Two new species of Tardigrada from moss cushions (Grimmia sp.) in a xerothermic habitat in northeast Tennessee (USA, North America), with the first identification of males in the genus Viridiscus

Diane R. Nelson; Rebecca Adkins Fletcher; Roberto Guidetti; Milena Roszkowska; Daria Grobys; Łukasz Kaczmarek

doi:10.7717/peerj.10251

Two new species of Tardigrada from moss cushions (Grimmia sp.) in a xerothermic habitat in northeast Tennessee (USA, North America), with the first identification of males in the genus Viridiscus

Diane R. Nelson ¹, Rebecca Adkins Fletcher², Roberto Guidetti³, Milena Roszkowska⁴, Daria Grobys⁵, Łukasz Kaczmarek⁶

1Department of Biological Sciences, East Tennessee State University, Johnson City, TN, United States of America

2Department of Appalachian Studies, East Tennessee State University, Johnson City, TN, United States of America

3Department of Life Sciences, University of Modena and Reggio Emilia, Modena, Italy

4Department of Animal Taxonomy and Ecology, Department of Bioenergetics, Adam Mickiewicz University, Poznań, Poland

5Department of Bioenergetics, Adam Mickiewicz University, Poznań, Poland

6Department of Animal Taxonomy and Ecology, Adam Mickiewicz University, Poznań, Poland

DOI: 10.7717/peerj.10251

Published: 2020-11-23
Accepted: 2020-10-06
Received: 2020-06-29

Academic Editor: Michael Wink

Subject Areas: Biodiversity, Ecology, Molecular Biology, Taxonomy, Zoology
Keywords: Dioecious, Eutardigrada, Heterotardigrada, Systematics, Taxonomy, Water bears, Morphology, Molecular technique, Parthenogenesis, Anhydrobiosis

Copyright: © 2020 Nelson et al.
Licence: This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, reproduction and adaptation in any medium and for any purpose provided that it is properly attributed. For attribution, the original author(s), title, publication source (PeerJ) and either DOI or URL of the article must be cited.

Cite this article: Nelson DR, Adkins Fletcher R, Guidetti R, Roszkowska M, Grobys D, Kaczmarek Ł. 2020. Two new species of Tardigrada from moss cushions (Grimmia sp.) in a xerothermic habitat in northeast Tennessee (USA, North America), with the first identification of males in the genus Viridiscus. PeerJ 8:e10251 https://doi.org/10.7717/peerj.10251

Abstract

Background

The phylum Tardigrada consists of over 1,300 species that inhabit terrestrial, freshwater and marine environments throughout the world. In terrestrial habitats they live primarily in mosses, lichens, leaf litter and soil, whereas tardigrades in freshwater and marine environments are mainly found in sediments and on aquatic plants. More than 65 species have been previously reported in the state of Tennessee, USA.

Methods

Tardigrades present in moss cushions (Grimmia sp.) collected from a xerothermic habitat on the East Tennessee State University campus, Johnson City, TN, USA, were extracted, mounted on slides, identified, and counted. Additional samples of fresh dried moss were used for integrative analyses, including morphological analysis with phase contrast (PCM) and scanning electron microscopy (SEM), as well as molecular analyses of COI, 18S rRNA, 28S rRNA, and ITS-2 of the Macrobiotus and Milnesium species.

Results

Five species were found, including two species new to science: Viridiscus miraviridis sp. nov. and Macrobiotus basiatus sp. nov. Viridiscus miraviridis sp. nov. differs from other members of the genus mainly by having a different type of dorsal cuticle and some other, more subtle, morphometric characters. In addition to the two new species, Viridiscus perviridis and Viridiscus viridissimus were present, and males of Vir. viridissimus were found for the first time, the first record of males in the genus Viridiscus. Macrobiotus basiatus sp. nov. is most similar to Macrobiotus nelsonae, but it differs from Mac. nelsonae mainly by the stylet supports being situated in a more anterior position, shorter and narrower egg processes, and a smaller number of areoles around the egg processes. Moreover, the identification of Milnesium inceptum was confirmed as the first record for the USA by analysis of COI.

Introduction

The phylum Tardigrada consists of over 1,300 species (Guidetti & Bertolani, 2005; Degma & Guidetti, 2007; Degma, Bertolani & Guidetti, 2009–2020) that inhabit terrestrial, freshwater and marine environments throughout the world. In terrestrial habitats they live primarily in mosses, lichens, leaf litter and soil, whereas in freshwater and marine environments tardigrades are found mainly in sediments and on aquatic plants (Nelson, Guidetti & Rebecchi, 2015).

Studies on the distribution of terrestrial and aquatic tardigrades have been conducted primarily in Europe and North America (see e.g., McInnes, 1994; Meyer, 2013; Kaczmarek, Michalczyk & McInnes, 2014; Kaczmarek, Michalczyk & McInnes, 2015; Kaczmarek, Michalczyk & McInnes, 2016; McInnes, Michalczyk & Kaczmarek, 2017). In addition to a few specimens reported by Riggin (1962) and Maucci (1987), surveys of the distribution of tardigrades in Tennessee have been conducted by Nelson (1975) and her students and colleagues (Kathman & Nelson, 1987; Nelson, Kincer & Williams, 1987; Nelson & McGlothlin, 1993; Nelson & McGlothlin, 1996; Kendall-Fite & Nelson, 1996; Guidetti, 1998; Bertolani, Marley & Nelson, 1999; Guidetti, Bertolani & Nelson, 1999; Nelson, Marley & Bertolani, 1999; Nelson & Adkins, 2001; Bartels & Nelson, 2006; Bartels & Nelson, 2007; Bartels & Nelson, 2012; Bartels et al., 2007; Bartels et al., 2008; Bartels et al., 2009; Bartels et al., 2011; Bartels, Nelson & Exline, 2011; Bartels et al., 2014; Bartels, Mormino & Nelson, 2017; Nelson & Bartels, 2007; Nelson & Bartels, 2013; Nelson, Bartels & Guil, 2019). Thus far for the state of Tennessee, more than 65 species have been reported (Kaczmarek, Michalczyk & McInnes, 2016; Meyer, 2013).

The genus Viridiscus was recently established by Gąsiorek & Michalczyk in Gąsiorek et al. (2019) in an integrative analysis of the “arctomys group” of Echiniscus, in which five new genera were erected. The widely distributed “Echiniscus viridis group” (now Viridiscus), with the very characteristic fully or partially green cuticle and lacking all lateral and dorsal appendages except cirrus A currently includes six species: Viridiscus clavispinosus (Fontoura, Pilato & Lisi, 2011), Viridiscus perviridis (Ramazzotti, 1959), Viridiscus rufoviridis (Du Bois Reymond Marcus, 1944), Viridiscus viridianus (Pilato, Fontoura & Lisi, 2007), Viridiscus viridis (Murray, 1910), and Viridiscus viridissimus (Péterfi, 1956). In this paper, we provide a description of a new species of Viridiscus, and the first confirmation of the presence of males in the genus.

We also provide the first record of Milnesium inceptum Morek et al., 2019 from the USA (Tennessee, Washington County), confirmed by molecular analysis. Milnesium Doyère, 1840 was originally believed to be a monospecific genus, but multiple species have been described in recent years with over 40 known species to date (Morek et al., 2020).

In addition, based on an integrative approach, we also provide a description of a new species in the genus Macrobiotus. The cosmopolitan genus Macrobiotus C.A.S. Schultze, 1834, the first described genus of tardigrades, comprises a large complex of species, some of which have been allocated to several new genera in recent years.

Materials & Methods

Sample processing

In a previous study (Nelson & Adkins, 2001), 60 samples of the moss Grimmia sp. were collected from vertical surfaces of concrete caps on six brick fence posts (10 samples per post), fully exposed to sun, wind and precipitation, located on the north side of the East Tennessee State University campus (Johnson City, Washington County, Tennessee, USA) (Figs. 1A–1C), from 14 May 1995 to 30 September 1995 (for more details see Nelson & Adkins, 2001).

Sampling site and type locality of Viridiscus miraviridis sp. nov. and Macrobiotus basiatus sp. nov. in moss cushions (Grimmia sp.) on the campus of East Tennessee State University, Johnson City, Tennessee. — Figure 1: Sampling site and type locality of *Viridiscus miraviridis* sp. nov. and *Macrobiotus basiatus* sp. nov. in moss cushions (*Grimmia* sp.) on the campus of East Tennessee State University, Johnson City, Tennessee.
(A) Overall view of one of the brick fence posts on the campus. (B) Close up of the concrete cap on the fence post. (C) Close up of the moss showing the cushion growth form and leaf structures. Photo credit: Diane R. Nelson.

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DOI: 10.7717/peerj.10251/fig-1

The samples were soaked in tap water in a small jar for 30 min, then agitated vigorously. Afterwards, the sample was poured into a beaker through a wire screen to remove the moss. When the beaker contents settled, the top layer of water was decanted, and boiling alcohol was added to preserve the sample. The beaker contents were then poured through a 45 µm sieve. The remaining sample on the sieve was washed into a petri dish and examined for tardigrades under a stereomicroscope. All tardigrades and eggs found in each sample were extracted with an Irwin loop. To describe the two new species with an integrative approach, additional samples of freshly dried moss from the same posts were collected in 2019 and mailed to Adam Mickiewicz University, Poznań, Poland, for processing, imaging and morphometrics.

Microscopy and imaging

Specimens from the previous study were mounted on microscope slides in Hoyer’s mounting medium, with the coverslip sealed with epoxy paint when dry. All specimens and eggs were identified to species level with phase contrast light microscopy (PCM). Microphotographs of specimens from the fresh samples were obtained with Olympus BX41 associated with an ARTCAM–300Mi digital camera (Olympus Corporation, Shinjuku–ku, Japan). A total of 10 adults of Viridiscus miraviridis sp. nov. and 25 specimens and 15 eggs of Macrobiotus basiatus sp. nov. from the fresh samples were prepared for SEM imaging according to Roszkowska et al. (2018). These animals and eggs were examined under high vacuum in a Hitachi S3000N Scanning Electron Microscope.

