Next Article in Journal
Heterochlamydomonas uralensis sp. nov. (Chlorophyta, Chlamydomonadaceae), New Species Described from the Mountain Tundra Community in the Subpolar Urals (Russia)
Next Article in Special Issue
A New Species of Vampirolepis (Cestoda: Cyclophyllidea: Hymenolepididae) from the Bat Artibeus lituratus (Chiroptera: Phyllostomidae) in the Amazon Rainforest, Brazil
Previous Article in Journal
Defining Evolutionary Conservation Units in the Macedonian Crested Newt, Triturus macedonicus (Amphibia; Salamandridae), in a Biodiversity Hotspot
Previous Article in Special Issue
Trematodes of Land Birds from the Republic of Mordovia with a Checklist of Avian Trematodes of the Middle Volga Region (European Russia)
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Diversity
Volume 15
Issue 5
10.3390/d15050672
diversity-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Diversity and Distribution of Helminths in Wild Ruminants of the Russian Arctic: Reindeer (Rangifer tarandus), Muskoxen (Ovibos moschatus), and Snow Sheep (Ovis nivicola)

by
Olga A. Loginova
1,*,
Sofya B. Rozenfeld
1,
Taras P. Sipko
1,
Ivan A. Mizin
2,
Danila V. Panchenko
3,
Kasim A. Laishev
4,
Mikhail G. Bondar
5,
Leonid A. Kolpashchikov
5,
Aleksandr R. Gruzdev
6,
Pavel S. Kulemeev
6,
Dennis I. Litovka
7,
Mariia N. Semerikova
7,
Viktor N. Mamontov
8,
Evgeniy G. Mamaev
9 and
Sergei E. Spiridonov
1
1
A. N. Severtsov Institute of Ecology and Evolution of the Russian Academy of Sciences, 119071 Moscow, Russia
2
National Park “Russian Arctic”, 163000 Arkhangelsk, Russia
3
Institute of Biology of Karelian Research Centre of the Russian Academy of Sciences, 185910 Petrozavodsk, Russia
4
St. Petersburg Federal Research Center of the Russian Academy of Sciences, 199178 St. Petersburg, Russia
5
Joint Directorate of Taimyr Nature Reserves, 663305 Norilsk, Russia
6
Wrangel Island State Nature Reserve, 689400 Pevek, Russia
7
Chukotka Arctic Scientific Center, 689000 Anadyr, Russia
8
N. Laverov Federal Center for Integrated Arctic Research of the Ural Branch of the Russian Academy of Sciences, 163020 Arkhangelsk, Russia
9
Commander Islands Nature and Biosphere Reserve, 684500 Nikolskoye, Russia
*
Author to whom correspondence should be addressed.
Diversity 2023, 15(5), 672; https://doi.org/10.3390/d15050672
Submission received: 10 April 2023 / Revised: 12 May 2023 / Accepted: 15 May 2023 / Published: 16 May 2023
(This article belongs to the Special Issue Biodiversity of Parasites in Vertebrates in the Wildlife)
Browse Figures
Review Reports Versions Notes

Abstract

:
The Russian Arctic supports wild sympatric ruminants and their data-deficient helminths. In this study, we: (1) collected fecal samples of wild and semiwild reindeer (Rangifer tarandus), muskoxen (Ovibos moschatus), and snow sheep (Ovis nivicola) across Palearctic North territories: Arkhangelsk Oblast (including Novaya Zemlya archipelago), Karelia and Sakha Republics, Kola, Yamal, Taimyr, and Chukotka Peninsulas, Bering, Svalbard, and Wrangel Islands; (2) conducted a coprological survey (noninvasive life-time method preferable for protected animals) to obtain eggs and larvae of helminths inhabiting digestive, respiratory, nervous, and muscular systems; (3) identified helminths according to their morphology and DNA sequences; (4) estimated parasite load per host; (5) analyzed our findings. Varestrongylus eleguneniensis (in reindeer) was reported for the Palearctic for the first time, while Orthostrongylus sp. was reported both for R. tarandus and for the Palearctic for the first time. Capillarid-type eggs were reported for snow sheep for the first time. The question of the role of wild Arctic ruminants as vectors for rotifers was raised.

Graphical Abstract

1. Introduction

The relatively low biodiversity of terrestrial animal taxa is a hallmark of Polar regions [1,2], but this pattern is not obvious for the parasites of Arctic ruminants [3]. Possibly, there are geological and biological factors that influence the diversity of ruminant parasites in the Northern Palearctic. For example, the continuity of the tundra throughout the area allows animals to move freely: mountain ranges are rare and low, and large rivers are covered with ice for most of the year. In the Pleistocene, the ice sheet was located only in the west of the Palearctic [4,5], and the rest of the territory was continuously inhabited by ruminants. Human activities (including the work of herdsmen, hunters, biologists, and veterinarians) can also have an impact on parasite diversity and distribution. The presence of only three main ruminant species, reindeer (Rangifer tarandus) [6,7,8,9,10], muskox (Ovibos moschatus) [11,12], and snow sheep (Ovis nivicola) [13,14], facilitates the analysis of possible helminth exchange in the northern Palearctic. Reindeer occupy this territory fairly evenly in contrast to the more limited distribution of snow sheep. Muskoxen became extinct in the Palearctic in the Late Pleistocene (12,000 years ago) and have only recently been reintroduced from the Nearctic (1970s) [15,16,17], and their range is still expanding. The aim of this study is to obtain reference data for further monitoring, as this Arctic area is now undergoing active transformation due to climate warming, use of mineral resources, development of transport logistics, etc. Previous helminthological studies were carried out more than 60 years ago [18] for reindeer (mostly domestic) [19] and snow sheep [13,14], and, to the authors’ knowledge, never for muskoxen in Russia. Moreover, these studies never included DNA analysis (because it was not available at the time), and their results were mostly published in Russian journals and conference proceedings, including so-called gray literature [20]. We aim to fill this gap and provide the English-speaking reader the current state of the helminth fauna of selected wild ruminants in the Russian Arctic.

2. Materials and Methods

2.1. Helminth Recovery

Fecal sampling was done for three host species, reindeer (R. tarandus), muskoxen (O. moschatus), and snow sheep (O. nivicola), inhabiting the Arctic area of Russia (Figure 1, Table 1). Territory was defined according to the Conservation of Arctic Flora and Fauna (CAFF) boundary and the Russian Arctic zone boundary. Additionally, we included a site at Svalbard (Norway) due to the Svalbard Treaty recognizing right of Russia (along with many other countries) to engage in commercial activity on the islands of the Archipelago of Spitsbergen, and the actual use of this right.
Semiwild reindeer were also included in this study as representatives of R. tarandus. Being formally domestic animals along with cattle, sheep, and goats, they are not handled like other farm ruminants. The reindeer in question led their life as close to the natural one as it was possible. They were also migrating long distances (with their herder) and were exposed to many natural threats (weather, biting insects, and so on). Wild and semiwild reindeer are often sympatric.
Feces were collected from the ground shortly after defecation. They were stored chilled, frozen, or dried. Prior to analyses, the frozen material thawed out naturally, while the dried material had been humidified for at least 2 h. A total of 519 fecal samples were examined for the presence of helminths (at the stage of egg or larva) via complex coprological survey according to protocols of the National Standard of the Russian Federation (GOST R 54627-2011) Ruminant animals—Methods of Laboratory Helminthological Diagnostics [21]. Qualitative methods consisted of larvoscopy (Vajda’s method), ovoscopy (flotation with Darling’s solution and sedimentation in tap water), and coproculture (for nematode larvae, if possible). Briefly, Vajda’s method prescribes to place 3–4 fecal pellets on a microscope slide, then to add around 1 mL of 40 °C tap water to wash out the pellets, to leave it for 30 min, to remove the pellets, and study the remaining liquid. Flotation with Darling’s solution requires double centrifugation and the usage of the 1:1 mixture of glycerin and saturated sodium chloride solution, followed by supernatant microscopy. Sedimentation and coproculture were standard [22].
The quantitative method required the VIGIS chamber (analogue of the McMaster device) of the Diapar kit (VIGIS, Moscow, Russia).
Morphology of eggs and larvae obtained from feces was studied via light microscopy (LM) with the optical microscope Micmed-6 (LOMO-MA, St. Petersburg, Russia) equipped with lenses of 4× (to navigate the slide), 10×, 20×, 40×, and 100× magnifications (the latter with oil immersion). Micrographs were taken with the digital photo camera 5D Mark II (Canon, Tokyo, Japan) connected to the microscope with the C-mount adapter (LOMO-MA, Russia). Morphometry was performed based on the obtained pictures via Figi/ImageJ Version 1.2.4 RRID:SCR_003070 software (National Institutes of Health, Bethesda, MD, USA). The program was set using the microscope calibration slide (transmitted light object micrometer) OMP (LOMO-MA, Russia). We used Straight Line mode to measure eggs, and Segmented Line mode for larvae.

