sea belong to a distinct species (see remarks on P. verruculata).

Geographic distribution is discussed in detail with each species description. The map of species distributions only illustrates the records that are regarded as correct (Fig. 6). Most of those correct records correspond to the specimens included in our molecular analyses. However, they also include types as well as historical museum specimens and records from the literature which could be positively identified using anatomical traits (e.g., intestinal loops, length of the spine of the accessory penial gland).

The secondary literature was read with great attention, especially in cases where it could provide geographical records not included in our material. Every record found in the literature is commented on (in the species remarks). Records from the literature are certainly not taken for granted because the secondary literature is plagued with two major issues. First, past authors did not always take the time to examine type specimens. For instance, Labbé (1934a) did not examine the types of Onchidium verruculatum and Onchidium peronii which are preserved at the Paris Museum (he did not list them in the material examined for these species), even though his study was almost exclusively based on material from that institution. Second, because there was no proper knowledge about intraspecific character variation, nobody knew which character could help distinguish species or not. For instance, Hoffmann’s (1928: 73) record of Peronia verruculata from Hawaii was never questioned, but Peronia slugs from Hawaii are all characterized by intestinal loops of type V, which means that they cannot belong to P. verruculata (which is characterized by intestinal loops of type I).