All figures were assembled in Corel Photo-Paint 17. For deep structures that could not be fully focused in a single photograph, a series of 2–10 images were taken every ca. 0.5 µm and then assembled into a single stacked image manually in Corel Photo-Paint 17.

Morphometrics and morphological nomenclature

Structures were measured only if their orientation was suitable. Body length was measured from the anterior extremity to the end of the body, excluding the hind legs. The types of buccal-pharyngeal apparatuses and claws were classified according to Pilato & Binda (2010). The terminology used to describe the genus Viridiscus followed Gąsiorek et al. (2019). Lengths of the claw branches for Viridiscus were measured from the base of the claw to the top of the branch. The sp index, the ratio of the length of a given structure to the length of the scapular plate (scp), was calculated according to Dastych (1999). All measurements of adults and eggs of Macrobiotus were prepared according to Kaczmarek & Michalczyk (2017). Terminology describing the oral cavity armature followed Michalczyk & Kaczmarek (2003). The macroplacoid length sequence was determined according to Kaczmarek et al. (2014). The pt ratio, the length of a given structure to the length of the buccal tube expressed as a percentage (Pilato, 1981), was also calculated. The age structure of the Viridiscus populations was determined according to Bertolani et al. (1984): 1st instar, anus and gonopore absent, 2 claws per leg present; 2nd instar, anus present but gonopore absent or rudimentary, 4 claws per leg present; adult, anus and gonopore present, 4 claws per leg present.

Morphometric data were handled using the “Parachela” ver. 1.1 and “Echiniscoidea” ver. 1.1 templates available from the Tardigrada Register (Michalczyk & Kaczmarek, 2013). Tardigrade taxonomy followed Bertolani et al. (2014). Genera were abbreviated according to Perry, Miller & Kaczmarek (2019).

Genotyping

All specimens of Macrobiotus and Milnesium were preliminarily identified using light microscopy before DNA extraction. DNA was extracted from individual animals following a Chelex^®100 resin (Bio-Rad) extraction method (Casquet, Thebaud & RG, 2012) modified to obtain tardigrade exoskeletons according to Kaczmarek et al. (2019). For Macrobiotus specimens, we sequenced four DNA fragments differing in mutation rates: the cytochrome oxidase subunit I (COI, mtDNA), the internal transcribed spacer (ITS-2, nDNA), the large ribosome subunit (28S rRNA, nDNA) and the small ribosome subunit (18S rRNA, nDNA). For Milnesium specimens we sequenced only the COI fragment. All fragments were amplified and sequenced with the protocols described in Kaczmarek et al. (2019). Amplification of 28S rRNA and 18S rRNA fragments was conducted in the reaction mixture with addition of 3% DMSO. Primers used for PCR, and later sequencing, with their original references, are listed in Table 1. PCR programs used for the amplification of COI, ITS-2, 28S rRNA and 18S rRNA are listed in Table 2. For analysis of COI, PCR products were purified by thermosensitive Exonuclease I and FastAP Alkaline Phosphatase (Fermentas, Thermo Scientific), whereas ITS-2, 28S rRNA and 18S rRNA (after excision from the gel) were treated with NucleoSpin Gel and PCR Clean-up Kit (MACHERY-NAGEL). For sequencing COI, ITS-2 and 28S rRNA, the same primers were used as in the amplification reaction, while a few additional primers were required to sequence the 18S rRNA (see Table 1).

Table 1:

Primers used for amplification and sequencing of DNA fragments.

DNA fragment	Application	Direction	Code	Sequence (5′–3′)	Reference
COI	PCR & sequencing	Forward	LCO 1490	GGTCAACAAATCATAAAGATATTGG	Folmer et al. (1994)
COI	PCR & sequencing	Reverse	HCO 2198	TAAACTTCAGGGTGACCAAAAAATCA	Folmer et al. (1994)
ITS-2	PCR & sequencing	Forward	ITS2_Eutar_Ff	CGTAACGTGAATTGCAGGAC	Stec, Kristensen & Michalczyk (2018)
ITS-2	PCR & sequencing	Reverse	ITS2_Eutar_Rr	TGATATGCTTAAGTTCAGCGG	Stec, Kristensen & Michalczyk (2018)
28S rRNA	PCR & sequencing	Forward	28SF0001	ACCCVCYNAATTTAAGCATAT	Mironov, Dabert & Dabert (2012)
28S rRNA	PCR & sequencing	Reverse	28SR0990	CCTTGGTCCGTGTTTCAAGAC	Mironov, Dabert & Dabert (2012)
18S rRNA	PCR & sequencing	Forward	Euk-A	AACCTGGTTGATCCTGCCAGT	Koid et al. (2012)
	PCR & sequencing	Reverse	Euk-B	GATCCTTCTGCAGGTTCACCTAC	Koid et al. (2012)
	sequencing	Forward	fw390	AATCAGGGTTCGATTCCGGAGA	Dabert et al. (2010)
		Forward	for_770	ACTTTGAAAAAATTAGAGTGC
		Reverse	rev930	GACGGTCCAAGAATTTCAC
		Reverse	rev_1460	CATCACAGACCTGTTATTGC

DOI: 10.7717/peerj.10251/table-1

The sequences were edited and manually checked against non-conservative alignments using BioEdit, version 7.0.5. (Hall, 1999). The 18S rRNA sequence was assembled using the CAP contig assembly program implemented in BioEdit. All sequences were submitted to GenBank (see ‘Results’ section).

We were not able to obtain DNA from the Viridiscus specimens since more than one morphologically very similar species of Viridiscus were present and insufficient numbers were present in the new samples from the study sites.

Comparative molecular analysis

The obtained sequences were analysed by Standard Nucleotide BLAST to confirm their uniqueness. Then, a comparison was performed with the COI sequence of the genus Milnesium, as well as COI, ITS-2, 18S rRNA, and 28S rRNA sequences of the genus Macrobiotus, deposited in GenBank, using only the sequences of good quality and length. All sequences were aligned with the ClustalW Multiple Alignment tool (Thompson, Higgins & Gibson, 1994) implemented in BioEdit. Alignment sequences were trimmed to 759, 678, 276 and 518 bp for 18S rRNA (21 sequences, 17 species), 28S rRNA (18 sequences, 11 species), ITS2 (21 sequences, 12 species) and COI (32 haplotypes, 59 sequences, 16 species) barcodes, respectively, and calculation for the uncorrected p-distances was performed using MEGA X (Kumar et al., 2018). Uncorrected pairwise distances are provided in Table S1. Table S2 provides the name, accession number and link for the third-party database where the sequences have been deposited.

Table 2:

PCR programs used for the amplification of COI, ITS-2, 28S rRNA and 18S rRNA.

	COI and ITS-2			28S rRNA			18S rRNA
Step	Cycles	Time [min.:sec.]	Temp. [°C]	Cycles	Time [min:sec]	Temp. [°C]	Cycles	Time [min:sec]	Temp. [°C]
initial denaturation	–	03:00	98	–	05:00	98	–	05:00	98
denaturation	40	00:15	98	40	00:30	98	40	00:30	98
annealing		00:30	50		01:00	50		01:00	60
extension		00:30	72		01:00	72		01:30	72
final extension	–	07:00	72	–	07:00	72	–	07:00	72

DOI: 10.7717/peerj.10251/table-2

Comparative material

For comparison with the new species, the following specimens were used: Vir. viridis collected in Hawaii (Schuster collection at the Bohart Museum, University of California-Davis, USA); Vir. viridianus (paratype 060511PF) collected in Azores Is. (Fontoura Collection, Department of Zoology and Anthropology, University of Porto, Portugal); Vir. rufoviridis collected in Ecuador (slides 13775, 13779, 13814, 13817); Vir. viridissimus collected in Johnson City, TN, USA (slides 7731, 7735, 7843); Vir. viridis collected in Ecuador (slide 13821); Vir. perviridis collected in Johnson City, TN, USA (slide 7744) and in Italy (slide 4564) (Maucci collection at the Natural History Museum of Verona, Italy); and holotype and paratypes of Macrobiotus nelsonae (Guidetti, 1998) collected on Roan Mountain, Tennessee (slides 5N05A-S10) (Bertolani Collection, Department of Life Science, University of Modena and Reggio Emilia, Italy).

In addition to the key to Viridiscus in Fontoura, Pilato & Lisi (2011), the following original descriptions or redescriptions, as well as more recent papers, were used: Murray (1910), Du Bois Reymond Marcus (1944), Péterfi (1956), Ramazzotti (1959), Pilato, Fontoura & Lisi (2007); Pilato, Fontoura & Lisi (2008), and Morek et al. (2019).

For the differential diagnosis of the new Macrobiotus species, the following original descriptions or redescriptions were used: Maucci (1954), Guidetti (1998), Binda et al. (2001) and Meyer et al. (2017).

For the correct identification of Mil. inceptum, we used the original description in Morek et al. (2019).

The electronic version of this article in Portable Document Format (PDF) will represent a published work according to the International Commission on Zoological Nomenclature (ICZN), and hence the new names contained in the electronic version are effectively published under that Code from the electronic edition alone. This published work and the nomenclatural acts it contains have been registered in ZooBank, the online registration system for the ICZN. The ZooBank LSIDs (Life Science Identifiers) can be resolved and the associated information viewed through any standard web browser by appending the LSID to the prefix http://zoobank.org/ . The LSID for this publication is: urn:lsid:zoobank.org:pub:7E91563A-395E-4B52-8AD1-009CFABBD472. The online version of this work is archived and available from the following digital repositories: PeerJ, PubMed Central and CLOCKSS.

Results

Five tardigrade species from three genera were found in the present study: Mil. inceptum (Eutardigrada, Apochela), Macrobiotus basiatus sp. nov. (Eutardigrada, Parachela), and three species of Heterotardigrada, Vir. miraviridis sp. nov., Vir. perviridis, and Vir. viridissimus.

In terms of relative abundance, Vir. viridissimus was the most numerous species, followed by Mil. inceptum; both species were present on all six posts. The other three species were much less numerous with Macrobiotus basiatus sp. nov. found on four of the posts, whereas Vir. perviridis and Vir. miraviridis sp. nov. were present on only two of the posts.