2.2. DNA Analysis

The DNA was extracted individually from L1 or L3 by digestion with Proteinase K [23]. For species identification, a PCR was performed. To obtain a partial sequence of the internal transcribed spacer (ITS rDNA), we used the forward primer 18S (TTG ATT AGG TCC CTG CCC TTT) and the reverse primer 26S (TTT CAC TCG CCG TTA CTA AGG) [24]. The amplification conditions used were: an initial 5 min denaturation at 94 °C, followed by 35 cycles of denaturation at 94 °C for 30 s, annealing at 55 °C for 60 s, and extension at 72 °C for 60 s, with a final elongation step at 72 °C for 5 min.
The ITS2 region of the internal transcriber spacer was amplified using the forward primer NC1 (ACG TCT GGT TCA GGG TTG TT) and the reverse primer NC2 (ATG CTT AAG TTC AGC GGG T) designed by Gasser et al. [25]. The amplification conditions used were: an initial 3 min denaturation at 95 °C, followed by 35 cycles of denaturation at 95 °C for 30 s, annealing at 54 °C for 45 s, and extension at 72 °C for 60 s, with a final elongation step at 72 °C for 5 min.
For the partial sequence of the D2 and D3 domains of LSU rDNA (28S rDNA), we used the forward primer LSU391 (AGC GGA GGA AAA GAA ACT AA) [26] and the reverse primer LSU501 (TCG GAA GGA ACC AGC TAC TA) [27]. The amplification conditions used were: an initial 3 min denaturation at 95 °C, followed by 35 cycles of denaturation at 95 °C for 30 s, annealing at 52 °C for 35 s, and extension at 72 °C for 60 s, with a final elongation step at 72 °C for 5 min.
Targeting the mitochondrial cytochrome c oxidase subunit I (COX1 mtDNA), we used the forward primer COI_F1 (CCT ACT ATG ATT GGT GGT TTT GGT AAT TG) and the reverse primer COI_R2 (GTA GCA GAC GTA AAA TAA GCA CG) [28]. The amplification conditions used were: an initial 5 min denaturation at 95 °C, followed by 35 cycles of denaturation at 95 °C for 45 s, annealing at 52 °C for 50 s, and extension at 72 °C for 60 s, with a final elongation step at 72 °C for 5 min.
For each run, the final elongation phase was followed by a cooling to 12 °C. Reagent-only (no DNA) reactions were used as negative controls to detect potential contamination.
The results of the PCR were visualized in a 1% agarose gel. Bands of expected molecular weight were excised, and DNA was extracted using the Wizard SV Gel and PCR Clean-Up System (Promega, Madison, WI, USA). After cleaning the PCR product with an ethanol precipitation in the presence of ammonium acetate, the samples were directly sequenced by Genotech® (Moscow, Russia). Obtained chromatograms were analyzed using Chromas 2.6.6. RRID:SCR_000598 (Technelysium Pty Ltd., Brisbane, Australia). The sequences similar to those of the studied material were searched for in GenBank using BLASTN 2.13.0+ [29,30].

3. Results

Eggs of trematodes, cestodes, and nematodes, as well as the first-stage larvae (L1) of nematodes, were found in reindeer feces. Third-stage larvae (L3) were obtained via coproculture in some cases. Parasites were preliminarily identified based on their morphology and morphometric data. Where possible, DNA-based identification was performed (Table 2).
Diagnostic stages of helminths obtained from reindeer feces are shown in Figure 2.
Eggs of cestodes and nematodes, as well as first-stage larvae (L1) of nematodes, were found in the feces of muskoxen and snow sheep. Parasites were preliminarily identified based on their morphology and morphometric data. Diagnostic stages of these helminths are shown in Figure 3 and Figure 4, respectively.
The diversity and distribution of helminths found in reindeer, muskoxen, and snow sheep via coproscopy are summarized in Table 3.
Except for one case, all the samples showed low parasitic intensity (in terms of GOST R 54627-2011). We found 1–10 eggs of trematodes per 1 g of feces, and 1–100 eggs/larvae of cestodes/nematodes per 1 g of feces. Only one sample from semiwild reindeer (#12) contained 45 eggs of Paramphistomum sp. per 1 g. The latter is interpreted as medium intensity (GOST R 54627-2011).