Taxonomic account

Phylum: Tardigrada Doyère, 1840

Class: Heterotardigrada Marcus, 1927

Family: Echiniscidae Thulin, 1928

Genus:Viridiscus Gąsiorek & Michalczyk, 2019, in Gąsiorek et al., 2019

1. Viridiscus miraviridis sp. nov. Genus name: urn:lsid:zoobank.org:act:2FCD58BD-5429-451A-BE2D-D6BA33C1ED31, Species name: urn:lsid:zoobank.org:act:0A9781C0-8F03-4A68-BE35-46DDB066E246. Publication LSID: urn:lsid:zoobank.org:pub:7E91563A-395E-4B52-8AD1-009CFABBD472.

(Table 3, Figs. 2–4)

Table 3:

Measurements and sp values of selected morphological structures of individuals of Viridiscus miraviridis sp. nov. mounted in Hoyer’s medium.

CHARACTER	N	RANGE		MEAN		SD		Holotype
		μ m^a	sp	μ m^a	sp	μ m^a	sp	μ m^a	sp
Body length	30	133–247	–	200	–	29	14	191	–
Scapular plate length	30	30.1–53.3	–	43.8	–	6.2	–	42.7	–
Head appendages lengths
Cirrus internus	30	8.9–17.8	25.4–34.4	13.5	30.7	2.3	2.3	13.5	31.6
Cephalic papilla	29	4.2–7.6	12.4–15.9	6.1	14.0	0.9	0.8	6.1	14.3
Cirrus externus	29	10.0–21.0	30.5–40.6	15.0	34.4	2.5	2.3	14.9	34.9
Clava	30	3.8–7.2	12.6–16.4	5.9	13.6	0.7	0.9	5.8	13.6
Cirrus A	27	31.2–65.4	103.7–126.5	50.9	114.9	7.8	6.2	50.3	117.8
Cirrus A/Body length ratio	27	21%–30%	–	25%	–	2%	–	26%	–
Cirrus int/ext length ratio	29	77%–96%	–	89%	–	4%	–	91%	–
Spine on leg I length	29	1.8–3.5	5.5–7.1	2.8	6.3	0.5	0.4	2.8	6.6
Papilla on leg IV length	23	3.0–4.7	8.3–10.4	4.0	9.1	0.4	0.5	3.9	9.1
Number of teeth on the collar	25	8–11	–	9.4	–	0.7	–	10	–
Notch length	24	11.4–19.3	33.9–39.9	16.1	36.7	2.1	1.8	16.6	38.9
Claw 1 lengths
Branch	24	10.0–17.9	29.9–35.6	14.1	32.3	1.9	1.6	14.6	34.2
Spur length	10	1.1–1.9	2.7–3.6	1.4	3.2	0.3	0.3	1.3	3.0
Spur/branch length ratio	10	9%–11%	–	10%	–	1%	–	0	–
Claw 2 lengths
Branch	24	9.0–15.4	27.8–34.3	13.1	30.7	1.7	1.8	13.2	30.9
Spur length	12	0.9–1.7	2.9 – 4.2	1.5	3.3	0.2	0.4	?	?
Spur/branch length ratio	12	9%–13%	–	11%	–	1%	–	?	–
Claw 3 lengths
Branch	20	9.8–16.6	27.9–33.5	13.4	30.5	1.6	1.6	13.6	31.9
Spur length	9	1.2–1.5	2.5 – 3.8	1.4	3.0	0.1	0.3	1.3	3.0
Spur/branch length ratio	9	9%–12%	–	10%	–	1%	–	0	–
Claw 4 lengths
Branch	25	11.4–19.7	34.3–40.7	16.2	37.1	2.1	1.7	15.1	35.4
Spur length	20	1.2–2.1	3.2 – 4.7	1.7	3.9	0.2	0.4	1.7	4.0
Spur/branch length ratio	20	9%–13%	–	11%	–	1%	–	0	–

DOI: 10.7717/peerj.10251/table-3

Notes:

N: number of specimens/structures measured, RANGE refers to the smallest and the largest structure among all measured specimens
SD: standard deviation
?: trait oriented unsuitably for measurement
sp: ratio of the length of a given structure to the length of the scapular plate

aexcept for ratios, which are presented as percentages, and the number of teeth in dentate collar.

Viridiscus miraviridis sp. nov. — Figure 2: *Viridiscus miraviridis* sp. nov.
(A) Habitus, dorsoventral assembled image, empty arrow indicates the anterior part of the paired plates, empty arrowhead indicates the posterior part of the paired plates (holotype, PCM). (B) Dorsal view (paratype, SEM). (C) Close up of the dorsal cuticle, empty arrow indicates the anterior part of the paired plates with large, dark unconnected granules, empty arrowhead indicates the posterior part of the paired plates with a mesh of irregular dark granules connected by dark lines, filled arrow indicates large, irregular dark granules, filled arrowhead indicates irregular light depressions between a mesh of dark granules, filled indented arrowhead indicates the absence of median plate 3, mp 1 and mp 2 indicate median plate 1 and 2, respectively (paratype, PCM). (D) Close up of the terminal plate, filled arrow indicates large, irregular dark granules, empty indented arrowhead indicates a dense background of fine dots, filled arrowhead indicates irregular light depressions between a mesh of dark granules (paratype, PCM). (E) Close up of the dorsal cuticle, mp 1 and mp 2 indicate median plate 1 and 2, respectively, pp I and pp II indicate paired plates I and II, respectively (paratype, PCM). (F) Terminal plate with two notches (empty arrow), filled indented arrowhead indicates the absence of median plate 3 (paratype, PCM). Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-2

Comparison of the types of dorsal sculpture in the seven Viridiscus species (PCM). — Figure 4: Comparison of the types of dorsal sculpture in the seven *Viridiscus* species (PCM).
*Vir. miraviridis* sp. nov., *Vir. perviridis*, *Vir. viridissimus*, *Vir. rufoviridis*, *Vir. viridis*, *Vir. clavispinosus*, and *Vir. viridianus* are illustrated (green filter enhanced contrast/colour). The characters used in the comparison are: TP, terminal plate. II-PP, second paired plate. II-MP, second median plate. I-PP, first paired plate. SP, scapular plate. CP, cephalic plate. Scale bar = 20 µm. Photo credit: Roberto Guidetti, except *Vir. clavispinosus* by Giovanni Pilato.

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DOI: 10.7717/peerj.10251/fig-4

Type locality: 36°18′N, 82°22′W, 517 m asl; north side of the East Tennessee State University campus, Johnson City, Washington County, TN, USA, moss (Grimmia sp.) on concrete caps of brick fence posts; xerothermic habitat fully exposed to sun and wind.

Etymology: The species name is based on mira (L.) - wonderful, remarkable and viridis (L.) - green, which refers to the beautiful green color of the cuticle.

Type material: Holotype and 49 paratypes: 3 specimens of the 2nd instar; 20 specimens of adults) identified as females; and 27 individuals whose gender was not distinguished due to poor position on the slide or indistinct gonopore (i.e., as small, round circles that could not be definitely associated with a gender). No 1st instar.

Type depositories: The holotype and 9 paratypes are deposited in the National Museum of Natural History, Smithsonian Institution, Washington, DC, USA. (holotype and 4 paratypes, slide number USNM 1622558; 5 paratypes, slide number USNM 1622559). Additional paratypes are deposited in the Nelson Collection (28 slides), East Tennessee State University, Johnson City, TN, USA.; Bertolani Collection (2 slides with 7 specimens), University of Modena and Reggio Emilia, Modena, Italy; and Kaczmarek Collection (10 slides), Adam Mickiewicz University, Poznań, Poland.

Description of the new species

Animals (measurements and statistics in Table 3)

Females. Body (Figs. 2A–2B) green, eyes absent or not visible after mounted in Hoyer’s. Except for the head appendages (internal and external cirri and drop-shaped cephalic papillae), only lateral cirrus A is present (with finger-like clava near the base) (Figs. 2A–2B). Gonopore rosette-shaped, anterior to the anus.

Cuticle on dorsal plates in general appears “two-layered,” constituted by a surface with shallow depressions over a uniform background of small granules (dark dots) (Figs. 2C–2F, 3). Head plate with very light mesh sculpture over a background of fine dots. On the scapular plate, terminal plate, and posterior parts of paired plates, there are large irregular dark granules and shallow depressions (in PCM) (Figs. 2C–2D, filled arrows) over a dense background of fine dots (Fig. 2D, empty indented arrowhead). These granules appear connected by dark lines, forming a lattice-like network interspersed with irregular light depressions (Figs. 2C–2D, filled arrowheads). Meshes are more irregular on the terminal plate (Fig. 2D). Dorsal plates are well developed. Head and scapular plates are not faceted. Paired plates I and II are divided into two parts, with anterior portion having a background of fine dots and large dark granules that are not connected (Figs. 2A, 2C, empty arrow), and the posterior portion with the sculpture as described above (Figs. 2A, 2C, empty arrowhead). Additionally, on lateral sides of the anterior parts of paired plates I and II, a small area without granules is present (but with background of fine dots). Median plates 1 and 2 are divided into two parts with different types of sculpture (Figs. 2C, 2E). Median plate 1 and 2 have cuticular pattern similar to the scapular plate, terminal plate, and posterior parts of paired plates. Median plate 3 is absent; instead there are dark, round granules over a very poorly visible background of fine dots (Figs. 2C, 2F, filled indented arrowhead). Terminal plate has two notches (Fig. 2F, empty arrow). Ventral plates are absent, but the cuticle has a tiny, regular granulation caused by dense cuticular pillars (Fig. 3A).

All legs have a narrow band of very fine granules (Fig. 3B, empty arrows). Dentate collar on legs IV has fine granulation and 8–14 teeth (teeth usually have only a single point and are slightly irregular) (Figs. 2F, 3D). Triangular spine on leg I (Fig. 3C, filled arrow) and finger-like papilla on leg IV are present (Fig. 3D, filled arrowhead). External claws of all legs are smooth, internal have spurs directed downwards; spurs are larger on the claws on the hind legs.

Males and eggs unknown.

Differential diagnosis

Based on the specific morphology of the cuticle on the dorsal plates, the new species differs from all other species in the viridis group (Fig. 3). More specifically Vir. miraviridis sp. nov. differs from:

1. Viridiscus clavispinosus, known only from its type locality in Republic of Cape Verde (Santo Antão Island off West Africa in the Atlantic Ocean) (Fontoura, Pilato & Lisi, 2011), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs slightly raised dark tubercles that are not connected in Vir. clavispinosus); different shape of clava (finger-like clava in the new species vs spine-like clava in Vir. clavispinosus); area between paired plates II and terminal plate is sculptured vs smooth in Vir. clavispinosus; slightly smaller sp index of the clava (12.6–16.4 in the new species vs 16.5–18.1 in Vir. clavispinosus); larger sp index of cirrus A (103.7–126.5 in the new species vs 54.5–66.4 in Vir. clavispinosus); and larger cirrus A/body length ratio (21–30% in the new species vs 10.9–13.4% in Vir. clavispinosus).

2. Viridiscus perviridis, known from Italy and the USA (Ramazzotti, 1959; Kaczmarek, Michalczyk & McInnes, 2016), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs polygonal tubercles not connected in Vir. perviridis); and shorter cirrus A (31.2–65.4 µm in the new species vs 114.0–170.0 µm in Vir. perviridis).

3. Viridiscus rufoviridis, known from Argentina and Brazil (Kaczmarek, Michalczyk & McInnes, 2015), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs large dorsal granules absent in Vir. rufoviridis); entire body green (only caudal region of the body of green colour in Vir. rufoviridis); median plate 3 absent; terminal plate not faceted; and presence of spurs on internal claws.

4. Viridiscus viridianus, known from the Azores Islands (Portugal) and the USA (Pilato, Fontoura & Lisi, 2007), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs polygonal tubercles not connected in Vir. viridianus); and cirrus A longer than scapular plate (cirrus A shorter than scapular plate in Vir. viridianus). Viridiscus viridianus from the Azores Islands was reported to be dark grey in colour (Fontoura, 1985), rather than green as in the species from the USA and the other species of Viridiscus.

5. Viridiscus viridis, with confirmed localities only from the USA (Hawaiian Islands) (Pilato, Fontoura & Lisi, 2008), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs polygonal tubercles not connected in Vir. viridis); longer cirrus A (31.2–65.4 µm in the new species vs 16.3–18.2 µm in Vir. viridis); larger sp index of cirrus A (103.7–126.5 in the new species vs 36.1–43.5 in Vir. viridis); and larger cirrus A/body length ratio (21–30% in the new species vs 7.4–9.6% in Vir. viridis).

6. Viridiscus viridissimus, known from Romania, the USA and Venezuela (Péterfi, 1956; Kaczmarek, Michalczyk & McInnes, 2015; Kaczmarek, Michalczyk & McInnes, 2016), by: different type of dorsal cuticle (in general dorsal granules appear connected by dark lines in PCM, forming a lattice-like network interspersed with irregular light depressions in the new species vs large dorsal granules absent in Vir. viridissimus) and shorter cirrus A (cirrus A/body length ratio is 21–30% (31.2–65.4 µm) in the new species vs 36% (85.0 µm in specimen 234.0 µm long) in Vir. viridissimus).

2. Viridiscus perviridis Ramazzotti, 1959

Type locality: Italy

Material examined: 137 specimens.

Remarks: Viridiscus perviridis has previously been reported from Italy and Tennessee (USA) (Ramazzotti, 1959; Maucci, 1987; McInnes, 1994; Nelson & Bartels, 2007; Kaczmarek, Michalczyk & McInnes, 2016).

No males were identified in this study. Based on the presence or absence of the anus and gonopore, the age structure of the population was as follows: 20, 1st instar; 22, 2nd instar; and 35, adults. For 60 individuals, 2nd instars and adults could not be distinguished.

3. Viridiscus viridissimusPéterfi, 1956

Type locality: Romania

Material examined: 2,165 specimens.

Remarks: Viridiscus viridissimus has previously been reported from Europe (Romania), North America (Alabama, North Carolina, and Tennessee; USA) and South America (Venezuela) (Péterfi, 1956; Word, Folkerts & Mason, 1976; Nelson, 1979; Grigarick, Schuster & Nelson, 1983; Maucci, 1987; Nelson, Kincer & Williams, 1987; Dewel, Dewel & Nelson, 1992; Dewel, Dewel & Roush, 1992; McInnes, 1994; Dewel & Dewel, 1996; Nelson & Adkins, 2001; Dewel & Eibye-Jacobsen, 2006; Kaczmarek, Michalczyk & McInnes, 2015; Kaczmarek, Michalczyk & McInnes, 2016).

Male gonopore (filled arrowhead) of Viridiscus viridissimus. — Figure 5: Male gonopore (filled arrowhead) of *Viridiscus viridissimus*.
Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-5

Males were present, but rare. The ratio of males to females for individuals whose gender could definitely be determined was 0.04 (29 males to 653 females). This is the first report of males in the genus Viridiscus. The male gonopore (Fig. 5) is oval, slightly raised, and located more posterior than the female gonopore, which has the typical rosette-shape.

The number of individuals that could be determined in each age class was as follows: 178, 1st instar; 299, 2nd instar; and 682, adults. Distinction between 2nd instar and adults could not be made for 1006 individuals due to the position of the gonopore (e.g., hidden by a leg) or due to the immaturity of the animal.

Class: Eutardigrada Richters, 1926

Order: Apochela Schuster et al., 1980

Family: Milnesiidae Ramazzotti, 1962

Genus:MilnesiumDoyère, 1840

4. Milnesium inceptumMorek et al., 2019

Type locality: Germany

Material examined: Total specimens: 1,164; sexually dimorphic males, 126.

Remarks: A pseudocryptic species, morphologically extremely similar to Milnesium alpigenum (Ehrenberg, 1853) but clearly different based on molecular data (Morek et al., 2019). This is a first record of this species from the USA. No males of Mil. inceptum were found by Morek et al. (2019) in populations from Bulgaria, Germany or Switzerland, and culturing of isolated virgin females confirmed that the type population is parthenogenetic. However, males were found to appear spontaneously in an otherwise parthenogenetic culture of the Japanese strain (Suzuki, 2008), suggesting that the species is facultatively parthenogenetic with males appearing only occasionally (Morek et al., 2019). Mature males have modified claws only on the first pair of legs, with the secondary (basal) branch of both claws shaped like a robust hook bearing a small spur (Rebecchi & Nelson, 1998).

DNA sequences

A good quality sequence for COI described here is accessible via GenBank accession number MT502117. Since the COI sequence was almost identical [only two nucleotide substitutions and p-distance 0.3% with sequence MK628723.1 (Morek et al., 2019)] to Mil. inceptum, we did not sequence 28S rRNA or ITS-2.

Class: Eutardigrada Richters, 1926

Order: Parachela Schuster et al., 1980

Superfamily: Macrobiotoidea Thulin, 1928

Family: Macrobiotidae Thulin, 1928

Genus:Macrobiotus C.A.S. Schultze, 1834

Macrobiotus basiatussp. nov. Genus name: urn:lsid:zoobank.org:act:42DFC178-D1BB-48EC-81F6-5B6BE3539AF2, Species name: urn:lsid:zoobank.org:act:95167BDA-B668-4D49-BFD2-1B314080F2F5. Publication LSID: urn:lsid:zoobank.org:pub:7E91563A-395E-4B52-8AD1-009CFABBD472.

(Tables 4–5; Figs. 6–10)

Table 4:

Measurements and pt values of selected morphological structures of individuals of Macrobiotus basiatus sp. nov. mounted in Hoyer’s medium.

CHARACTER	N	RANGE		MEAN		SD		Holotype
		µm	pt	µm	pt	µm	pt	µm	pt
Body length	30	276–540	–	430	–	80	–	461	–
Buccalpharyngeal tube
Buccal tube length	30	29.3–52.6	–	44.4	–	5.8	–	47.6	–
Stylet support insertion point	30	22.7–40.7	74.4–78.9	34.5	77.8	4.5	1.1	35.7	75.0
Buccal tube external width	30	4.6–10.4	15.0–21.0	8.1	18.2	1.5	1.8	7.3	15.3
Buccal tube internal width	28	3.5–8.0	11.3–17.4	6.3	14.3	1.2	1.6	5.8	12.2
Ventral lamina length	29	19.6–31.5	55.7–63.7	26.9	60.0	3.0	2.0	27.3	57.4
Placoid lengths
Macroplacoid 1	30	7.3–17.5	24.6–35.0	13.2	29.6	2.6	2.8	13.6	28.6
Macroplacoid 2	30	4.8–11.0	14.2 – 22.7	8.2	18.4	1.7	2.2	8.5	17.9
Microplacoid	30	2.3–5.1	6.6–10.3	3.8	8.6	0.8	1.0	3.8	8.0
Macroplacoid row	30	13.5–30.9	44.4–61.8	23.6	52.9	4.4	4.6	25.0	52.5
Placoid row	28	17.2–37.8	53.6–75.6	28.9	65.3	5.4	5.2	30.2	63.4
Claw 1 lengths
External primary branch	22	7.7–12.4	21.4–26.3	10.8	24.5	1.5	1.2	11.9	25.0
External secondary branch	18	6.0–10.3	16.4–21.4	8.3	18.7	1.2	1.3	9.5	20.0
Internal primary branch	22	7.3–11.5	20.3–24.9	10.1	23.0	1.4	1.2	11.0	23.1
Internal secondary branch	18	5.8–9.1	15.3–18.9	7.8	17.3	1.0	1.0	8.2	17.2
Claw 2 lengths
External primary branch	26	7.9–13.4	22.5–28.3	11.4	26.1	1.8	1.5	13.2	27.7
External secondary branch	24	6.1–10.3	16.2–21.6	8.5	19.6	1.4	1.2	10.3	21.6
Internal primary branch	25	7.3–12.5	20.7–26.7	10.6	24.2	1.7	1.4	11.4	23.9
Internal secondary branch	22	5.4–9.7	16.0–20.5	8.0	18.5	1.3	1.1	9.4	19.7
Claw 3 lengths
External primary branch	27	8.2–13.8	22.7–29.1	11.7	26.2	1.7	1.5	13.0	27.3
External secondary branch	27	6.5–11.1	16.2–23.3	8.9	19.9	1.3	1.5	11.1	23.3
Internal primary branch	25	8.0–13.0	22.8–27.4	10.9	24.5	1.5	1.2	12.5	26.3
Internal secondary branch	21	5.5–10.1	16.3–21.3	8.4	18.8	1.3	1.4	9.8	20.6
Claw 4 lengths
Anterior primary branch	23	9.0–14.9	21.8–31.0	12.1	27.2	1.7	1.8	12.0	25.2
Anterior secondary branch	20	6.5–11.1	17.9–23.4	8.8	19.9	1.3	1.2	8.8	18.5
Posterior primary branch	22	9.1–14.8	24.8–31.6	12.7	28.7	1.7	1.5	13.5	28.4
Posterior secondary branch	21	6.4–11.2	17.9–23.6	9.3	21.1	1.4	1.6	10.4	21.8

DOI: 10.7717/peerj.10251/table-4

Notes:

N: number of specimens/structures measured, RANGE refers to the smallest and the largest structure among all measured specimens
SD: standard deviation
pt: ratio of the length of a given structure to the length of the buccal tube expressed as a percentage

Type locality: 36°18′N, 82°22′W, 517 m asl; north side of East Tennessee State University campus, Johnson City, Washington County, TN, USA, moss (Grimmia sp.) on concrete caps of brick fence posts; xerothermic habitat fully exposed to sun and wind.