4. Discussion

The following trematodes were reported for R. tarandus: Cotylophoron skrjabini, Dicrocoelium sp., Fasciola hepatica, Fascioloides magna, Fischoederius elongatus, Liorchis sp., and Paramphistomum sp. [14,19,32,33] (C. skrjabini and Liorchis sp. were subsequently placed in synonymy with Paramphistomum sp. [34]). All of them can be detected via coproscopy. The eggs of rumen flukes F. elongatus and Paramphistomum sp. are highly similar. Their size ranges overlap: length and width of F. elongatus (125–135 × 65–70 μm) fit in with the criteria for P. cervi (114–176 × 60–90 μm) [35]. However, F. elongatus was reported only once for reindeer in Buryatia (Russia) [36], of which the southern boundary is 57.09183 N. Thus, we assumed that all typical-looking eggs found in wild and semiwild reindeer feces belonged to Paramphistomum sp. (we aim to investigate them genetically in the future). Feces were collected at different times of day, which could have affected the egg count, as a rise of egg excretion was reported for the afternoon time [37]. D. dendriticum (syn. D. lanceatum) and D. chinensis (cyn. D. orientalis) were both reported for reindeer. The dimensions of their eggs were also overlapping and host-species (host-size)-dependent [38]. Therefore, egg morphology and morphometrics allows us only genus identification. Among the cestodes available for coproscopy in reindeer, there are: Avitellina arctica, Moniezia baeri, M. benedeni, M. expansa, M. mizkewitschi, M. rangiferina, M. taymirica, Thysaniezia giardi, and possibly Thysanosoma sp. [14,19,32,33]. Only the eggs of Moniezia were detected in our study. Given that the Moniezia taxonomy requires thorough investigation (due to the recovery of cryptic species), prior to further DNA analysis, nothing can be added to our identification [39]. Nematodes are the most widespread helminths in reindeer, and the majority of them can be revealed through feces examination. Those include: gastrointestinal nematodes (GINs) (fam. Cooperiidae, Haemonchidae, Molineidae, and Trichostrongylidae), lungworms of the genus Dictyocaulus, representatives of Protostrongylidae, the pinworm Skrjabinema tarandi, the whipworm Trichuris sp., a few species of Capillaria, and Ascaris mosgovoyi [14,19,32,33]. GINs include Cooperia, Haemonchus, Ostertagia, Trichostrongylus, and other genera. The differential traits of their eggs are subtle and lack certain genera. Therefore, eggs of this type are called Strongyle-type eggs [22]. They were the most numerous in this study. In one case, we managed to get L3 and perform its DNA analysis. It resulted in O. arctica (cyn. O. gruehneri) detection in reindeer from the Bering Island (sample set #13). Marshallagia sp. eggs were recovered only in Svalbard (Norway) reindeer, which is consistent with scientific literature data reporting no findings of Marshallagia sp. in Russian reindeer [14,19,32,33]. Alternatively, the absence of Marshallagia in Russian reindeer feces can be due to its specific habitat requirements (high Arctic islands with low rainfalls) combined with seasonal egg production (highest during winter) [32,33,40]. Thus, it is desirable to sample feces from the most northern territories in the winter time, which is extremely difficult. For Svalbard reindeer, M. marshalli was reported [32,33,40]. Egg dimensions for this species were estimated as follows: 180–187 × 80–100 μm [41]. Meanwhile, in this study, the recovered Marshallagia egg (the only one found) was significantly bigger: 217 × 91 μm. Specific egg and L1 traits [42] support genus identification, but DNA analysis is highly desirable for this polymorphic helminth [43,44]. Reported Nematodirus species in reindeer included mostly these two: N. skrjabini and N. tarandi [19,32,33], and their synonymy is suggested. In this study, we discovered Nematodirus eggs of at least three morphological types; hence, further investigation of Nematodirus diversity in reindeer is needed and DNA analysis is essential. Nematodirella longispiculata and N. longissimespiculata are known for R. tarandus [19,32,33]. We identified our findings based on their morphology only up to the genus [45,46]. Three species of Capillaria were reported for reindeer: C. brevipes, C. rangiferi, and Capillaria sp., found by Kadenazii in 1963 [19]. The egg size ranges of these species overlap and their validity is questionable. Therefore, all the capillarid-type eggs in the reindeer were identified as Capillaria sp. Numerous findings of Capillaria, GINs (including Nematodirus and Nematodirella), and Moniezia in semiwild reindeer in contrast to the wild ones have brought up a suggestion that the physiology and tolerance of semiwild reindeer against some parasites may be genetically constrained (due to boreal conditions and the tundra-type origin of domestic animals) [33,47,48,49,50]. In fact, we have also found Capillaria, Moniezia, and Nematodirella only in semidomestic reindeer (Table 3). Yet, the interpretation of this fact requires accuracy to avoid survivorship bias [51]. Capillaria, GINs (including Nematodirus and Nematodirella), and Moniezia are more often found in calves than adults [19,33,49]. Fecal sampling of domestic reindeer involves many calves due to the specificity of this industry. Animals are most available during preslaughter examination. The most promising (healthy) reindeer are supposed to stay in the main herd (which can consist of a thousand animals), whereas “excessive” yearlings and weak old reindeer are supposed to be killed for meat and other products. Thus, more infected animals are fecal-sampled. In wild reindeer, the situation is usually quite reverse. Herds are also divided throughout the year (naturally) so that males are separate from females with calves. It is mostly small groups of males that are fecal-sampled alive, whereas the most weak and infected animals die themselves or become the prey of predators out of sight of researchers, and thus escape inclusion in the studies. Only sample set #1 was obtained from a group of wild reindeer including calves, and it also showed a higher parasitic load. Seasonal egg production of different parasites, as well as the sampling season, should be considered while interpreting the results [19,33]. The same holds for Dictyocaulus L1s that we found only in semiwild reindeer (including feralized animals from the Bering Island). DNA analyses are required to clarify the species (D. murmanensis and D. eckerti were reported [19]), especially in light of the recent discovery of new species of this genus in cervides [52,53]. Among the fam. Protostrongylidae, only Elaphostrongylus rangiferi was reported previously for reindeer in Russia [19]. We have found its DSL in semiwild reindeer (sample set #10 and 12) and supported our findings with DNA analyses (Table 2). Moreover, DSL were found in six other sample sets (both from wild and semiwild animals). Surprisingly, our attempt to confirm E. rangiferi for L1 from sample set #1 resulted in identifying Varestrongylus eleguneniensis—another (newly described) protostrongylid reported for R. tarandus in the Nearctic [54,55]. Even more surprising was a finding of non-DSL L1 of Protostrongylidae in the same sample set. DNA analysis resulted in Orthostrongylus sp. identification (Table 2). To date, the only species known from this genus is O. macrotis [56]. Orthostrongylus sp. has never been reported in R. tarandus before, nor has it been reported in the Palearctic [31,57]. Trichuris baskakowi, T. globulosa, T. longispiculus, T. massino, T. ovis, and T. tarandi were reported for reindeer [19]. Their egg ranges overlap, and the validity is questionable. Therefore, we identified our findings (both in wild and semiwild reindeer) as Trichuris sp. These worms were also found more frequently in young animals [33], which corresponds at least to sample set #10 (where we know that calves were present). Skrjabinema tarandi is a serendipitous finding during coproscopy due to the specific life cycle of this parasite [19,33]. More reliable methods include examinations of perianal deposits or cellophane (Scotch) tape tests [58]. However, it is still found from time to time [33,59]. In this study, S. tarandi was found only twice in semiwild reindeer (sample set #8 and #16) [60]. Ascaris mosgovoyi is an Ascaris-type helminth described for reindeer in 1959 [19]. It was described only via males; no pictures or descriptions of its eggs are available. That might be a reason why scientists were not reporting it (or looking for it) later on. However, an egg shown in Figure 2K is, in our opinion, A. mosgovoyi due to its specific traits (including larvae), size, and host. Further investigation and DNA analysis are highly desirable.
No trematodes were found in the muskoxen feces. Known cestodes reported for them and available for coprology include: M. expansa and Moniezia sp. [32]. M. expansa eggs tend to be triangular in shape in slides; therefore, we identify our findings as Moniezia sp., since they looked squarer. GINs, found in the muskoxen feces, were Strongyle-type nematodes, Nematodirus sp., and Nematodirella sp., which corresponds to the literature data. In particular, Teladorsagia boreoarcticus (Strongyle-type), Nematodirella alcidis, N. gazelli, N. longissimespiculata, as well as Nematodirus helvetianus and N. tarandi, were reported for muskoxen in North America [32]. The following protostrongylids were reported for muskoxen: Protostrongylus stilesi, Umingmakstrongylus pallikuukensis, and V. eleguneniensis [32]. The latter two species produce DSL. Given that all Russian muskoxen are offspring of animals (re-)introduced from North America back in the XX century [15,16,17,18], we expect that the DSL found might belong to one (or both) of these species. Taking into account V. eleguneniensis found in reindeer, we suggest that introduced animals could have been the source of Nearctic protostrongylids. Further thoughtful investigation (including DNA analysis and animal travel history) is needed. Trichuris spp. were found at a low prevalence both on the North American mainland (yet no Banks Islands) [32] and in this study.
Regarding snow sheep, the following helminths were reported for them: flatworm Stilesia sp., tapeworms Moniezia benedeni, M. expansa, M. rangiferina, M. taymirica, and Moniezia sp., GINs (including Marshallagia mongolica, Nematodurus oiratianus, Nematodirus sp., Nematodirella longissimespiculata, and Nematodirella sp., as well as Ostertagia arctica Ostertagia sp. and Teladorsagia circumcincta), protostrongylids (Protostrongylus sp. and Spiculocaulus leuckarti), Skrjabinema ovis, S. chubuki, Truchuris skrjabini, T. ovis, and Trichuris sp. [13,14]. We found Moniezia sp., GINs (including Marshallagia sp., Nematodurus sp., and Strongyle-type), protostrongylids (Protostrongylus sp.), Truchuris sp., and the Capillarid-type helminth (Figure 4). Notably, the Protostrongylus L1s were of two morphological types: the ones with a longer tail spike (from the sample set #23) and the ones with a twice-shorter tail spike (sample set #24). Marshallagia sp. eggs found were also (as in Svalbard reindeer) bigger than expected. Further investigation is needed.
Ranges of snow sheep in this study were independent and separated from each other and other northern ungulates. Therefore, their helminth fauna probably suggests certain autonomy. Wild reindeer from Taymyr (sample set #1) might have been sympatric to introduced muskoxen (#20, 21). Yamal muskoxen (#18, 22) might be sympatric to Yamal semiwild reindeer (these were not studied here). Muskoxen on the Wrangel Island (#19) were sympatric with introduced and ferallized semiwild reindeer. However, by 2023, those reindeer herds died out. Wild reindeer from the Murmansk Oblast (sample set #5) were not supposed to be in sympatry with domestic herds. Information regarding other populations and their sympatry with domestic and wild ungulates is fragmentary and barely reliable.
Apart from the true parasites (helminths) of the studied wild ruminants, spurious and pseudoparasites were also found. Among them were Bdelloid rotifers at their different stages (Figure A1). In dried fecal samples, they revived after adding water [61]. They could have contaminated the feces after excretion (since material was often picked up from the ground). Alternatively, rotifers were accidentally ingested and managed to survive inside the ruminants. To the authors’ knowledge, Arctic ruminants have never been considered as vectors for rotifers. Given that reindeer cover long distances daily, they might play a major role in the rotifers’ distribution. We welcome specialists on rotifers to collaborate.