Etymology: This species is named for the shape of the projections on the eggs, which resemble the famous chocolate candy, Hershey’s Kisses. From basiatus (L.), which means “kissed”

Type material: Holotype and 234 paratypes (199 specimens, 35 eggs –2 embryonated).

Type depositories: The holotype and 9 paratypes are deposited in the National Museum of Natural History, Smithsonian Institution, Washington, DC, USA (holotype slide number USNM 1622560; paratype slide numbers USNM 1622561, USNM 1622562, USNM 1622563, USNM 1622564, USNM 1622565 (2 specimens), USNM 1622566, USNM 1622567, USNM 1622568). Additional paratypes are deposited in the Nelson Collection (180 slides), East Tennessee State University, Johnson City, TN, USA; Bertolani Collection (12 slides with 20 animals and 2 eggs), University of Modena and Reggio Emilia, Modena, Italy; and Kaczmarek Collection (34 slides), Adam Mickiewicz University, Poznań, Poland.

Table 5:

Measurements of selected morphological structures of eggs of Macrobiotus basiatus sp. nov.^a

CHARACTER	N	RANGE (μ m)	MEAN (μ m)	SD (μ m)
Diameter of egg without processes	8	72.3–88.9	82.2	6.6
Diameter of egg with processes	8	96.0–113.3	104.6	5.9
Process height	25	10.9–17.0	13.8	1.5
Process base width	25	14.2–19.0	16.4	1.2
Process base/height ratio	25	101%–142%	120%	12%
Distance between processes	23	3.9–7.5	5.3	0.9
Number of processes on the egg circumference	8	14–14	14.0	0.0

DOI: 10.7717/peerj.10251/table-5

Notes:

aexcept for ratios, which are presented as percentages, and the number of processes on the egg circumference.

Description of the new species

Animals (measurements and statistics in Table 4)

Body colour white (transparent after fixation in Hoyer’s medium; Figs. 6A–6B). Larger specimens with several orange or brown irregular excretory granules. Eyes present in 97% of measured specimens; eyes were absent only in one specimen. Entire cuticle covered with conspicuous small, round pores (1.0–1.5 in diameter) distributed randomly but more clearly visible on the anterior part of the body (Figs. 6C–6E). Clearly visible granulation present on all legs, but much more indistinct on legs I-III (Figs. 7A–7C). Buccal-pharyngeal apparatus of the Macrobiotus type, with the ventral lamina and ten peribuccal lamellae (Figs. 8A–8B). Mouth antero-ventral. The oral cavity armature of the hufelandi type (Kaczmarek & Michalczyk, 2017), with three bands of teeth visible under PCM (Figs. 8C–8D). The first band of teeth composed of very small granules (PCM), arranged in a few rows and situated on the basal parts of the lamellae and just below them. The second band of teeth composed of small granules (PCM), arranged in a few rows, just above the third band of teeth. The third band of teeth composed of three dorsal and three ventral teeth in the shape of transverse ridges (PCM). Median tooth on ventral side in shape of an oval granule that can sometimes be divided into two parts. Pharyngeal bulb spherical with triangular apophyses, two rod-shaped macroplacoids and a thin microplacoid. Macroplacoid length configuration 2<1. The first macroplacoid with a central constriction (Fig. 8E). The second macroplacoid without constriction, but with latero-terminal globular projections. Claws of the hufelandi type, stout (Figs. 9A–9D). Primary branches with distinct accessory points. Lunules I–III smooth, but IV slightly indented, with very tiny teeth mainly visible with SEM (Figs. 9C insert and 9D). Thin paired bars under claws I–III present. Other cuticular thickenings on legs absent.

Macrobiotus basiatus sp. nov. — Figure 6: *Macrobiotus basiatus* sp. nov.
(A) Habitus, dorsoventral assembled image (holotype, PCM). (B) Animal in dorsal view (paratype, SEM). (C) Cuticle covered with pores (holotype, PCM). (D) Cuticle covered with pores (paratype, SEM). (E) Ring of pores visible posterior to the mouth opening. Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-6

Macrobiotus basiatus sp. nov. granulation on legs (filled arrows). — Figure 7: *Macrobiotus basiatus* sp. nov. granulation on legs (filled arrows).
(A) Leg III (paratype, PCM). (B) Leg IV (paratype, PCM). (C) Leg IV (paratype, SEM). Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-7

Eggs (measurements and statistics in Table 5)

Eggs spherical, ornamented and laid freely (Figs. 10A–10B). Processes conical with short and sometimes a slightly flexible elongated tip (Figs. 10A–10F). In PCM, the surface of the processes and arms of the areoles have a reticular pattern with a regular rounded mesh (Figs. 10C–10D). In SEM, the processes and arms of areoles are smooth and visibly raised above the internal surface of the egg and connected with it by branched columns (Figs. 10E– 10F). Each process is surrounded by 9–10 areoles, smooth inside (Figs. 10D–10H), not always visible in PCM (Fig. 10B). The number of areoles between two neighboring processes is always two (Fig. 10H).

Macrobiotus basiatus sp. nov. claws. — Figure 8: *Macrobiotus basiatus* sp. nov. claws.
(A) Mouth opening with ten peribuccal lamellae, filled arrowhead indicates teeth of the first band, filled arrow indicates teeth of the second band and indented filled arrowhead indicates teeth of the third band (paratype, SEM). (B) Buccalpharyngeal apparatus (dorsoventral assembled image) (paratype, PCM). (C) Oral cavity armature (dorsal view), filled arrowhead indicates teeth of the first band, filled arrow indicates teeth of the second band and indented filled arrowhead indicates teeth of the third band. (D) Oral cavity armature (ventral view), empty arrowhead indicates median tooth of the third band on ventral side in shape of oval granule (paratype, PCM). (E) Macroplacoids, filled arrowhead indicates the first macroplacoid with a central constriction (paratype, PCM). Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-8

Differential diagnosis

The new species, based on the morphology of the eggs and adults, is most similar to Macrobiotus deceptor Meyer et al., 2017, Macrobiotus nelsonae Guidetti, 1998, Macrobiotus pallarii Maucci, 1954, and Macrobiotus ragonesei Binda et al., 2001, but it differs specifically from:

Macrobiotus deceptor, known only from Saint Mary Parish, Antigua (Meyer et al., 2017), by: presence of eyes, cuticular pores, and granulation on legs in the new species, oval areoles around egg processes (polygonal in Mac. deceptor), shorter egg processes (10.9–17.0 µm in the new species vs 21.2–30.3 µm in Mac. deceptor) and smaller number of areoles around egg processes (ca. 8 in the new species vs 10–12 in Mac. deceptor).
Macrobiotus nelsonae, known only from Roan Mountain, Tennessee, USA (Guidetti, 1998), by: the stylet supports situated in more anterior position (pt 74.4–78.9 in new species vs 79.4–84.5 in Mac. nelsonae), shorter egg processes (10.9–17.0 µm in new species vs 20.8–31.7 µm in Mac. nelsonae), oval areoles around egg processes in the new species (polygonal in Mac. nelsonae), narrower egg processes (14.2–19.0 µm in new species vs 20.0–34.7 µm in Mac. nelsonae) and smaller number of areoles around egg processes (ca. 8 in new species vs 11–12 in Mac. nelsonae).
Macrobiotus pallarii, reported from Asia, Europe and North America (McInnes, 1994), by: the presence of slightly indented with very tiny teeth on lunules (visible with SEM) on legs IV (smooth lunules in Macrobiotus pallarii) and presence of two rows of areoles around each egg processes (only one row of areoles in Mac. pallarii). Macrobiotus pallarii, is poorly described and needs revision, but we have included it here for comparison with the new species.
Macrobiotus ragonesei, reported from North Kivu Province, Democratic Republic of the Congo (Binda et al., 2001), by: the presence of granulation on legs I–III, the presence of regular reticular mesh on the entire surface of the egg processes (egg processes with only the basal part covered with regular reticular mesh in Mac. ragonesei), higher number of processes on the egg circumference (14 in the new species vs 10–12 in Mac. ragonesei), smaller eggs (72.3–88.9 µm and 96.0–113.3 µm with and without processes, respectively, in the new species vs 92.0–100.0 µm and 126.0–136.0 µm with and without processes, respectively, in Mac. ragonesei), shorter egg processes (10.9–17.0 µm in the new species vs 20.0–23.0 µm in Mac. ragonesei), and narrower egg processes (14.2–19.0 µm in the new species vs 20.0–26.0 µm in Mac. ragonesei).