5. Conclusions

The diversity of helminths in wild and semiwild reindeer, muskoxen, and snow sheep was studied using a noninvasive method (coproscopy). In R. tarandus, two genera of trematodes (Paramphistomum and Dicrocoelium) were found, as well as one genus of cestodes (Moniezia), and various nematodes: small gastrointestinal nematodes, Marshallagia sp., Nematodirus spp. (three different morphological types), Nematodirella sp. (including N. longissimespiculata), Skrjabinema tarandi, Trichuris sp., Capillaria spp., (possibly) Ascaris mosgovoyi, Dictyocaulus sp., Elaphostrongylus rangiferi, Orthostrongylus sp. (possibly, O. macrotis), and Varestrongylus eleguneniensis. A micrograph of what we believe to be A. mosgovoyi is presented for the first time. This is the first report of V. eleguneniensis in the Palearctic. This is also the first report of Orthostrongylus sp. both in relation to R. tarandus and to the Palearctic.
In O. moschatus, no trematodes, one genus of cestodes (Moniezia), and various nematodes (small GINs, Nematodirus sp., Nematodirella sp., Trichuris sp., and the DSL of Protostrongylidae) were found.
In O. nivicola, no trematodes, one genus of cestodes (Moniezia), and various nematodes (small GINs, Marshallagia sp., Nematodirus sp., Trichuris sp., Capillarid-type, and Protostrongylus spp.) were found. This is the first report of a Capillarid-type nematode (and its egg micrograph) in snow sheep.

Author Contributions

Conceptualization, O.A.L., S.B.R., T.P.S., I.A.M., D.V.P., M.G.B. and S.E.S.; methodology, O.A.L. and S.E.S.; formal analysis, O.A.L.; investigation, O.A.L. and S.E.S.; resources, S.B.R., T.P.S., I.A.M., D.V.P., K.A.L., M.G.B., L.A.K., A.R.G., P.S.K., D.I.L., M.N.S., V.N.M. and E.G.M.; writing—original draft preparation, O.A.L.; writing—review and editing, S.B.R., T.P.S., I.A.M., D.V.P., K.A.L., M.G.B., L.A.K., A.R.G., P.S.K., D.I.L., M.N.S., V.N.M., E.G.M. and S.E.S.; visualization, O.A.L.; supervision, S.E.S.; project administration, S.E.S.; funding acquisition, D.V.P. and S.E.S. All authors have read and agreed to the published version of the manuscript.

Funding

The study related to Murmansk Oblast wild reindeer was performed under state order (project FMEN-2022–0003). An application of molecular techniques for the identification of nematodes in this study was supported by RSF grant № 19-74-20147 for S. E. Spiridonov.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article.

Acknowledgments

Authors thank all the colleagues and collectors, in particular Andrey Podkorytov, Andrey Przhiboro, Ivan Belokobylskiy, and Maksim Kropotov for their assistance.

Conflicts of Interest

The authors declare no conflict of interest.

Appendix A

Rotifers found in feces (including dried ones) of studied Arctic ruminants are shown in Figure A1.
Diversity 15 00672 g0a1 550
Figure A1. Bdelloid rotifers obtained from feces of reindeer (R. tarandus), muskoxen (O. moschatus), and snow sheep (O. nivicola). (A) Active rotifer from reindeer; (B) Active rotifer with an egg (arrow) from snow sheep; (C) Oval rotifer egg from reindeer; (D) Oval rotifer egg with polar protrusions from reindeer; (E) Oval rotifer egg with polar protrusion from muskox; (F) Lemon-shaped rotifer egg from reindeer; (G) Angular rotifer egg with 8 visible protrusions from reindeer; (H) Angular rotifer egg with 10 visible protrusions from reindeer; (I) Inert rotifer from reindeer (reviving); (J) Inert rotifer from snow sheep (reviving); (K) Inert rotifer from muskox (reviving); (L) Inert rotifer (reviving) with an egg (arrow) from reindeer; (M) Inert rotifer (reviving) with an egg (arrow) from muskox. Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Figure A1. Bdelloid rotifers obtained from feces of reindeer (R. tarandus), muskoxen (O. moschatus), and snow sheep (O. nivicola). (A) Active rotifer from reindeer; (B) Active rotifer with an egg (arrow) from snow sheep; (C) Oval rotifer egg from reindeer; (D) Oval rotifer egg with polar protrusions from reindeer; (E) Oval rotifer egg with polar protrusion from muskox; (F) Lemon-shaped rotifer egg from reindeer; (G) Angular rotifer egg with 8 visible protrusions from reindeer; (H) Angular rotifer egg with 10 visible protrusions from reindeer; (I) Inert rotifer from reindeer (reviving); (J) Inert rotifer from snow sheep (reviving); (K) Inert rotifer from muskox (reviving); (L) Inert rotifer (reviving) with an egg (arrow) from reindeer; (M) Inert rotifer (reviving) with an egg (arrow) from muskox. Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Diversity 15 00672 g0a1