DNA sequences

Good quality sequences described here are accessible via GenBank accession numbers MT488397, MT488398, MT488399, MT488400, MT488401, MT488402 for 28S rRNA, MT498094 for 18S rRNA, MT505165, MT505166, MT505167, MT505168, MT505169 and MT505170 for ITS-2, and MT502116 for COI. We obtained two exoskeletons of Mac. basiatus sp. nov. after DNA isolation, which are deposited in the Kaczmarek collection, labelled USA/NEL/4/S and USA/NEL/6/S.

Molecular differential diagnosis

All the obtained DNA barcode sequences of Mac. basiatus sp. nov. were unique and distinct from those deposited in GenBank.

The p-distances between new species and other deposited in GenBank are:

18S rRNA: 0.4–3.6% (1.9% on average), with the most similar being the Mac. hufelandi group (FJ435738 and FJ435739) (Guil & Giribet, 2012) and the least similar being Macrobiotus polypiformis Roszkowska et al., 2017 (KX810008) (Roszkowska et al., 2017).
28S rRNA: 1.8–11.8% (6.6% on average), with the most similar being Mac. hufelandi group (FJ435751 and FJ435754 –FJ435755) (Guil & Giribet, 2012) and the least similar being Mac. polypiformis (KX810009) (Roszkowska et al., 2017).
COI: 15.3–23.4% (19.1% on average), with the most similar being Macrobiotus terminalis Bertolani & Rebecchi, 1993 (AY598775) (Guidetti et al., 2005) and the least similar being Mac. polypiformis (KX810012) (Roszkowska et al., 2017).
ITS-2: 0.5–30.6% (17.2% on average), with the most similar being Macrobiotus canaricus (Stec, Krzywański & Michalczyk, 2018) (MH063929) (Stec, Krzywański & Michalczyk, 2018) and the least similar being Macrobiotus scoticus Stec et al., 2017 (KY797268) (Stec et al., 2017).

Discussion

The most numerous species, Vir. viridissimus, was obviously a well-established species on the fence posts. With the second highest relative abundance, Mil. inceptum was also considered highly successful in this xerothermic habitat. Previously found in samples from this site (Nelson & Adkins, 2001) but undescribed until now, Mac. basiatus sp. nov. was relatively low in abundance.

Macrobiotus basiatus sp. nov. eggs. — Figure 10: *Macrobiotus basiatus* sp. nov. eggs.
(A) Entire egg (SEM). (B) Entire egg (PCM). (C–D) Egg processes, indented arrowheads indicate visible arms of areoles (PCM). (E–F) Egg processes, indented arrowheads indicate branched columns under processes and arms of areoles (SEM). (G) Egg surface with areoles (PCM). (H) Egg surface with areoles (SEM). Scale bars in micrometres [µm]. Photo credit: Milena Roszkowska.

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DOI: 10.7717/peerj.10251/fig-10

The large number of Mil. inceptum individuals is interesting since this species is a predator. If Milnesium individuals fed on Viridiscus or Macrobiotus specimens, then cuticular parts of the animals (e.g., claws, buccal tubes, dorsal plates) should have been visible within the gut, however, no evidence of this was found. Thus, the major food supply for such a large population of this predator is probably constituted by the numerous rotifers and nematodes, verified by the presence of rotifer jaws in guts of some Mil. inceptum specimens.

The genus Viridiscus was recently established by Gąsiorek & Michalczyk in Gąsiorek et al. (2019) for the viridis group of species previously included in the genus Echiniscus. Viridiscus is characterized by species with a fully or partially green cuticle (although sometimes brown, gray, or almost black), a dorsal plate sculpture composed of an intracuticular foam-like layer and small, flat and densely arranged epicuticular granules, no cuticular appendages other than cephalic appendages and lateral cirri A, well-developed sabre-like claws, absence of males, and inhabiting terrestrial ecosystems. The genus includes Vir. clavispinosus, Vir. perviridis, Vir. rufoviridis, Vir. viridis, Vir. viridianus, Vir. viridissimus, and now Vir. miraviridis sp. nov. The dorsal cuticle is characterized by an evident surface (with “pits”, granules, meshes, etc.) characteristic of each species, overlying a quite uniform background of fine dots. In Viridiscus a ventral cuticular pattern is also present, but it is also present in species from different heterotardigrade genera; this character has not yet been evaluated for its taxonomic value in the genus. The presence of a green cuticle, a primary morphological criterion for Viridiscus, is also found in Echiniscus pooensis Rodriguez-Roda, 1948, which has lateral and dorsal spines on the cuticular plates and therefore was not included in the genus in Gąsiorek et al. (2019).

We noted also that the specimens of Vir. rufoviridis of the Maucci collection present a different structure of the dorsal cuticle of the paired plates and second median plate than any other species of Viridiscus, which was also reported by Pilato, Fontoura & Lisi (2008). In fact, the granules present in the anterior position of these plates are not overlying the background of fine dots, but these granules become smaller (around the middle portion of each plate) forming the background of small dots present under the posterior surface of the plates characterized by white “pits.”

Males are very rare in the genera derived from the evolutionary line of the former Echiniscus “arctomys group” according to Gąsiorek et al. (2019). Only 37 years ago, males were considered completely absent in the widespread genus Echiniscus, at that time composed of about 112 species (Ramazzotti & Maucci, 1983), including the former viridis group (now the genus Viridiscus) within the “arctomys group”. The first report of males in Echiniscus species was published by Dastych (1987) and later by others (e.g., Claxton, 1996; Miller, Claxton & Heatwole, 1999; Mitchell & Romano III, 2007; Claxton & Dastych, 2017, who reported males in several species of Echiniscus. Miller, Claxton & Heatwole (1999) hypothesized a Gondwana origin of bisexual Echiniscus species, comprising those of the “arctomys group.” Although Mitchell & Romano III (2007) reported the first males in the former Echiniscus mauccii [now Claxtonia mauccii (Ramazzotti, 1956)] in North America, our finding of males in Vir. miraviridis sp. nov. confirms the presence of males in the genus Viridiscus on the continent. Therefore, the proposal of a Gondwana origin of bisexual species in this evolutionary line needs to be re-evaluated. In the description of the new genus Viridiscus in Gąsiorek et al. (2019), the species in the genus were reported to reproduce by parthenogenesis. However, in our study, males of Vir. viridissimus were rare (0.04%) but present, raising questions about that one aspect of the generic description. Additional observations of gonopores may reveal rare males in other species of Viridiscus.

The distribution of the genus Viridiscus is global, occurring throughout Europe, North and South America and on Pacific and Atlantic islands, leading to the hypothesis that the genus is also present in areas not yet well investigated. Although Vir. viridis has been widely reported from Europe, Hawaii (Oahu Is.) USA, North and South America, at present only the Hawaiian reports are correct according to Pilato, Fontoura & Lisi (2008). A similar wide range (Europe, North and South America) has been reported for Vir. viridissimus, but probably this species also has a narrower distribution. Viridiscus perviridis is reported from Europe and North America, Vir. viridianus from North America and the Azores Islands (Atlantic Ocean), and Vir. rufoviridis only from South America. Viridiscus clavispinosus is currently considered endemic to Santo Antão Island (Republic of Cape Verde, Atlantic Ocean, off West Africa) and Viridiscus miraviridis sp. nov. is known only from Tennessee (USA, North America) (McInnes, 1994; Fontoura, Pilato & Lisi, 2011; Kaczmarek, Michalczyk & McInnes, 2015; Kaczmarek, Michalczyk & McInnes, 2016). The records of the widely distributed species need further confirmation.

Conclusions

Five species of tardigrades, including two species new to science, were found in moss from a xerothermic habitat in Tennessee: Milnesium inceptum (Eutardigrada), Macrobiotus basiatus sp. nov. (Eutardigrada), and three species of Heterotardigrada, Viridiscus. miraviridis sp. nov., Vir. perviridis, and Vir. viridissimus. This is the first record of Mil. inceptum from North America, confirmed by COI analysis, and the first record of males in the genus Viridiscus. The two new species were identified by an integrative analysis of morphological and/or molecular characters. This research increases our knowledge of tardigrades in Tennessee and North America and adds to the database on biogeography and distribution. Further investigations could examine the tardigrade community structure, analysing the co-existence of multiple Viridiscus species in the isolated habitats, as well as determining the chemical composition of the green pigment in the Viridiscus cuticle, as little is known of the chemical composition of various pigments in tardigrades.

Supplemental Information

Sequences and GenBank numbers for 18S rRNA, 28S rRNa, ITS2, and COI for Macrobiotus basiatus n sp. and COI for Milnesium tardigradum

DOI: 10.7717/peerj.10251/supp-4

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Sequences of 18S rRNA for Macrobiotus basiatus n. sp

DOI: 10.7717/peerj.10251/supp-5

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Sequences of COI for Macrobiotus basiatus n. sp

DOI: 10.7717/peerj.10251/supp-6

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Sequences of ITS2 for Macrobiotus basiatus n. sp

DOI: 10.7717/peerj.10251/supp-7

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Sequences of 28S rRNA for Macrobiotus basiatus n. sp

DOI: 10.7717/peerj.10251/supp-8

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[1] Bartels PJ, Mormino CJ, Nelson DR. 2017. Aquatic tardigrades of North Carolina with a new North American record of a lotic tardigrade, Dactylobiotus haplonyx Maucci, 1980 (Eutardigrada: Macrobiotidae) Journal of the North Carolina Academy of Science 132:26-30

[2] Bartels PJ, Nelson DR. 2006. A large-scale, multihabitat inventory of the Phylum Tardigrada in the Great Smoky Mountains National Park, USA: a preliminary report. Hydrobiologia 558:111-118

[3] Bartels PJ, Nelson DR. 2007. An evaluation of species richness estimators for tardigrades of the Great Smoky Mountains National Park, TN and NC, USA. Journal of Limnology 66(Suppl. 1):104-110

[4] Bartels PJ, Nelson DR. 2012. An online key and field guide to the tardigrades of the Great Smoky Mountains National Park (North Carolina & Tennessee, USA, North America) with taxonomic revisions of two species. Zootaxa 3249:67-68

[5] Bartels PJ, Nelson DR, Exline RP. 2011. Allometry and the removal of body size effects in the morphometric analysis of tardigrades. Journal of Zoological Systematics and Evolutionary Research 49:17-25