References

  1. Taylor, J.J.; Lawler, J.P.; Aronsson, M.; Barry, T.; Bjorkman, A.D.; Christensen, T.; Coulson, S.J.; Cuyler, C.; Ehrich, D.; Falk, K.; et al. Arctic terrestrial biodiversity status and trends: A synopsis of science supporting the CBMP State of Arctic Terrestrial Biodiversity Report. Ambio 2020, 49, 833–847. [Google Scholar] [CrossRef]
  2. Convey, P. Antarctic terrestrial biodiversity in a changing world. Polar Biol. 2011, 34, 1629–1641. [Google Scholar] [CrossRef]
  3. Carlson, C.J.; Burgio, K.R.; Dougherty, E.R.; Phillips, A.J.; Bueno, V.M.; Clements, C.F.; Castaldo, G.; Dallas, T.A.; Cizauskas, C.A.; Cumming, G.S.; et al. Parasite biodiversity faces extinction and redistribution in a changing climate. Sci. Adv. 2017, 3, e1602422. [Google Scholar] [CrossRef]
  4. Zykin, V.S.; Zykina, V.S.; Orlova, L.A.; Stratigrafiya, I. Osnovnye zakonomernosti izmeneniya prirodnoy sredy I klimata v pleistotsene I golotsene Zapadnoy Sibiri (Stratigraphy and the main regularities of changes in the natural environment and climate in the Pleistocene and Holocene of Western Siberia). Arkheologiya 2000, 1, 3–22. (In Russian) [Google Scholar]
  5. European Glacial Landscapes. Maximum Extent of Glaciations; Palacois, D.; Hughes, P.D.; Garcia-Ruiz, J.M.; Andres, N. (Eds.) Elsevier: Amsterdam, The Netherlands, 2021; p. 546. ISBN 978-0-12-823498-3. [Google Scholar]
  6. Holand, Ø.; Mizin, I.; Weladji, R.B. Reindeer Rangifer tarandus (Linnaeus, 1758). In Handbook of the Mammals of Europe; Hackländer, K., Zachos, F.E., Eds.; Springer: Cham, Switzerland, 2022; pp. 247–277. ISBN 978-3-030-24474-3. [Google Scholar] [CrossRef]
  7. Mizin, I.A.; Sipko, T.P.; Davydov, A.V.; Gruzdev, A.R. The wild reindeer (Rangifer tarandus: Cervidae, Mammalia) on the Arctic Islands of Russia: A Review. Nat. Conserv. Res. 2018, 3, 1–14. [Google Scholar] [CrossRef]
  8. Danilov, P.I.; Panchenko, D.V.; Tirronen, K.F. Severnyy Olen’ Vostochnoy Fennoskandii (Reindeer of Eastern Fennoscandia); KarNTS RAN: Petrozavodsk, Russia, 2020; p. 187. ISBN 978-5-9274-0879-5. (In Russian) [Google Scholar]
  9. Korolev, A.N.; Mamontov, V.N.; Kholodova, M.V.; Baranova, A.I.; Shadrin, D.M.; Poroshin, E.A.; Efimov, V.A.; Kochanov, S.K. Polymorphism of the mtDNA control region in Reindeer (Rangifer tarandus) from the mainland of the Northeastern part of European Russia. Biol. Bull. 2017, 44, 882–893. [Google Scholar] [CrossRef]
  10. Safronov, V.M. Ekologiyai Ispol’zovaniye Dikogo Severnogo Olenya v Yakutii (Ecology and Use of Wild Reindeer in Yakutia); Izdatel’stvo SO RAN: Yakutsk, Russia, 2005; p. 188. ISBN 5-463-00087-5. (In Russian) [Google Scholar]
  11. Schmidt, N.M.; Stelvig, M. Muskox Ovibos moschatus (Zimmermann, 1780). In Handbook of the Mammals of Europe; Hackländer, K., Zachos, F.E., Eds.; Springer: Cham, Switzerland, 2022; pp. 313–325. ISBN 978-3-030-24474-3. [Google Scholar]
  12. Cuyler, C.; Rowell, J.; Adamczewski, J.; Anderson, M.; Blake, J.; Bretten, T.; Brodeur, V.; Campbel, M.; Checkley, S.L.; Cluff, H.D.; et al. Muskox status, recent variation, and uncertain future. Ambio 2020, 49, 805–819. [Google Scholar] [CrossRef]
  13. Revin, T.V.; Sopin, L.V.; Zheleznov, N.K. Snezhnyy Baran. Morfologiya, Sistematika, Ekologiya, Okhrana (Snow Sheep. Morphology, Systematics, Ecology, Protection); Nauka: Novosibirsk, Russia, 1988; p. 193. ISBN 5-02-029417-9. (In Russian) [Google Scholar]
  14. Baskin, L.; Danell, K. Ecology of Ungulates: A Handbook of Species in Eastern Europe and Northern and Central Asia; Springer: Berlin/Heidelberg, Germany; New York, NY, USA, 2003; pp. 378–385. ISBN 3-540-43804-1. [Google Scholar]
  15. Sipko, T.P.; Gruzdev, A.R.; Egorov, S.S.; Tikhonov, V.G. Analis protsessa introduktsii ovtsebyka na severe Evrazii (The Analysis of Muskox Introduction in Northern Asia). Zool. Zhurnal 2007, 86, 620–627. (In Russian) [Google Scholar]
  16. Gordeeva, N.V.; Sipko, T.P.; Gruzdev, A.R. Izmenchivost’ mikrosatellitnoy DNK v populyatsiyakh ovtsebykov (Ovibos moschatus), akklimatizirovannykh v Rossii (Microsatellite DNA Variability in the Populations of Muskoxen (Ovibos moschatus) Transplanted into the Russian North). Genetika 2009, 45, 932–940. (In Russian) [Google Scholar]
  17. Argunov, A.V. Alien Species of the Mammalian Fauna in Yakutia. Russ. J. Biol. Invasions. 2018, 9, 313–326. [Google Scholar] [CrossRef]
  18. Poulin, R.; Presswell, B.; Bennett, J.; de Angeli Dutra, D.; Salloum, P.M. Biases in parasite biodiversity research: Why some helminth species attract more research than others. Int. J. Parasitol. Parasites Wildl. 2023, 21, 89–98. [Google Scholar] [CrossRef] [PubMed]
  19. Mizkewitsch, V.Y. Gel’minty Severnogo Olenya i Vyzyvayemyye Imi Zabolevaniya (Reindeer Helminths and the Diseases They Cause); Kolos: Leningrad, Russia, 1967; 308p. (In Russian) [Google Scholar]
  20. Emelyanova, A.; Savolainen, A.; Oksanen, A.; Nieminen, P.; Loginova, O.; Abass, K.; Rautio, A. Research on Selected Wildlife Infections in the Circumpolar Arctic—A Bibliometric Review. Int. J. Environ. Res. Public Health 2022, 19, 11260. [Google Scholar] [CrossRef]
  21. Latypov, D.G.; Timerbayeva, R.R.; Kirillov, E.G. Parazitologiya i Invazionnyye Bolezni Zhvachnykh Zhivotnykh (Parasitology and Invasive Diseases of Ruminants); Lan: St. Petersburg, Russia, 2019; pp. 59–67. ISBN 978-5-8114-3561-6. (In Russian) [Google Scholar]
  22. Verocai, G.G.; Chaudhry, U.N.; Lejeune, M. Diagnostic Methods for Detecting Internal Parasites of Livestock. Vet. Clin. Food. Anim. 2020, 36, 125–143. [Google Scholar] [CrossRef] [PubMed]
  23. Holterman, M.; van der Wurff, A.; van den Elsen, S.; van Megen, H.; Bongers, T.; Holovachov, O.; Bakker, J.; Helder, J. Phylum-Wide Analysis of SSU rDNA Reveals Deep Phylogenetic Relationships among Nematodes and Accelerated Evolution toward Crown Clades. Mol. Biol. Evol. 2006, 23, 1792–1800. [Google Scholar] [CrossRef]
  24. Vrain, T.C.; Wakarchuk, D.A.; Levesque, A.C.; Hamilton, R.I. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundam. Appl. Nematol. 1992, 15, 563–573. [Google Scholar]
  25. Gasser, R.B.; Chilton, N.B.; Hoste, H.; Beveridge, I. Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res. 1993, 21, 2525–2526. [Google Scholar] [CrossRef] [PubMed]
  26. Nadler, S.A.; Hudspeth, D.S.S. Phylogeny of the Ascaridoidea (Nematoda: Ascaridida) based on three genes and morphology: Hypotheses of structural and sequence evolution. J. Parasitol. 2000, 86, 380–393. [Google Scholar] [CrossRef]
  27. Thomas, W.K.; Vida, J.T.; Frisse, L.M.; Mundo, M.; Baldwin, J.G. DNA sequences from formalin-fixed nematodes: Integrating molecular and morphological approaches to taxonomy. J. Nematol. 1997, 29, 250–254. [Google Scholar]
  28. Kanzaki, N.; Futai, K. A PCR primer set for determination of phylogenetic relationships of Bursaphelenchus species within the xylophilus group. Nematology 2002, 4, 35–41. [Google Scholar] [CrossRef]
  29. Zhang, Z.; Schwartz, S.; Wagner, L.; Miller, W. A greedy algorithm for aligning DNA sequences. J. Comput. Biol. 2004, 7, 203–214. [Google Scholar] [CrossRef]
  30. Morgulis, A.; Coulouris, G.; Raytselis, Y.; Madden, T.L.; Agarwala, R.; Schäffer, A.A. Database indexing for production MegaBLAST searches. Bioinformatics 2008, 24, 1757–1764. [Google Scholar] [CrossRef]
  31. Loginova, O.A.; Kolpashchikov, L.A.; Spiridonov, S.E. First report of Orthostrongylus sp. (Nematoda: Protostrongylidae) in wild reindeer (Rangifer tarandus) from the Taimyr, Russia: Nearctic parasites in a Palearctic host. Parasitol. Res. 2023, 122, 685–689. [Google Scholar] [CrossRef]
  32. Kutz, S.J.; Ducrocq, J.; Verocai, G.G.; Hoar, B.M.; Colwell, D.D.; Beckmen, K.B.; Polley, L.; Elkin, B.T.; Hoberg, E.P. Parasites in ungulates of Arctic North America and Greenland: A view of contemporary diversity, ecology, and impact in a world under change. Adv. Parasitol. 2012, 79, 99–252. [Google Scholar] [CrossRef] [PubMed]
  33. Kutz, S.J.; Laaksonen, S.; Asbakk, K.; Nilssen, A.C. Parasitic Infections and Diseases. In Reindeer and Caribou. Health and Disease; Tryland, M., Kutz., S.J., Eds.; CRC Press (Taylor and Francis Group): Boca Raton, FL, USA; London, UK; New York, NY, USA, 2019; pp. 177–235. ISBN 978-1-4822-5068-8. [Google Scholar]
  34. Eduardo, S.L. The taxonomy of the family Paramphistomatidae Fischoeder, 1901 with special references to the morphology of species occurring in ruminants. II. Revision of the genus Paramphistomum Fischoeder, 1901. Syst. Parasitol. 1982, 4, 189–238. [Google Scholar] [CrossRef]
  35. Skrjabin, K.I. Trematody Zhivotnykh i Cheloveka. Osnovy Trematodologii (Trematodes of Animals and Man. Principles of Trematodology); Izdatelstvo Akademii Nauk SSSR: Moscow, Russia, 1949; Volume 3, p. 600. (In Russian) [Google Scholar]
  36. Zhaltsanova, D.S. Occurrence of Fischoederius elongatus in the Buryat ASSR. Mater. Nauchnykh Issled. Chlenov Vsesoyuznogo Obs. Gel’mintologov 1972, 24, 49–50. (In Russian) [Google Scholar]
  37. Willmott, S.; Pester, F.R.N. Variations in Faecal Egg-counts in Paramphistome Infections as Determined by a New Technique. J. Helminthol. 1952, 26, 2–3+147–156. [Google Scholar] [CrossRef]
  38. Skrjabin, K.I. Trematody Zhivotnykh i Cheloveka. Osnovy Trematodologii (Trematodes of Animals and Man. Principles of Trematodology); Izdatelstvo Akademii Nauk SSSR: Moscow, Russia, 1952; Volume 7, p. 762. (In Russian) [Google Scholar]
  39. Haukisalmi, V.; Laaksonen, S.; Oksanen, A.; Beckmen, K.; Halajian, A.; Yanagida, T.; Nakao, M. Molecular taxonomy and subgeneric classification of tapeworms of the genus Moniezia Blanchard, 1891 (Cestoda, Anoplocephalidae) in northern cervids (Alces and Rangifer). Parasitol. Int. 2018, 67, 218–224. [Google Scholar] [CrossRef]
  40. Meradi, S.; Bentounsi, B.; Zouyed, I.; Cabaret, J. The steppe species of gastrointestinal nematodes of small ruminants, with a focus on Marshallagia: Climate as a key determinant. Parasite J. De La Société Française De Parasitol. 2011, 183, 261–269. [Google Scholar] [CrossRef]
  41. Skrjabin, K.I.; Shikhobalova, N.P.; Schultz, R.S. Trikhostrongilidy Zhivotnykh i cheloveka. Osnovy Nematodologii (Trichostrongylids of Animals and Man. Principles of Nematodology); Izdatelstvo Akademii Nauk SSSR: Moscow, Russian, 1954; Volume 3, p. 684. (In Russian) [Google Scholar]
  42. Loginova, O.A.; Chuprak, D.I.; Rozenfel’d, S.B. O samostoyatel’nom diagnosticheskom znachenii obolochki yaytsa Marshallagia spp. Nnematoda: Strongylida) (On independent diagnostic value of egg membranes of Marshallagia spp. (Nematoda: Strongylida)). In Proceedings of the Modern Issues of General and Applied Parasitology Conference, Voronezh, Russia, 27–28 October 2022; pp. 56–61. (In Russian). [Google Scholar] [CrossRef]
  43. Dallas, J.F.; Irvine, R.J.; Halvorsen, O. DNA evidence that Marshallagia marshalli Ransom, 1907 and M. occidentalis Ransom, 1907 (Nematoda: Ostertagiinae) from Svalbard reindeer are conspecific. Syst. Parasitol. 2001, 50, 101–103. [Google Scholar] [CrossRef]
  44. Kuchboev, A.; Sobirova, K.; Karimova, R.; Amirov, O.; von Samson-Himmelstjerna, G.; Krücken, J. Molecular analysis of polymorphic species of the genus Marshallagia (Nematoda: Ostertagiinae). Parasites Vectors 2020, 13, 1–12. [Google Scholar] [CrossRef]
  45. Fruetel, M.; Lankester, M.W. Gastrointestinal helminths of woodland and barren ground caribou (Rangifer tarandus) in Canada, with keys to species. Can. J. Zool. 1989, 67, 2253–2269. [Google Scholar] [CrossRef]
  46. Thomas, R.J. A comparative study of the infective larvae of Nematodirus species parasitic in sheep. Parasitology 1957, 47, 60–65. [Google Scholar] [CrossRef]
  47. Baskin, L.M. Odomashnivaniye Severnogo Olenya. Ot Okhotnika Do Pastukha i Ranchevoda (Reindeer Domestication. from Hunter to Herdsman and Rancher); Tovarishchestvo Nauchnykh Izdaniy KMK: Moscow, Russia, 2021; p. 280. ISBN 978-5-907372-89-4. (In Russian) [Google Scholar]
  48. Hrabok, J.T.; Oksanen, A.; Nieminen, M.; Waller, P.J. Population dynamics of nematode parasites of reindeer in the sub-arctic. Vet. Parasitol. 2006, 142, 301–311. [Google Scholar] [CrossRef]
  49. Jokelainen, P.; Moroni, B.; Hoberg, E.; Oksanen, A.; Laaksonen, S. Gastrointestinal parasites in reindeer (Rangifer tarandus tarandus): A review focusing on Fennoscandia. Vet. Parasitol. Reg. Stud. Rep. 2019, 17, 100317. [Google Scholar] [CrossRef]
  50. Turgeon, G.; Kutz, S.J.; Lejeune, M.; St-Laurent, M.H.; Pelletier, F. Parasite prevalence, infection intensity and richness in an endangered population, the Atlantic-Gaspésie caribou. Int. J. Parasitol. Parasites Wildl. 2018, 7, 90–94. [Google Scholar] [CrossRef] [PubMed]
  51. Brown, S.J.; Goetzmann, W.; Ibbotson, R.G.; Ross, S.A. Survivorship bias in performance studies. Rev. Financ. Stud. 1992, 5, 553–580. [Google Scholar] [CrossRef]
  52. Pyziel, A.M.; Laskowski, Z.; Demiaszkiewicz, A.W.; Höglund, J. Interrelationships of Dictyocaulus spp. in wild ruminants with morphological description of Dictyocaulus cervi n. sp.(Nematoda: Trichostrongyloidea) from red deer, Cervus elaphus. J. Parasitol. 2017, 103, 506–518. [Google Scholar] [CrossRef]
  53. Filip-Hutsch, K.; Demiaszkiewicz, A.W.; Chęcińska, A.; Hutsch, T.; Czopowicz, M.; Pyziel, A.M. First report of a newly-described lungworm, Dictyocaulus cervi (Nematoda: Trichostrongyloidea), in moose (Alces alces) in central Europe. Int. J. Parasitol. Parasites Wildl. 2020, 13, 275–282. [Google Scholar] [CrossRef]
  54. Verocai, G.G.; Kutz, S.J.; Simard, M.; Hoberg, E.P. Varestrongylus eleguneniensis sp. n. (Nematoda: Protostrongylidae): A widespread, multi-host lungworm of wild North American ungulates, with an emended diagnosis for the genus and explorations of biogeography. Parasites Vectors 2014, 7, 1–22. [Google Scholar] [CrossRef]
  55. Verocai, G.G.; Hoberg, E.P.; Simard, M.; Beckmen, K.B.; Musiani, M.; Wasser, S.; Cuyler, C.; Manseau, M.; Chaudhry, U.N.; Kashivakura, C.K.; et al. The biogeography of the caribou lungworm, Varestrongylus eleguneniensis (Nematoda: Protostrongylidae) across northern North America. Int. J. Parasitol. Parasites Wildl. 2020, 11, 93–102. [Google Scholar] [CrossRef]
  56. Boev, S.N. Protostrongilidy. Osnovy Nematodologii (Protostrongylids. Principles of Nematodology); Izdatelstvo Akademii Nauk SSSR: Moscow, Russian, 1975; Volume 25, p. 338. (In Russian) [Google Scholar]
  57. Verocai, G.G.; Kafle, P.; Sulliotti, V.; Lejeune, M.; Hoberg, E.P.; Kutz, S.J. Morphometry of first-stage larva of Orthostrongylus macrotis (Nematoda: Protostrongylidae), lungworm of wild ungulates from Western North America. J. Parasitol. 2022, 108, 322–329. [Google Scholar] [CrossRef] [PubMed]
  58. Greiner, E.C. Laboratory Diagnostics of Parasitic Diseases. In Equine Infectious Diseases, 2nd ed.; Sellon, D.C., Long, M.T., Eds.; Elsevier: Amsterdam, The Netherlands, 2014; pp. 449–456. ISBN 978-1455708918. [Google Scholar]
  59. Oksanen, A. Endectocide treatment of the reindeer. Rangifer 1999, 19, 7–50. [Google Scholar] [CrossRef]
  60. Loginova, O.A. D-shaped nematode eggs in the feces of Rangifer tarandus. RJN 2023. submitted. [Google Scholar]
  61. Rebecchi, L.; Boschetti, C.; Nelson, D.R. Extreme-tolerance mechanisms in meiofaunal organisms: A case study with tardigrades, rotifers and nematodes. Hydrobiologia 2020, 847, 2779–2799. [Google Scholar] [CrossRef]
Diversity 15 00672 g001 550
Figure 1. Map of Palearctic indicating the sampling sites. Numbers correspond to identification numbers in Table 1.
Figure 1. Map of Palearctic indicating the sampling sites. Numbers correspond to identification numbers in Table 1.
Diversity 15 00672 g001
Diversity 15 00672 g002 550
Figure 2. Diagnostic stages of helminths obtained from feces of reindeer (R. tarandus). (A) Paramphistomum sp. egg; (B) Dicrocoelium sp. egg; (C) Moniezia sp. egg; (D) Strongyle-type egg; (E) Marshallagia sp. egg (embryonated, dead); (F) Nematodirus sp. egg; (G) Nematodirella longissimespiculata egg; (H) Skrjabinema tarandi egg; (I) Trichuris sp. egg; (J) Capillaria sp. egg; (K) Possibly, Ascaris mosgovoyi egg (embryonated, dead); (L) Dictyocaulus sp. first-stage larva (L1); (M) Elaphostrongylus rangiferi L1; (N) Orthostrongylus sp. (possibly, O. macrotis) L1; (O) Varestrongylus eleguneniensis L1 (dead). Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Figure 2. Diagnostic stages of helminths obtained from feces of reindeer (R. tarandus). (A) Paramphistomum sp. egg; (B) Dicrocoelium sp. egg; (C) Moniezia sp. egg; (D) Strongyle-type egg; (E) Marshallagia sp. egg (embryonated, dead); (F) Nematodirus sp. egg; (G) Nematodirella longissimespiculata egg; (H) Skrjabinema tarandi egg; (I) Trichuris sp. egg; (J) Capillaria sp. egg; (K) Possibly, Ascaris mosgovoyi egg (embryonated, dead); (L) Dictyocaulus sp. first-stage larva (L1); (M) Elaphostrongylus rangiferi L1; (N) Orthostrongylus sp. (possibly, O. macrotis) L1; (O) Varestrongylus eleguneniensis L1 (dead). Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Diversity 15 00672 g002