[6] Bartels PJ, Nelson DR, Kaczmarek Ł, Michalczyk Ł. 2007. Doryphoribius smokiensis, a new species of Eutardigrada (Hypsibiidae) from the Great Smoky Mountains National Park TN, USA (North America) Zootaxa 1646:59-65

[7] Bartels PJ, Nelson DR, Kaczmarek Ł, Michalczyk Ł. 2008. Three new species and one new record of the genus Doryphoribius Pilato, 1969 (Tardigrada: Eutardigrada: Hypsibiidae) from the Great Smoky Mountains National Park (Tennessee, USA) Journal of Natural History 42:2595-2608

[8] Bartels PJ, Nelson DR, Kaczmarek Ł, Michalczyk Ł. 2011. Ramazzottius belubellus, a new species of Tardigrada (Eutardigrada: Parachela: Hypsibiidae) from the Great Smoky Mountains National Park (North Carolina, USA) Proceedings of the Biological Society of Washington 124:23-27

[9] Bartels PJ, Nelson DR, Kaczmarek Ł, Michalczyk Ł. 2014. The genus Milnesium (Tardigrada: Eutardigrada: Milnesiidae) in the Great Smoky Mountains National Park (North Carolina and Tennessee, USA), with the description of Milnesium bohleberi sp. nov. Zootaxa 3826:356-368

[10] Bartels PJ, Pilato G, Lisi O, Nelson DR. 2009. Macrobiotus (Eutardigrada, Macrobiotidae) from the Great Smoky Mountains National Park, Tennessee/North Carolina, USA (North America): two new species and six new records. Zootaxa 2022:45-57

[11] Bertolani R, Grimaldi de Zio S, D’Addabbo Gallo M, Morone De Lucia MR. 1984. Postembryonic development in Heterotardigrades. Monitore Zoologico Italiano 18:307-320

[12] Bertolani R, Guidetti R, Marchioro T, Altiero T, Rebecchi L, Cesari M. 2014. Phylogeny of Eutardigrada: new molecular data and their morphological support lead to the identification of new evolutionary lineages. Molecular Phylogenetics and Evolution 76:110-126

[13] Bertolani R, Marley N, Nelson DR. 1999. Re-description of the genus Thulinia (Eutardigrada, Hypsibiidae) and of Thulinia augusti (Murray, 1907) comb. n. Zoologischer Anzeiger 238:139-145

[14] Bertolani R, Rebecchi L. 1993. A revision of the Macrobiotus hufelandi group (Tardigrada, Macrobiotidae), with some observations on the taxonomic characters of eutardigrades. Zoologica Scripta 22:127-152

[15] Binda MG, Pilato G, Moncada E, Napolitano A. 2001. Some tardigrades from Central Africa with the description of two new species: Macrobiotus ragonesei and M. priviterae (Eutardigrada Macrobiotidae) Tropical Zoology 14:233-242

[16] Casquet J, Thebaud C, Gillespie RG. 2012. Chelex without boiling, a rapid and easy technique to obtain stable amplifiable DNA from small amounts of ethanol-stored spiders. Molecular Ecology Resources 12:136-141

[17] Claxton SK. 1996. Sexual dimorphism in Australian Echiniscus (Tardigrada, Echiniscidae) with descriptions of three new species. Zoological Journal of the Linnean Society 116(1–2):13-33

[18] Claxton SK, Dastych H. 2017. A new bisexual species of Echiniscus C.A.S. Schultze, 1840 (Heterotardigrada: Echiniscidae) from Tasmania, Australia. Entomologie Heute 29:105-119

[19] Dabert M, Witalinski W, Kazmierski A, Olszanowski Z, Dabert J. 2010. Molecular phylogeny of acariform mites (Acari, Arachnida): strong conflict between phylogenetic signal and long-branch attraction artifacts. Molecular Phylogenetics and Evolution 56:222-241

[20] Dastych H. 1987. Two new species of Tardigrada from the Canadian Subarctic with some notes on sexual dimorphism in the family Echiniscidae. Entomologische Mitteilungen aus dem Zoologischen Museum Hamburg 8(129):319-334

[21] Dastych H. 1999. A new species of the genus Mopsechniscus Du-Bois Reymond Marcus, 1944 (Tardigrada) from the Venezuelan Andes. Acta Biologica Benrodis 10:91-101

[22] Degma P, Bertolani R, Guidetti R. 2009–2020. Actual checklist of Tardigrada species. (Version 37: Edition: 07-08-2019)

[23] Degma P, Guidetti R. 2007. Notes to the current checklist of Tardigrada. Zootaxa 1579:41-53

[24] Dewel RA, Dewel WC. 1996. The brain of Echiniscus viridissimus Péterfi 1956 (Heterotardigrada): a key to understanding the phylogenetic position of tardigrades and the evolution of the arthropod head. Zoological Journal of the Linnean Society 116:35-49

[25] Dewel RA, Dewel WC, Nelson DR. 1992. Cephalic sensory structures of the heterotardigrade Echiniscus viridissimus. American Zoologist 32:122

[26] Dewel RA, Dewel WC, Roush BG. 1992. Unusual cuticle-associated organs in the heterotardigrade, Echiniscus viridissimus. Journal of Morphology 212:123-140

[27] Dewel RA, Eibye-Jacobsen J. 2006. The mouth cone and mouth ring of Echiniscus viridissimus Péterfi, 1956 (Heterotardigrada) with comparisons to corresponding structures in other tardigrades. Hydrobiologia 558:41-51

[28] Doyère LMF. 1840. Mémoire sur les Tardigrades I. Annales des Sciences Naturelles, Paris, Series 2 14:269-362

[29] du Bois Reymond Marcus E. 1944. Sobre tardigrados Brasileiros. Comunicaciones zoológicas del Museo de Historia Natural de Montevideo 1:1-19

[30] Ehrenberg CG. 1853. Diagnoses novarum formarum. Verhandlungen der Königlich Preussische Akademie der Wissenschaften zu Berlin 8:526-533

[31] Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology 3(5):294-299

[32] Fontoura P, Pilato G, Lisi O. 2011. Tardigrada from Santo Antão Island (Archipelago of Cape Verde, West Africa) with the description of a new species. Zootaxa 2838:30-40

[33] Fontoura P. 1985. Primeiro contributo para o conhecimento da fauna de tardigrados dos Açores. Arquipélago 4:45-56

[34] Gąsiorek P, Morek W, Stec D, Michalczyk Ł. 2019. Untangling the Echiniscus Gordian knot: paraphyly of the “arctomys” group (Heterotardigrada: Echiniscidae) Cladistics 2019:1-21

[35] Grigarick AA, Schuster RO, Nelson DR. 1983. Heterotardigrada of Venezuela (Tardigrada) Pan-Pacific Entomology 59:64-77

[36] Guidetti R. 1998. Two new species of Macrobiotidae (Tardigrada: Eutardigrada) from the United States of America, and some taxonomic considerations of the genus Murrayon. Proceedings of the Biological Society of Washington 111(3):663-673

[37] Guidetti R, Bertolani R. 2005. Tardigrade taxonomy: an updated check list of the taxa and a list of characters for their identification. Zootaxa 845:1-46

[38] Guidetti R, Bertolani R, Nelson DR. 1999. Ecological and faunistic studies on tardigrades of leaf litter of beech forests. Zoologischer Anzeiger 238:215-233

[39] Guidetti R, Gandolfi A, Rossi V, Bertolani R. 2005. Phylogenetic analysis of Macrobiotidae (Eutardigrada, Parachela): a combined morphological and molecular approach. Zoologica Scripta 34:235-244

[40] Guil N, Giribet G. 2012. A comprehensive molecular phylogeny of tardigrades - adding genes and taxa to a poorly resolved phylum-level phylogeny. Cladistics 28(1):21-49

[41] Hall TA. 1999. BioEdit: a user friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series 41:95-98

[42] Kaczmarek Ł, Cytan J, Zawierucha K, Diduszko D, Michalczyk Ł. 2014. Tardigrades from Peru (South America), with descriptions of three new species of Parachela. Zootaxa 3790(2):357-379

[43] Kaczmarek Ł, Grobys D, Kulpa A, Bartylak T, Kmita H, Kepel M, Kepel A, Roszkowska M. 2019. Two new species of the genus Milnesium Doyère, 1840 (Tardigrada) from Madagascar. Zookeys 884:1-22

[44] Kaczmarek Ł, Michalczyk Ł. 2017. The Macrobiotus hufelandi (Tardigrada) group revisited. Zootaxa 4363:101-123

[45] Kaczmarek Ł, Michalczyk Ł, McInnes SJ. 2014. Annotated zoogeography of non-marine Tardigrada. Part I: Central America. Zootaxa 3763:1-62

[46] Kaczmarek Ł, Michalczyk Ł, McInnes SJ. 2015. Annotated zoogeography of non-marine Tardigrada. Part II: South America. Zootaxa 3923:1-107

[47] Kaczmarek Ł, Michalczyk Ł, McInnes SJ. 2016. Annotated zoogeography of non-marine Tardigrada. Part III: North America and Greenland. Zootaxa 4203:1-249

[48] Kathman RD, Nelson DR. 1987. Population trends in the aquatic tardigrade Pseudobiotus augusti (Marray) Selected Symposia and Monographs, Unione Zoologica Italiana 1:155-168

[49] Kendall-Fite KL, Nelson DR. 1996. Two new species of tardigrades from Short Mountain, Tennessee, U.S.A. Zoological Journal of the Linnean Society 116:205-214

[50] Koid A, Nelson WC, Mraz A, Heidelberg KB. 2012. Comparative analysis of eukaryotic marine microbial assemblages from 18S rRNA gene and gene transcript clone libraries by using different methods of extraction. Applied Environmental Microbiology 78(11):3958-3965

[51] Kumar S, Stecher G, Li M, Knyaz C, Tamura K. 2018. MEGA X: molecular evolutionary genetics analysis across computing platforms. Molecular Biology and Evolution 35(6):1547-1549

[52] Marcus E. 1927. Zur Anatomie und Ökologie mariner Tardigraden. Zoologische Jahrbücher. Abteilung für Systematik, Geographie und Biologie der Tiere 53:487-558

[53] Maucci W. 1954. Tardigradi nuovi della fauna Italiana. Atti della Società Italiana di Scienze Naturali e del Museo Civico di Storia Naturale di Milano 93:576-585

[54] Maucci W. 1987. A contribution to the knowledge of the North American Tardigrada with emphasis on the fauna of Yellowstone National Park (Wyoming) Selected Symposia and Monographs, Unione Zoologica Italiana 1:187-210

[55] McInnes SJ. 1994. Zoogeographic distribution of terrestrial/freshwater tardigrades from current literature. Journal of Natural History 28:257-352

[56] McInnes SJ, Michalczyk Ł, Kaczmarek Ł. 2017. Annotated zoogeography of non-marine Tardigrada, Part IV: Africa. Zootaxa 4284:1-74

[57] Meyer H. 2013. Terrestrial and freshwater Tardigrada of the Americas. Zootaxa 3747:01-71

[58] Meyer HA, Hinton JG, Gladney WL, Klumpp MC. 2017. Water bears from the Caribbean island of Antigua, with the description of a new Macrobiotus species (Tardigrada: Eutardigrada: Macrobiotidae) Caribbean Naturalist 39:1-13

[59] Michalczyk Ł, Kaczmarek Ł. 2003. A description of the new tardigrade Macrobiotus reinhardti (Eutardigrada: Macrobiotidae, harmsworthi group) with some remarks on the oral cavity armature within the genus Macrobiotus Schultze. Zootaxa 331:1-24

[60] Michalczyk Ł, Kaczmarek Ł. 2013. The Tardigrada Register: a comprehensive online data repository for tardigrade taxonomy. Journal of Limnology 72(S1):5-181

[61] Miller WR, Claxton SK, Heatwole HF. 1999. Tardigrades of the Australian Antarctic territories: males in the genus Echiniscus (Tardigrada: Heterotardigrada) Zoologischer Anzeiger 238:303-309

[62] Mironov SV, Dabert J, Dabert M. 2012. A new feather mite species of the genus Proctophyllodes Robin, 1877 (Astigmata: Proctophyllodidae) from the Long–tailed Tit Aegithalos caudatus (Passeriformes: Aegithalidae): morphological description with DNA barcode data. Zootaxa 3253:54-61

[63] Mitchell CR, Romano III FA. 2007. Sexual dimorphism, population dynamics and some aspects of life history of Echiniscus mauccii (Tardigrada; Heterotardigrada) Journal of Limnology 66:126-131

[64] Morek W, Blagden B, Kristensen RM, Michalczyk Ł. 2020. The analysis of inter- and intrapopulation variability of Milnesium eurystomum Maucci, 1991 reveals high genetic divergence and a novel type of ontogenetic variation in the order Apochela. Systematics and Biodiversity 18(5):461-479

[65] Morek W, Suzuki A, Schill RO, Georgiev D, Yankova M, Marley N, Michalczyk Ł. 2019. Redescription of Milnesium alpigenum Ehrenberg, 1853 (Tardigrada: Apochela) and a description of Milnesium inceptum sp. nov., a tardigrade laboratory model. Zootaxa 4586:35-64

[66] Murray J. 1910. Tardigrada. British Antarctic Expedition 1907–1909. Reports on the Scientific Investigations. Vol. 1 Biology (Part V). 83-87

[67] Nelson DR. 1975. Ecological distribution of tardigrades on Roan Mountain, Tennessee-North Carolina. Memorie dell’Istituto Italiano di Idrobiologia 32(Suppl.):225-276

[68] Nelson DR. 1979. First identification and SEM analysis of Echiniscus viridissimus (Phylum Tardigrada) outside its type locality. The Journal of the Tennessee Academy of Science 54:71

[69] Nelson DR, Adkins RG. 2001. Distribution of tardigrades within a moss cushion: do tardigrades migrate in response to changing moisture conditions? Zoologischer Anzeiger 240:493-500

[70] Nelson DR, Bartels PJ. 2007. “Smoky Bears”—Tardigrades of the Great Smoky Mountains National Park. Southeastern Naturalist 6(Special Issue 1):229-238

[71] Nelson DR, Bartels PJ. 2013. Species richness of soil and leaf litter tardigrades in the Great Smoky Mountains National Park (North Carolina/Tennessee, USA) Journal of Limnology 72:144-151

[72] Nelson DR, Bartels PJ, Guil N. 2019. Chapter 7, Tardigrade ecology. In: Schill RO, ed. Water bears: the biology of tardigrades. Zoological monographs. Zurich, Switzerland: Springer Nature. Vol. 2:163-210

[73] Nelson DR, Guidetti R, Rebecchi L. 2015. Phylum Tardigrada. In: Thorp J, Rogers DC, eds. Ecology and general biology: Thorp and Covich’s freshwater invertebrates. Oxford, UK: Elsevier Inc.. 347-380

[74] Nelson DR, Kincer CJ, Williams TC. 1987. Populations trends in the aquatic tardigrade Pseudobiotus augusti (Murray) Selected Symposia and Monographs, Unione Zoologica Italiana 1:141-153

[75] Nelson DR, Marley NJ, Bertolani R. 1999. Re-description of the genus Pseudobiotus (Eutardigrada, Hypsibiidae) and of the new type species Pseudobiotus kathmanae sp. n. Zoologischer Anzeiger 238:311-317

[76] Nelson DR, McGlothlin KL. 1993. A new species of Hypsibius (Phylum Tardigrada) from Roan Mountain, Tennessee, U.S.A. Transactions of the American Microscopical Society 112(2):140-144

[77] Nelson DR, McGlothlin KL. 1996. A new species of Calohypsibius (Phylum Tardigrada, Eutardigrada) from Roan Mountain, Tennessee-North Carolina, U.S.A. Zoological Journal of the Linnean Society 116:167-174

[78] Perry E, Miller WR, Kaczmarek Ł. 2019. Recommended abbreviations for the names of genera of the phylum Tardigrada. Zootaxa 4608:145-154

[79] Péterfi F. 1956. Contributioni la Cunoasterea Tardigradelor din R. P. R. Studii si Ceretari di Biolog 7:149-155

[80] Pilato G. 1981. Analisi di nuovi caratteri nello studio degli Eutardigradi. Animalia 8(1/3):51-57

[81] Pilato G, Binda MG. 2010. Definition of families, subfamilies, genera and subgenera of the Eutardigrada, and keys to their identification. Zootaxa 2404:1-52

[82] Pilato G, Fontoura P, Lisi O. 2007. Remarks on the Echiniscus viridis group, with the description of a new species (Tardigrada, Echiniscoidea) Journal of Limnology 66(S1):33-39

[83] Pilato G, Fontoura P, Lisi O. 2008. New description of Echiniscus viridis Murray 1910 and remarks on the viridis group. New Zealand Journal of Zoology 35:85-92

[84] Ramazzotti G. 1956. Tre nouve specie di Tardigradi ed altre specie poco comuni. Atti della Societa Italiana di Scienze Naturali e del Museo Civico di Storia Naturale di Milano 95:284-291

[85] Ramazzotti G. 1959. Il gruppo dell’ Echiniscus viridis con la nuova species E. perviridis e Macrobiotus pustulatus altra nuova specie (Tardigrada) Atti della Società Italiana di Scienze Naturali e del Museo Civico di Storia Naturale di Milano 98:303-309

[86] Ramazzotti G. 1962. Tardigradi del Cile, con descrizione di quattro nuove specie e di una nuova varietà. Atti della Societa Italiana di Scienze Naturali e del Museo Civico di Storia Naturale di Milano 101:275-287

[87] Ramazzotti G, Maucci W. 1983. Il Phylum Tardigrada. Memorie delli Istituto Italiano di Idrobiologia 41:1-1012

[88] Rebecchi L, Nelson DR. 1998. Evaluation of a secondary sex character in eutardigrades. Invertebrate Biology 117(3):194-198

[89] Richters F. 1926. Tardigrada. In: Kükenthalund W, Krumbach T, eds. Handbuch der Zoologie, vol. 3. Berlin: Walter de Gruyter & Co.. 1-68

[90] Riggin GT. 1962. Tardigrada of Southwest Virginia: with the addition of a description of a new marine species from Florida. Virginia Agricultural Experiment Station Technical Bulletin 152:1-145

[91] Roszkowska M, Ostrowska M, Stec D, Janko K, Kaczmarek Ł. 2017. Macrobiotus polypiformis sp. nov., a new tardigrade (Macrobiotidae; hufelandi group) from the Ecuadorian Pacific coast, with remarks on the claw abnormalities in eutardigrades. European Journal of Taxonomy 327:1-19

[92] Roszkowska M, Stec D, Gawlak M, Kaczmarek Ł. 2018. An integrative description of a new tardigrade species Mesobiotus romani sp. nov. (Macrobiotidae: harmsworthi group) from the Ecuadorian Pacific coast. Zootaxa 4450(5):550-564

[93] Schultze CAS. 1834. Macrobiotus Hufelandii animal e crustaceorum classe novum, reviviscendi post diuturnam asphixiam et aridiatem potens. Berolini Apud Carolum Curths 1834:1-8

[94] Schuster RO, Nelson DR, Grigarick AA, Christenberry D. 1980. Systematic criteria of Eutardigrada. Transactions of the American Microscopical Society 99:284-303

[95] Stec D, Kristensen RM, Michalczyk Ł. 2018. Integrative taxonomy identifies Macrobiotus papei, a new tardigrade species of the Macrobiotus hufelandi complex (Eutardigrada: Macrobiotidae) from the Udzungwa Mountains National Park (Tanzania) Zootaxa 4446(2):273-291

[96] Stec D, Krzywański Ł, Michalczyk Ł. 2018. Integrative description of Macrobiotus canaricus sp, nov. with notes on M. recens (Eutardigrada: Macrobiotidae) European Journal of Taxonomy 452:1-36

[97] Stec D, Morek W, Gąsiorek P, Blagden B, Michalczyk Ł. 2017. Description of Macrobiotus scoticus sp. nov. (Tardigrada: Macrobiotidae: hufelandi group) from Scotland by means of integrative taxonomy. Annales Zoologici 67(2):181-197