Diversity 15 00672 g003 550
Figure 3. Diagnostic stages of helminths obtained from feces of muskoxen (O. moschatus). (A) Moniezia sp. egg; (B) Strongyle-type egg; (C) Nematodirus sp. egg; (D) Nematodirella sp. egg; (E) Trichuris sp. egg; (F) first-stage larva (L1) of the family Protostrongylidae. Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Figure 3. Diagnostic stages of helminths obtained from feces of muskoxen (O. moschatus). (A) Moniezia sp. egg; (B) Strongyle-type egg; (C) Nematodirus sp. egg; (D) Nematodirella sp. egg; (E) Trichuris sp. egg; (F) first-stage larva (L1) of the family Protostrongylidae. Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Diversity 15 00672 g003
Diversity 15 00672 g004 550
Figure 4. Diagnostic stages of helminths obtained from feces of snow sheep (O. nivicola). (A) Moniezia sp. egg; (B) Strongyle-type egg; (C) Marshallagia sp. egg (embryonated, dead); (D) Nematodirus sp. egg (embryonated); (E) Trichuris sp. egg; (F) Capillarid-type egg; (G) Protostrongylus sp. first stage larva (L1) with a longer tail spike (sample set #23); (H) Protostrongylus sp. L1 with a shorter tail spike (sample set #24). Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Figure 4. Diagnostic stages of helminths obtained from feces of snow sheep (O. nivicola). (A) Moniezia sp. egg; (B) Strongyle-type egg; (C) Marshallagia sp. egg (embryonated, dead); (D) Nematodirus sp. egg (embryonated); (E) Trichuris sp. egg; (F) Capillarid-type egg; (G) Protostrongylus sp. first stage larva (L1) with a longer tail spike (sample set #23); (H) Protostrongylus sp. L1 with a shorter tail spike (sample set #24). Bright field microscopy, 40× objective lens magnification. Scale bar equals 50 μm.
Diversity 15 00672 g004
Table
Table 1. Collection data for fecal samples from reindeer, muskoxen, and snow sheep across the Northern Palearctic.
Table 1. Collection data for fecal samples from reindeer, muskoxen, and snow sheep across the Northern Palearctic.
Sample Set IDMap ReferenceHost SpeciesNumber of Fecal SamplesLocationCoordinates
(Decimal Degrees)		Date Collected
11Wild reindeer (Rangifer tarandus)19Taymyr Peninsula
(Krasnoyarsk Krai)		71.10780,
98.04150		June 2020
223Novaya Zemlya,
Arkhangelsk Oblast		76.90835,
68.44880		August 2021
322Novaya Zemlya,
Arkhangelsk Oblast		76.90835,
68.44880		July 2022
4315Barentsbugr, Svalbard 1
(Norway)		78.04729,
14.28717		August 2022
5421Murmansk Oblast68.50699,
30.03720		September 2022
6511Arkhangelsk Oblast63.86779,
44.43266		September 2022
76Semiwild reindeer (R. tarandus)50Yamalo-Nenets
Autonomous Okrug		66.62200,
65.71646		August 2018
8759Nenets Autonomous Okrug67.97568,
52.91955		October 2018
982Khanty-Mansi
Autonomous Okrug		65.63307,
61.94561		October 2019
10930Murmansk Oblast68.00424,
35.01228		December 2020
11931Murmansk Oblast68.00424,
35.01228		March 2021
12820Khanty-Mansi
Autonomous Okrug		65.63307,
61.94561		August 2021
131042Bering Island 2
Kamchatka Krai		55.26152,
165.93711		October 2021
14117Republic of Karelia66.30072,
33.08216		June 2022
151210Republic of Sakha (Yakutia)65.34745,
140.08864		June 2022
161330Chukotka
Autonomous Okrug		65.46168,
−173.47478		June 2022
17148Chukotka
Autonomous Okrug		65.90614,
−178.84644		October 2022
1815Muskoxen (Ovibos moschatus)35Yamalo-Nenets
Autonomous Okrug		67.58240,
66.25280		March 2022
191624Wrangel Island,
Chukotka
Autonomous Okrug		71.34890,
−178.66722		March 2022
20176Taymyr Peninsula
(Krasnoyarsk Krai)		73.10336,
95.11816		July 2022
211825Taymyr Peninsula
(Krasnoyarsk Krai)		74.17696,
107.46475		August 2022
221530Yamalo-Nenets
Autonomous Okrug		67.58240,
66.25280		October 2022
2319Snow sheep
(Ovis nivicola)		21Taymyr Peninsula
(Krasnoyarsk Krai)		68.64397,
95.66562		July 2016
242013Republic of Sakha (Yakutia)70.51859,
129.50336		June 2022
25215Taymyr Peninsula
(Krasnoyarsk Krai)		69.33130,
93.58480		July 2022
1 Non-Russian territory included due to Svalbard Treaty. 2 Site with originally domestic reindeer that eventually feralized.
Table
Table 2. Species of parasitic nematodes recovered from reindeer feces and identified genetically.
Table 2. Species of parasitic nematodes recovered from reindeer feces and identified genetically.
Sample Set IDSpeciesGenBank 1Vouchers 2
1Orthostrongylus sp. 3OL700043 (for LSU)IPEE_Parasites 14284
OL700044 (for ITS2)IPEE_Parasites 14284
1Varestrongylus eleguneniensis3OM743794 (for ITS2)IPEE_Parasites 14298
OL743795 (for ITS2)IPEE_Parasites 14299
10Elaphostrongylus rangiferi 4OQ731723 (for ITS)IPEE_Parasites 14310
OQ730416 (for COX1)IPEE_Parasites 14310
OQ746340 (for LSU)IPEE_Parasites 14310
12Elaphostrongylus rangiferi 4OQ731722 (for ITS2)IPEE_Parasites 14309
13Ostertagia arctica 4OQ726410 (for ITS2)IPEE_Parasites 14311
13Nematodirella longissimespiculata 4OQ726409 (for ITS2)IPEE_Parasites 14312
1 GenBank accession numbers for sequences from the individual first larval stage (L1) or third larval stage (L3) of representative species. 2 Voucher specimens with definitive identifications and accession numbers archived in the Museum of Helminthological Collections of the Parasitology Center at the A. N. Severtsov Institute of Ecology and Evolution of the Russian Academy of Sciences (Moscow, Russia). 3 Possibly, O. macrotis. Sequence previously in GenBank as reported by Loginova et al. [31]. 4 Sequence information reported here for the first time.
Table
Table 3. Helminths found in the feces of reindeer, muskoxen, and snow sheep.
Table 3. Helminths found in the feces of reindeer, muskoxen, and snow sheep.
Sample Set IDHost SpeciesTrematodesCestodesNematodes
Paramphistomum sp.Dicrocoelium sp.Moniezia sp.Strongyle-TypeMarshallagia sp.Nematodirus sp.Nematodirella sp.Dictyocaulus sp.ProtostrongylidaeSkrjabinema sp.Trichuris sp.Capillarid-TypeAscaris-Type
1Wild reindeer (Rangifer tarandus)9 (47%)––15 (79%)–1 (5%)––1 (5%) 3
3 (16%) 4		––––
2–––1 (33%)–––––––––
3–––1 (50%)–––––––––
4–––11 (73%)1 (7%)––––––––
5–––14 (67%)––––4 (19%) 5––––
6–––11 (100%)––––––1 (9%)––
7Semiwild reindeer (R. tarandus)––2 (4%)44 (88%)–1 (2%)––6 (12%) 5––––
8–1 (2%)16 (27%)47 (80%)–18 (31%)11 (19%)–12 (21%) 52 (3%) 9–––
9––1 (50%)2 (100%)––––––1 (50%)––
10–––15 (50%)–7 (23%)––21 (70%) 6––12 (40%) 10 –
118 (26%)––6 (19%)–––1 (3%)16 (52%) 5––––
129 (45%)––1 (5%)–––1 (5%)6 (30%) 6––3 (15%) 10–
13––2 (5%)27 (64%) 1–4 (10%)5 (12%) 22 (5%)–––14 (33%) 10–
14–––4 (57%)––––4 (57%) 5––1 (14%) 10–
15–––4 (40%)–––––––––
16––2 (7%)18 (60%)–––––2 (7%) 9––1 (3%) 11
17–––––2 (25%)3 (38%)––––––
18Muskoxen (Ovibos moschatus)––2 (6%)––1 (3%)––19 (54%) 5–1 (3%)––
19––4 (17%)3 (13%)––1 (4%)––––––
20–––2 (33%)–––––––––
21–––6 (24%)–––––––––
22––1 (3%)9 (30%)–1 (3%)––8 (26%) 5––––
23Snow sheep (Ovis nivicola)––––4 (19%)–––16 (76%) 7–4 (19%)––
24––2 (15%)–7 (54%)1 (8%)––9 (69%) 8–5 (39%)1 (8%)–
25–––1 (20%)–––––––––
1 Including Ostertagia arctica. 2 Nematodirella longissimespiculata. 3 Orthostrongylus sp. (possibly, O. macrotis). 4 Varestrongylus eleguneniensis. 5 Dorsal-spined larvae (DSL). 6 Elaphostrongylus rangiferi. 7 Protostrongylus sp. with a longer tail spike. 8 Protostrongylus sp. with a shorter tail spike. 9 Skrjabinema tarandi. 10 Capillaria sp. 11 Possibly, Ascaris mosgovoyi.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Loginova, O.A.; Rozenfeld, S.B.; Sipko, T.P.; Mizin, I.A.; Panchenko, D.V.; Laishev, K.A.; Bondar, M.G.; Kolpashchikov, L.A.; Gruzdev, A.R.; Kulemeev, P.S.; et al. Diversity and Distribution of Helminths in Wild Ruminants of the Russian Arctic: Reindeer (Rangifer tarandus), Muskoxen (Ovibos moschatus), and Snow Sheep (Ovis nivicola). Diversity 2023, 15, 672. https://doi.org/10.3390/d15050672

AMA Style

Loginova OA, Rozenfeld SB, Sipko TP, Mizin IA, Panchenko DV, Laishev KA, Bondar MG, Kolpashchikov LA, Gruzdev AR, Kulemeev PS, et al. Diversity and Distribution of Helminths in Wild Ruminants of the Russian Arctic: Reindeer (Rangifer tarandus), Muskoxen (Ovibos moschatus), and Snow Sheep (Ovis nivicola). Diversity. 2023; 15(5):672. https://doi.org/10.3390/d15050672

Chicago/Turabian Style

Loginova, Olga A., Sofya B. Rozenfeld, Taras P. Sipko, Ivan A. Mizin, Danila V. Panchenko, Kasim A. Laishev, Mikhail G. Bondar, Leonid A. Kolpashchikov, Aleksandr R. Gruzdev, Pavel S. Kulemeev, and et al. 2023. "Diversity and Distribution of Helminths in Wild Ruminants of the Russian Arctic: Reindeer (Rangifer tarandus), Muskoxen (Ovibos moschatus), and Snow Sheep (Ovis nivicola)" Diversity 15, no. 5: 672. https://doi.org/10.3390/d15050672

APA Style

Loginova, O. A., Rozenfeld, S. B., Sipko, T. P., Mizin, I. A., Panchenko, D. V., Laishev, K. A., Bondar, M. G., Kolpashchikov, L. A., Gruzdev, A. R., Kulemeev, P. S., Litovka, D. I., Semerikova, M. N., Mamontov, V. N., Mamaev, E. G., & Spiridonov, S. E. (2023). Diversity and Distribution of Helminths in Wild Ruminants of the Russian Arctic: Reindeer (Rangifer tarandus), Muskoxen (Ovibos moschatus), and Snow Sheep (Ovis nivicola). Diversity, 15(5), 672. https://doi.org/10.3390/d15050672

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop