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Review

Trichuriasis in Selected Deer (Cervidae) Species: A Geographical Perspective

by
Kegan Romelle Jones
1,2
1
Department of Food Production (DFP), Faculty of Food and Agriculture (FFA), St. Augustine Campus, University of the West Indies (UWI), St. Augustine 999183, Trinidad and Tobago
2
Department of Basic Veterinary Sciences (DBVS), Faculty of Medical Sciences (FMS), St. Augustine Campus, School of Veterinary Medicine (SVM), University of the West Indies (UWI), St. Augustine 999183, Trinidad and Tobago
Ruminants 2021, 1(2), 178-190; https://doi.org/10.3390/ruminants1020013
Submission received: 19 October 2021 / Revised: 19 November 2021 / Accepted: 24 November 2021 / Published: 26 November 2021
(This article belongs to the Special Issue Feature Papers of Ruminants 2021-2022)
Versions Notes

Abstract

:
Trichuris spp. are endoparasites found in a wide range of mammalian species. Some of these host species include humans, non-human primates, dogs, cats, pigs, wild ruminants and domesticated ruminants. However, it had been noted that Trichuris are host specific, therefore the parasites that infects wild ruminant species may be transmitted to domesticated animals such as cattle, sheep and goat. Thus, the aim of this review was to identify species of Trichuris that parasitise deer species and to categorise the prevalence of this disease at various geographical locations. It must be noted that the prevalence and intensity of this parasite within deer species was low and rarely showed any signs of clinical disease. However, deer can be a source of infection to domesticated ruminants that may be housed in closed proximity. The review is divided into several sections based on the geographical location of the deer species. In summary, the review shows that most of the identification of various species of Trichuris in deer is based on morphological techniques. However, the use of molecular techniques in the identification of various species of Trichuris is more accurate. In closing, there is a need for more molecular investigations to be done in identifying the species of Trichuris that are present in deer living in the neo-tropical region.

1. Introduction

This review attempts to provide insight into the Trichuris species that inhabit the gastrointestinal tract of selected deer species. Deer species were chosen as they play a major ecological role in many wildlife reserves. In some locations, deer species are under threat due to the destruction of habitat as well as severe hunting pressure. Whipworm is one of the least studied endoparasites that affect Cervidae. The deer species that will be highlighted were categorized into four groups based on geographic regions. Recently, Jones et al. [1] reviewed endoparasites found in neo-tropical mammals and the red brocket deer (Mazama americana) was found to have Trichuris spp. present in its gastrointestinal tract but the species was not identified. Further to this, Jones [2] reviewed Trichuris spp. found in rodents. The review is a continuation of this series that investigated Trichuris spp. present in mammals.
It is well known that endoparasites can have a detrimental effect on wildlife populations as well as infect domestic ruminants, thus causing detrimental effects to health and performance. Trichuris spp. are commonly known as whipworm and they inhabit the caecum and colon of animals causing diarrhea when the level of infestation is high [3]. Hendrix and Robinson [4] stated that Trichuris ovis infections are usually asymptomatic but melena, anorexia and anemia can be seen with high levels of infestation. It is hypothesised that deer species that are located in the neo-tropics will have a higher prevalence of trichuriasis in comparison to other geographical regions. One reason may be the more favorable environmental conditions, which are conducive for ova development throughout the year. There has not been a published review that investigates the prevalence of Trichuris spp. in deer at various geographical locations. Thus, the objectives of this review were to show the prevalence and clinical signs of Trichuris spp. in selected species of deer at various localities. The laboratory techniques for the identification of whipworm will also be investigated. The disease is transmitted through the oro-faecal route; bi-polar eggs are produced in the faeces of the infected animal and contaminate the environment. Susceptible animals become infected when these eggs are consumed.

2. Methodology

An exhaustive literature search was conducted on several search engines. These engines included Google Scholar, UWI Linc, Pubmed and Scopus. The keywords used in the search included deer, Cervidae, Trichuris and Trichuriasis. The number of articles attained from these searches were 256. The inclusion criteria included prevalence as well as clinical signs of disease. Of the 256 articles searched, only 70 were appropriate to be included in this review. The literature search spanned over fifty years (1966–2021).

3. Trichuris in Deer Located in the Neo-Tropics

In this section, we will focus on Trichuriasis in two neo-tropical deer species, which are the brocket deer (Mazama spp.) and the white-tailed deer (Odocoileus virginianus). Knight et al. [5] analysed the morphometry of adult (male and female) Trichuris found in the caecum of the white-tailed deer. Based on the morphometry of this helminth, he proposed a new species called Trichuris odocoileus. Prior to this, endoparasites in white-tailed deer was studied and several authors identified Trichuris ovis in the caecum of deer (based on morphometry) [6,7,8]. Trichuris ovis and T skrjabini has also been reported in sheep that share similar pastures to deer [9]. The prevalence of Trichuris ovis in white-tailed deer is relatively low and it seems to have little clinical effect on the animal. The prevalence was reported as 3.3% [8], 15% [9], and 3.4% [7] (also see Table 1). Similar prevalence values have been reported in cattle: 3.8% [10], 1.26% [11], 7.3% and 13.2% [12]. Large ruminants such as the bison were found to have a low rate of infection, as low as 1% [13]. In contrast, small ruminants (sheep and goats) had a wide range of prevalence for Trichuris [14,15]. Using faecal flotation, sheep and goats had a prevalence of 40.46% and 50.51% [15], 4.9% and 4.1% [14]. Using morphological techniques, the prevalence in sheep and goat was 8.9% and 6.7% [14]. Further research on sheep reported a variable prevalence of 6.25% (using faecal flotation) and 27.42% (using morphology of adults) [16]. Recently, Yevstafieva et al. [17] reported 65% of sheep were infected with Trichuris spp. using morphological techniques. Knight and Tuff [18] identified Trichuris skrjabini in Sika deer (Cervus Nippon) based on morphological analysis but the animals showed no overt signs of disease. Cook et al. [19] noted that whipworms in the white-tailed deer had a prevalence of 4.76% (4/84) but failed to identify the species of the parasite. It must also be noted that one study failed to identify Trichuris spp. in white-tailed deer but found it in the Sambar deer living in the same population [19]. Interestingly, there were few studies that investigated the causes of morbidity and mortality of farmed white-tailed deer. In this review, based on morphological data, only one (1/347) case of Trichuris spp. infestation was identified as a cause of death in farmed white-tailed deer [20].
In recent times, archaeological studies have identified Trichuris spp. in deer that inhabited the neo-tropics [21,22]. These samples were found in Brazil and Argentina and shed some light on parasitism by Trichuris in wild deer species before colonisation. It also shows that these parasites were present in these animals before the arrival of domesticated livestock. Disease surveys have been conducted on free-ranging grey brocket deer in Bolivia and Brazil [23,24]. The authors noted that these animals were in good body condition before samples were taken. The prevalence was very low, which was similar to other reports in neo-tropical deer species with 9.09% (1/11) in Bolivia [23]. Lux Hoppe [24] was unable to detect Trichuris in deer samples that were collected. However, it must be noted that Lux Hoppe [24] did not analyse caecal or colonic contents which are the predilection sites for this parasite. Some work was done on the causes of mortality in Key deer (Odocoileus virginianus clavium) in Florida [25]. The authors identified several causes of morbidity and mortality in these animals such as haemonchosis, highway mortality and chronic purulent infection [25]. However, the effect of endoparasites on mortality was overlooked as well as the effect of individual parasites such as species of Trichuris. Trichuris spp. was identified in the gastrointestinal tracts of white-tailed deer and pampas deer in Mexico and Uruguay [26,27]. Thus, in these neo-tropical regions more work has to be done to investigate the prevalence of this parasite and its effect on wild populations. There were numerous investigations on gastrointestinal parasites found in deer in the neo-tropics. Most of these reports did not identify Trichuris spp. in the gastrointestinal tracts of white-tailed deer and fallow deer [28,29,30,31,32,33,34]. Interestingly, the studies mentioned above used morphological techniques as the method of parasite identification. This shows that the use of this technique as the sole means of identification of parasites can be inaccurate. In summary, the prevalence of this parasite found in deer within the neo-tropics is generally low (see Table 1)

4. Trichuris in Deer Located in Europe

Research was done in Czechoslovakia on endoparasites in roe deer. Two species of whipworm were identified as Trichocephalus (syn. Trichuris) capreoli and T. globulosa, which had a prevalence of 8.9% and 21.4%, respectively [42]. In Ireland, Trichuris spp. was found in red deer (Cervus elaphus) and fallow deer (Dama dama). Trichuris spp. occupied the large intestines of young or sick deer but never occurred in healthy deer. Trichuris ovis was found in red deer and an unidentified species of Trichuris was found in a fallow deer fawn. In the unidentified species, no males were recovered, which made identification based on morphology impossible [43]. In France, the relationship between helminth infestation and body condition in roe deer was investigated. Roe deer were hunted and the gastrointestinal tract and content were observed for adult helminths and eggs. Trichuris capreoli was found in the caecum and young animals had a higher infestation of helminths [44]. In a follow-up study done in roe deer, Trichuris capreoli infected males more often than females for each age and class category [45].
In Slovenia, wild fallow deer were hunted and gastrointestinal contents were analysed for helminths. These animals were clinically healthy when they were hunted with low number of parasites found in single animals [46]. Surprisingly, there were no gross pathological lesions attributed to the presence of nematodes in any of the infested digestive tracts. Three species of Trichuris were identified based on morphological analysis. They were T. globulosa, T. capreoli and T. ovis. Only a few deer were infected, 7% were infested with T. globulosa, 2% with T. capreoli and 2% with T. ovis [46]. Recently, a survey that lasted twenty months was done to investigate Cervidae kept in zoos in Belgium. The Cervidae under investigation included fallow deer (Dama dama), Dybowski’s deer (Cervus nippon dybiwski), puda (Puda puda) and reindeer (Rangifer tarandus tarandus) [47]. Adult helminths were collected from the carcasses of the animals and it was noted that the Trichuris spp. was present in one of the two zoos surveyed. The reindeer at the zoo were found to have a prevalence of 25% (n = 4). All other cervid species were negative for Trichuris spp. based on morphological identification of the adult worms [47].
Helminth fauna in cervids in Belarus was investigated and Trichuris ovis was identified in moose (Alces alces), roe deer (Capreolus capreolus) and red deer (Cervus elaphus). The intensity was low in the animals sampled and the prevalence of parasites in deer species were 33.3%, 37.5% and 37.5%, respectively [48]. In northern Poland, the prevalence in red deer and fallow deer was 1.9% and 3.64% respectively. However, Trichuris spp. was not detected in roe deer using morphological techniques. The maximum faecal egg load was 30 eggs per gram (EPG) [49]. Similar work was conducted in various countries and the prevalence in Turkey was 13.3% [50], 2.4% for Croatia [51] and 14.7% in Austria [52]. In the Iberian Peninsula, wild roe deer was found with T. capreoli and T. ovis in 53.1% and 10.4%, respectively, of the animals sampled. It was also noted that the prevalence of Trichuris was higher in males (59.4%) in comparison to females (22.6%) [53]. Free-ranging red deer was found to contain several endoparasites, one of which was Trichuris ovis. The authors showed that a low level of endoparasites caused reduction in the body condition of the animals [54].
In Norway, several studies have been done investigating the effect of endoparasites on body condition in moose (A. alces) [55] as well as the effect that supplemental feeding has on nematode infection [56]. The prevalence of Trichuris spp. was 2.2% [57] and 33.3% [56] with the level of endoparasites having an inverse relationship to body condition [57]. In addition, supplemental feeding had no impact on the level of endoparasites in the sampled animals [56]. Researchers also investigated the parasites in Norwegian red deer in an isolated reserve and found Trichuris globulosa in 30.8% of the samples, but the mean helminth count was relatively low [55].
In most of the investigations previously recorded, deer were usually hunted or reared semi-intensively with limited information on the health of the animal. In contrast, a study was done in Sweden investigating gastrointestinal parasitic infection in dead or debilitated moose (A. alces) [58]. Trichuris eggs were found in 38% of faecal samples but worms were found in 10% of caecum and 2% in the ascending colon [58] (See Table 2). Jokelainen et al. [59] reviewed gastrointestinal parasites in reindeer located in Fennoscandia (Finland, Sweden, Norway and Russia). It was stated that historically, the Trichuris spp. was not commonly found in reindeer or moose from Europe but recent evidence has shown that they can be found in these species. Recently, T. globulosa was found in 38.9% of roe deer that were sampled in Russia [60]. In summary, the prevalence of this parasite in European deer was quite variable. In some regions, the prevalence was high (>75%) whilst it was low (<2%) in others (See Table 2). Variability in the prevalence may be due to several reasons, such as the method of identification, geographical location and environmental factors such as season.

5. Trichuris in Deer Located in Canada

In Canada, the gastrointestinal tracts of wild moose (Alces alces) and elk (Cervus elaphus) were examined for endoparasites. These endoparasites were identified using morphological characteristics. Trichuris spp. were identified in the caecum of both the moose (n = 140) and the wapiti (n = 186) in 34% and 20% of the respective samples. Only female worms were identified and worms had characteristics similar to the vulva and uteri of T. ovis [70]. In eastern Ontario (Canada), two species of Trichuris were identified in wild moose (Alces alces) [71]. T. ovis and T. discolor were identified using morphological analysis and it was the first time the latter species had been identified in moose. The prevalence of T. ovis and T. discolor was 13% and 25%, respectively. The intensity of these parasites was not considered large enough to have affected the health of the animals. The two species of Trichuris identified have also been found in domestic ruminants and it should be mentioned that moose that were captured were present in a forest reserve in close proximity to agricultural areas [71]. Farmed and wild woodland caribou (Rangifer tarandus) in north-western Ontario (Canada) were found to have Trichuris spp. eggs in their faeces. Adult Trichuris ovis were discovered in the gastrointestinal tract of farmed woodland caribou [72]. The Atlantic-Gaspesie caribou (Rangifer tarandus caribou) is a small isolated population of an endangered species. Faecal samples were taken to assess the level of parasitism present in these animals. Trichuris eggs were found in 6% of the animals sampled with a low level of infection detected [73]. In summary, the prevalence of this parasite in Canadian deer was at an intermediate level (See Table 3).

6. Trichuris in Deer Located in Asia and Australia

The incidence of helminth infection was investigated in wild Axis deer in India. Direct smears of faecal samples were used to identify helminth ova. In this study, the prevalence of Trichuris spp. was 8.5% but the animals’ body condition was not recorded [74]. Prior to the published reported, McKenzie and Davidson [75] reported a 30% prevalence of Trichuris species in free-ranging Axis deer but these animals had low parasitic intensity and the sample size consisted of only ten animals. Upon post mortem examination, the animal was found to have large volumes of eggs in faeces based on faecal smears (prepared by direct method and faecal flotation) [76]. In Nepal, nine faecal samples were collected from the Musk deer (Moshus chrysogaster) and six samples were found to be positive for Trichuris spp. However, it must be stated that the small sample size of the above survey may bias the results. Most animals were recorded to have a light infection (<10%) of whipworms [77]. In China, the prevalence of endoparasites of musk deer at various locations and seasons were investigated [78]. The summer had the highest prevalence of Trichuris spp. (26.9%) with spring (13.6%) and winter (5.7%) having low values (see Table 4). With respect to location, the southern mountainous regions had higher values (31.1% and 35.0%) as compared to the eastern region (12.4%) [79]. In Indonesia, faecal sedimentation was applied to fifteen faecal samples from samba deer and seven faecal samples from spotted deer. The data showed only Trichuris spp. was found in 14.3% (n = 7) of spotted deer and none in the samba deer. The intensity of the infection in the spotted deer was categorised as light [80]. In New Zealand, the red deer (C. elaphus) was found to harbour Trichuris ovis and the fallow deer (D. dama) had an unidentified species of Trichuris [81]. Faecal samples were collected from captive wild ruminants in north eastern India. These ruminants included the barking deer, sika deer, mask deer, chital, brow-antlered deer and Himalayan serow. Of the samples taken, 1.77% (10/565) of ruminants were infected with Trichuris spp. [82]. A similar study was conducted on captive wild ruminants in the Punjab area. It revealed that most of the deer species were not infected with Trichuris spp. with only one spotted deer being infected (25%, n = 4) [83]. In summary, the prevalence of this parasite in Asian and Australian deer was quite variable. In some regions the prevalence was high (>70%) whilst it was low (<2%) in others (see Table 4). These results were similar to European deer.

7. Modern Techniques Used in Identifying Trichuris spp. in Cervids

At present, there is a dearth of information on the identification of Trichuris species in deer using molecular techniques. However, most of the molecular studies that were conducted were done with roe deer [86,87]. Trichuris species that were first found in roe deer used sequencing of the ITS1-5.8S-ITS2 ribosomal DNA fragment [86]. Comparisons were made between morphological and molecular techniques in the identification of this parasite. Interestingly, Trichuris globulosa and Trichuris ovis were identified using morphological techniques, but T. globulosa was identified as T. discolor using molecular techniques. T. ovis was confirmed using both methods (molecular and morphological) [86].
Further to this, a new molecular marker (ITS1-5.8S RNA-ITS2) was used to determine Trichuris spp. found in sheep (Ovis orientalis aries) and roe deer. T. ovis and T. discolor was found in both sheep and deer. T. ovis was predominantly present in sheep and T. discolor in roe deer [87]. Nechybova et al. [87] used the new molecular marker described previously to identify Trichuris species in wild ruminants in numerous localities in the Czech Republic. The ruminants investigated included roe deer, sika deer, red deer, fallow deer and mouflons, with T. discolor being the predominant trichurid and T. ovis identified less frequently [87]. The prevalence of T. discolor in roe deer, fallow deer, red deer and sika deer was 54.1%, 38.5%, 5.6% and 5.17%, respectively. T. ovis was not found in red deer but the prevalence for the other species of deer was 3.28% (roe deer), 7.69% (fallow deer) and 1.67% (sika deer) [87].
There are many simple techniques for the identification of gastrointestinal parasites. Examination of faeces grossly is not very sensitive, but in some cases adult helminths may be seen [4]. Microscopic examination of faeces and morphological analysis of adult worms are both insensitive in the determination of individual Trichuris spp. This is due to the variability noted in both egg and adult measurement of Trichuris spp. found in the same host. It must be noted that morphological techniques are needed to assess parasite burden. Some of the microscopic techniques include the use of direct faecal smears, faecal flotation and faecal sedimentation. Direct faecal smears are the most insensitive of the microscopic techniques. However, faecal smears allow for the identification of mobile protozoa. Faecal flotation and sedimentation are the most commonly used techniques. Faecal flotation allows eggs to float in solutions at specific gravity of more than 1.2 in most cases. Common faecal flotation solutions include zinc sulphate, sugar sulphate, sodium chloride (saturated solution). Some helminth eggs will not float, such as trematodes and in these cases, sedimentation techniques are more appropriate. Trichuris spp. eggs are easily identified using faecal flotation but some authors also used faecal sedimentation [4]. The use of faecal flotation as a tool to determine the prevalence of endoparasites may give false negative results in cases where the helminth population is immature (larval states) or consists of one sex (either males or females).
In most cases, Trichuris species were identified grossly using morphological tools. This method can be used as a screening test and there is a possibility of false negatives being reported. Faecal flotation and sedimentation were also used but molecular techniques are the gold standard for helminth identification. Therefore, the data presented in this review was done using microscopic and gross identification of helminth (adults and eggs) and there may be some under reporting of Trichuris-positive animals. Thus, the data described in this review should be used sparingly and future work should identify these helminths using molecular techniques to get a more accurate picture of the prevalence of this parasite at species level in various geographical regions.

8. Conclusions

The prevalence of the Trichuris spp. parasite in neo-tropical deer was low whilst in Europe, Asia, and Australia deer the prevalence was variable (2–80%). The prevalence in Canadian deer showed intermediary values (mid-range). Variability in the prevalence may be due to several reasons, such as the methods of identification, geographical location and environmental factors such as season. Morphological diagnosis (faecal flotation of eggs and adults) is not an accurate tool to determine the prevalence of these parasitic species and future studies should focus on molecular identification techniques.

9. Summary

This review revealed that Trichuris spp. were present in selected deer species. The parasite does not overtly cause clinical illness in wild or captive deer populations but the sub-clinical effects of this parasite are still unknown. Wild and captive-reared animals usually have a low prevalence and intensity of this parasite. The deer of neo-tropical origin have a low prevalence in comparison with other regions. In most cases, the identification technique used was based on morphometric features of eggs or adult helminths. These techniques were shown to be less accurate than modern molecular techniques. The species identified in this review appear to be T. skrjabini, T. odocoileus, T. ovis, T. globulosa, T. capreoli and T. discolor. Due to the use of morphological and biometric techniques in the identification of this parasite, it is difficult to state with any certainty which Trichuris species were present. This is because there has been variability in using morphological and biometric analysis of eggs and adults to determine individual species. Most of the studies failed to identify the species present in the samples and there was no published information on the molecular identification of Trichuris spp. with deer of neo-tropical origin. Some authors stated that the low prevalence and intensity of this parasite suggest it will not be transferred to domestic ruminants but this is a topic of major debate.

Funding

This work received no external funding.

Institutional Review Board Statement

This review didn’t involve the use of animals. Therefore, ethical approval was not required.

Data Availability Statement

Data supporting these results were presented in the article.

Conflicts of Interest

The author declares no conflict of interest.

References

  1. Jones, K.R.; Lall, K.R.; Garcia, G.W. Endoparasites of Selective Native Non-Domesticated Mammals in the Neotropics (New World Tropics). Vet. Sci. 2019, 6, 24. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Jones, K.R. Trichuris spp. in Animals with Specific Reference to Neo-Tropical Rodents. Vet. Sci. 2021, 8, 15. [Google Scholar] [CrossRef]
  3. Zajac, A.M.; Conboy, G.A. Veterinary Clinical Parasitology, 8th ed.; John Wiley & Sons Ltd.: Oxford, UK, 2012. [Google Scholar]
  4. Hendrix, C.M.; Robinson, E. Diagnostic Parasitology for Veterinary Technicians, 5th ed.; Elsevier Inc.: Amsterdam, The Netherlands, 2017. [Google Scholar]
  5. Knight, R.A. Trichuris odocoileus sp. N. (Nematoda: Trichuridae) from White Tailed Deer, Odocoileus virginianus, in South Eastern U.S., and a Key to Trichuris in North American Ruminants. J. Parasitol. 1983, 69, 1156–1159. [Google Scholar] [CrossRef] [PubMed]
  6. Samuel, W.M.; Beaudoin, R.L. Evaluation of Two Survey Methods for Detection of Helminth Infection in White Tailed Deer (Odocoileus virginianus). Bull. Wildl. Dis. Assoc. 1966, 2, 100–106. [Google Scholar] [CrossRef] [Green Version]
  7. Beaudoin, R.L.; Samuel, W.M.; Strome, C.P.A. A Comparative Study of the Parasites in two populations of White Tailed Deer. J. Wildl. Dis. 1970, 6, 56–63. [Google Scholar] [CrossRef]
  8. Heuer, D.E.; Phillips, J.H.; Rudersdorf, W.J.; Harley, P. Range Extention Records for Cooperia curticri, Ostertagia ostertagi, Setaria yehi and Trichuris ovis in White-Tailed Deer from Kentucky. Proc. Helminthol. Soc. Wash. 1975, 42, 141–142. [Google Scholar]
  9. Prestwood, A.K.; Pursglove, S.R.; Hayes, F.A. Parasitism among White-Tailed Deer and Domestic Sheep on Common Range. J. Wildl. Dis. 1976, 12, 380–385. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  10. Matsubayashi, M.; Kita, T.; Narushima, T.; Kimata, I.; Tani, H.; Sasai, K.; Baba, E. Coprological Survey of Parasite Infection in Pigs and Cattle in Slaughterhouse Osaka, Japan. J. Vet. Med. Sci. 2009, 71, 1073–1083. [Google Scholar] [CrossRef] [Green Version]
  11. Singh, N.K.; Singh, H.; Haque, J.M.; Rath, S.S. Prevalence of Parasitic Infections in Cattle of Ludhiana District, Punjab. J. Parasit. Dis. 2012, 36, 256–259. [Google Scholar] [CrossRef] [Green Version]
  12. Jimenez, A.E.; Montenegro, V.M.; Hernandez, J.; Dolz, G.; Maranda, L.; Galindo, J.; Epe, C.; Schnieder, T. Dynamics of Infections with Gastrointestinal Parasites and Dictyocaulus viviparus in Dairy and Beef Cattle from Costa Rica. Vet. Parasitol. 2007, 148, 262–271. [Google Scholar] [CrossRef]
  13. Woodbury, M.R.; Wagner, B.; Ben-Ezra, E.; Douma, D.; Wilkins, W. A Survey to Detect Toxocara Vitulorum and Other Gastrointestinal Parasites in Bison (Bison Bison) Herds from Manitoba and Saskatchewan. Can. Vet. J. 2014, 55, 870–874. [Google Scholar]
  14. Nwosu, C.O.; Madu, P.P.; Richards, W.S. Prevalence and Seasonal Changes in the Population of Gastrointestinal Nematodes of Small Ruminants in the Semi-Arid Zone Od North Eastern Nigeria. Vet. Parasitol. 2007, 144, 118–124. [Google Scholar] [CrossRef] [PubMed]
  15. Gul, N.; Tak, H. Prevalence of Trichuris spp. in Small Ruminants in Srinagar District (J & K). J. Parasit. Dis. 2016, 40, 741–744. [Google Scholar]
  16. Sousa, M.F.; Pimental-Netu, M.; da Silva, R.M.; Farias, A.C.B.; Guimaraes, M.P. Gastrointestinal Parasites of Sheep, Municipality of Lajes, Rio Grande Do Norte, Brazil. Rev. Bras. Parasitol. Vet. 2012, 21, 71–73. [Google Scholar] [CrossRef] [Green Version]
  17. Yevstafieva, V.A.; Yuskiv, I.D.; Melnychuk, V.V.; Yasnolob, I.O.; Kovalenko, V.A.; Horb, K.O. Nematodes of the Genus Trichuris (Nematoda, Trichuridae), Parasitizing Sheep in the Central and South-Eastern Regions of Ukraine. Vestnik. Zool. 2018, 52, 193–204. [Google Scholar] [CrossRef] [Green Version]
  18. Knight, R.A.; Tuff, D.W. Trichuris spp. (Nematoda: Trichuridae) in Sika Deer (Cervus nippon) in Texas. Proc. Helminthol. Soc. Wash. 1985, 51, 161–162. [Google Scholar]
  19. Cook, T.W.; Ridgeway, B.T.; Andrews, R.; Hodge, J. Gastro-Intestinal Helminths in White-Tailed Deer (Odocoileus virginianus) of Illinois. J. Wildl. Dis. 1979, 15, 405–408. [Google Scholar] [CrossRef] [Green Version]
  20. Haigh, J.; Berezowski, J.; Woodbury, M. A Cross-Sectional Study of the Causes of Morbidity and Mortality in Farmed White-Tailed Deer. Can. Vet. J. 2005, 46, 507–512. [Google Scholar]
  21. Sianto, L.; Duarte, A.N.; Charme, M.; Magalhaes, J.; Sousa, M.V.; Ferreira, L.F.; Araujo, A. Trichuris sp. from 1040 +/− 50-Year Old Cevidae Coprolites from Archaeological Site Furna Do Estrago, Pernambuco, Brazil. Mem. Inst. Oswaldo Cruz 2012, 107, 273–274. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Beltrame, M.O.; Tietze, E.; Perez, A.E.; Bellusci, A.; Sardella, N.H. Ancient Parasites form Endemic Deer from “CUEVA PARQUE DIANA” Archeological Site, Pantagonia, Argentina. Parasitol. Res. 2017, 116, 1523–1531. [Google Scholar] [CrossRef] [PubMed]
  23. Deem, S.L.; Noss, A.J.; Villarroel, R.; Uhart, M.M.; Karesh, W.B. Disease Survey of Free-Ranging Grey Brocket Deer (Mazama gouazoubira) in the Gran Chaco, Bolivia. J. Wildl. Dis. 2004, 40, 92–98. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  24. Lux Hoppe, E.G.; Tebaldi, J.H.; Nascimento, A.A. Helminthological Screening of Free-Range Grey Brocket Deer Mazama gouazoubira Fischer, 1817 (Cervidae: Odocoileini) from Brazilian Pentanal Wetlands, with Consideration on Pygarginema verrucosa (Molin, 1860) Kadenatzii, 1948 (Spirocercidae: Ascaropsinae). Braz. J. Biol. 2010, 70, 417–423. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. Nettles, V.F.; Quist, C.F.; Lopez, R.R.; Wilmers, T.J.; Frank, P.; Roberts, W.; Chitwood, S.; Davidson, W.R. Morbidity and Mortality Factors in Key Deer (Odocoileus virginianus clavium). J. Wildl. Dis. 2002, 38, 685–692. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  26. Montes Pérez, R.C.; Rodríguez Vivas, R.I.; Torres Acosta, J.F.; Ek Pech, L.G. Annual Monitoring of Gastrointestinal Parasitosis of White-Tailed Deer Odocoileus virginianus (Artiodactyla: Cervidae) in Captivity in Yucatán, México. J. Trop. Biol. 1998, 46, 821–827. [Google Scholar]
  27. Hernandez, Z.; Gonzalez, S. Parasitological Survey of the Uraguayan Population of Wild Pampas Deer (Ozotoceros bezoarticus L. 1758). Anim. Prod. Sci. 2012, 52, 781–785. [Google Scholar] [CrossRef]
  28. Carreno, R.A.; Durden, L.A.; Brooks, D.R.; Abrams, A.; Hoberg, E.P. Parelaphostrongylus Tenuis (Nematoda: Protostrongylidae) and Others Parasites of White-Tailed Deer (Odocoileus virginianus) in Costa Rica. Comp. Parasitol. 2001, 68, 177–184. [Google Scholar]
  29. Richardson, M.L.; Demarais, S. Parasites and Condition of Coexisting Populations of White-Tailed and Exotic Deer. J. Wildl. Dis. 1992, 29, 485–489. [Google Scholar] [CrossRef]
  30. Stubblefield, S.S.; Pence, D.B.; Warren, R.J. Visceral Helminth Communities of Sympatric Mule and White-Tailed Deer from the Davis Mountains of Texas. J. Wildl. Dis. 1987, 23, 113–120. [Google Scholar] [CrossRef] [Green Version]
  31. Waid, D.D.; Pence, D.B.; Warren, R.J. Effect of Season and Physical Condition on the Gastrointestinal Helminth Community of White-Tailed Deer from the Texas Edward Plateau. J. Wildl. Dis. 1985, 21, 264–273. [Google Scholar] [CrossRef]
  32. Davidson, W.R.; Crow, C.B. Parasites, Diseases and Health Status of Sympatric Populations of Sika Deer and White-Tailed Deer in Maryland and Virginia. J. Wildl. Dis. 1983, 19, 345–348. [Google Scholar] [CrossRef] [Green Version]
  33. Glazner, W.C.; Knowlton, F.F. Endoparasites Found in Welder Refuge Deer. J. Wildl. Manag. 1967, 31, 595–597. [Google Scholar] [CrossRef]
  34. Davidson, W.R.; Crum, J.M.; Blue, J.L.; Sharp, D.W.; Phillips, J.H. Parasites, Diseases and Health Status of Sympatric Populations of Fallow Deer and White-Tailed Deer in Kentucky. J. Wildl. Dis. 1985, 21, 153–159. [Google Scholar] [CrossRef] [Green Version]
  35. Wunschmann, A.; Armien, A.G.; Butler, E.; Schrage, M.; Stomberg, B.; Bender, J.B.; Firshman, A.M.; Carstensen, M. Necropsy Findings in 62 Opportunistically Collected Free-Ranging MOOSE (Alces alces) in Minnesota, USA (2003–13). J. Wildl. Dis. 2015, 51, 157–165. [Google Scholar] [CrossRef]
  36. Mukul-Yerves, J.M.; Zapata-Escobedo, M.R.; Montes-Perez, R.C.; Rodriguez-Vivas, R.I.; Torres-Acosta, J.F. Gastrointestinal and Ectoparasites in Wildlife-Ungulates under Captive and Free Living Conditions in the Mexican Tropic. Rev. Mex. Cienc. Pecu. 2014, 5, 459–469. [Google Scholar] [CrossRef] [Green Version]
  37. Morse, B.W.; Miller, D.L.; Miller, K.V.; Baldwin, C.A. Population Health of Fallow Deer (Dama dama) on Little St. Simon island, Geogria, USA. J. Wildl. Dis. 2009, 45, 411–421. [Google Scholar]
  38. Davidson, W.R.; Blue, J.L.; Flynn, L.B.; Shea, S.M.; Marchinton, R.L.; Lewis, J.A. Parasites, Diseases and Health Status of Sympatric Populations of Sambar Deer and White Tailed-Deer in Florida. J. Wildl. Dis. 1987, 23, 267–272. [Google Scholar] [CrossRef]
  39. Samuel, W.M.; Beaudoin, R.L. Identification of Eggs and Larvae of Nematodes Parasitic in Deer in Pennsylvania; Penn State University Press: University Park, PA, USA, 1966; Volume 39, pp. 73–77. [Google Scholar]
  40. Foreyt, W.J.; Trainer, D.O. Parasitism Changes in 2 Populatons of White-Tailed Deer in Wisconsin. J. Wildl. Manag. 1980, 44, 758–764. [Google Scholar] [CrossRef]
  41. Samuel, W.M.; Trainer, D.O. A Technique for Survey of Some Helminth and Protozoan Infection of White-Tailed Deer. J. Wildl. Manag. 1969, 33, 888–894. [Google Scholar] [CrossRef]
  42. Vetyska. V. Endoparasites of Roe Deer in the Strakonice Region. Acta Vet. Brno. 1980, 49, 91–103. [Google Scholar] [CrossRef] [Green Version]
  43. Sleeman, D.P. Parasites of Deer in Ireland. J. Life Sci. Dubl. Soc. 1983, 4, 203–210. [Google Scholar]
  44. Segonds-Pichon, A.; Ferte, H.; Gaillard, J.M.; Lamarque, F.; Duncan, P. Nematode Infestation and Body Condition in Roe Deer (Capreolus capreolus). Game Wildl. Sci. 2000, 17, 241–258. [Google Scholar]
  45. Body, G.; Ferte, H.; Gaillard, J.-M.; Delorme, D.; Klein, F.; Gilot-Fromont, E. Population Density and Phenotypic Attributes Influence the Level of Nematode Parasitism in Deer. Oecologia 2011, 167, 635–646. [Google Scholar] [CrossRef] [PubMed]
  46. Vengust, G.; Bidovec, A. Parasites of Fallow Deer (Dama dama) in Slovenia. Helminthologia 2003, 40, 161–164. [Google Scholar]
  47. Goossens, E.; Vercruysse, J.; Boomker, J.; Vercammen, F.; Dorny, P. A 12-Month Survey of Gastrointestinal Helminth Infections of Cervids Kept in Two Zoos in Belgium. J. Zoo Wildl. Med. 2005, 36, 470–478. [Google Scholar] [CrossRef] [PubMed]
  48. Shimalov, V.V.; Shimalov, V.T. Helminth Fauna of Cervids in Belorussian Polesie. Parasitol. Res. 2003, 89, 75–76. [Google Scholar]
  49. Burlinski, P.; Janiszewski, P.; Kroll, A.; Gonkowski, S. Parasitofauna in the Gastrointestinal Tract of the Cervids (Cervidae) in Northern Poland. Acta Vet. 2011, 61, 269–282. [Google Scholar] [CrossRef]
  50. Bolukbas, C.S.; Gurler, A.T.; Beyhan, Y.E.; Acici, M.; Umar, S. Helminths of Roe Deer (Capreolus capreolus) in the Middle Black Sea Region of Turkey. Parasitol. Int. 2012, 61, 729–730. [Google Scholar] [CrossRef]
  51. Kusak, R.R.; Spicic, S.; Slijepcevic, V.; Bosnic, S.; Janje, R.R.; Duvnjak, S.; Sindicic, M.; Majnaric, D.; Cvetnic, Z.; Huber, D. Health Status of Roe and Red Deer in Gorski kotar, Croatia. Vet. Arh. 2012, 82, 59–73. [Google Scholar]
  52. Rehbein, S.; Visser, M.; Jekel, I.; Silaghi, C. Endoparasites of Fallow Deer (Dama dama) of the Antheringer Au in Salzburg, Austria. Weir. Kin. Wochenschr. 2014, 126, 37–41. [Google Scholar] [CrossRef]
  53. Pato, F.J.; Vazquez, L.; Diez-Banos, N.; Lopez, C.; Sanchez-Andrade, R.; Fernandez, G.; Diez-Banos, P.; Panadero, R.; Diaz, P.; Morrondo, P. Gastrointestinal Nematode Infections in Roe Seer (Capreolus capreolus) from the NW of the Iberian Peninsula: Assessment of Some Risk Factors. Vet. Parasitol. 2013, 196, 136–142. [Google Scholar] [CrossRef]
  54. Irvine, R.J.; Corbishley, H.; Pilkington, J.G.; Albon, S.D. Low-Level Parasitic Worm Burdens May Reduce Body Condition in Free Ranging Red Deer (Cervus elaphus). Parasitology 2006, 133, 465–475. [Google Scholar] [CrossRef] [PubMed]
  55. Davidson, R.K.; Kutz, S.; Madsilen, K.; Hoberg, E.; Handeland, K. Gastrointestinal Parasites in an Isolated Norwegian Population of Wild Red Deer (Cervus elaphus). Acta Vet. Scand. 2014, 56, 59. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Milner, J.M.; Wedul, S.J.; Laaksonen, S.; Oksanen, A. Gastrointestinal Nematodes of Moose (Alces alces) in Relation to Supplementary Feeding. J. Wildl. Dis. 2013, 49, 69–79. [Google Scholar] [CrossRef] [Green Version]
  57. Davidson, R.K.; Licina, T.; Gorini, L.; Milner, J.S. Endoparasites in a Norwegian Moose (Alces alces) Population—Faunal Diversity Abundance and Body Condition. IJP Parasites Wildl. 2015, 4, 29–36. [Google Scholar] [CrossRef] [Green Version]
  58. Grandi, G.; Uhlhorn, H.; Agren, E.; Morner, T.; Righi, F.; Osterman-Lind, E.; Neimanis, A. Gastrointestinal Parasitic Infection in Dead or Debilitated Mose (Alces alces) in Sweden. J. Wildl. Dis. 2018, 54, 165–169. [Google Scholar] [CrossRef]
  59. 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] [PubMed]
  60. Kuznetsov, D.N.; Romashova, N.B.; Ramashov, B.V. Gastrointestinal Nematodes of European Roe Deer (Capreolus capreolus) in Russia. Russ. J. Theriol. 2020, 19, 85–93. [Google Scholar] [CrossRef]
  61. Tomczuk, K.; Szczepaniak, K.; Grzybek, M.; Studzinska, M.; Demkowska-Kutrzepa, M.; Lopuszynski, W.; Junkuszew, A.; Gruszecki, T.; Dudko, P.; Bojar, W. Internal Parasites in Roe Deer of Lubartow Forest Division in Post Mortem Studies. Med. Weter. 2017, 73, 726–730. [Google Scholar]
  62. Galecki, R.; Sokol, R.; Koziatek, S. Parasites of Wild Animals as a Potential Source of Hazard to Humans. Ann. Parasitol. 2015, 61, 105–108. [Google Scholar]
  63. Rehbein, S.; Lutz, W.; Visser, M.; Winter, R. Contributions to the Knowledge of the Parasite Fauna of the Wild in North Rhine-Westphalia. 1. The Endoparasite Infestation of the Roe Deer. J. Hunt. Sci. 2000, 46, 248–269. [Google Scholar]
  64. Filip-Hutsch, K.; Czopowicz, M.; Barc, A.; Demiaszkiewicz, A.W. Gastrointestinal Helminths of a European Moose Population in Poland. Pathogens 2021, 10, 456. [Google Scholar] [CrossRef]
  65. Popiołek, M.; Jarnecki, H.; Luczynski, T.; Macała, K.; Jagła, E. Endoparasites of Roe Deer (Capreolus capreolus L.) from Henryków Forest Inspectorate (Lower Silesia) Based on Faecal Analysis. Zesz. Nauk. Uniw. Przyr. We Wrocławiu-Biol. I Hod. Zwierząt 2009, 58, 139–149. [Google Scholar]
  66. Barth, D.; Matzke, P. Gastrointestinal Nematodes of Fallow Deer (Dama dama L.) in Germany. Vet. Parasitol. 1984, 16, 173–176. [Google Scholar] [CrossRef]
  67. Pato, F.J.; Díaz, P.; Panadero, R.; Díez-Baños, N.; Painceira, A.; López, C.; Díez-Baños, P.; Fernández, G.; Morrondo, P. Prevalence and Intensity of Infection by Gastrointestinal Nematodes in Roe Deer (Capreolus Capreolus): Differences According to the Sex. Mappe Parassitiologiche 2009, 18, 141. [Google Scholar]
  68. Pato, F.J.; Vázquez, L.; Painceira, A.; Díaz, P.; Uriarte, J.; Diez-Baños, N.; Dacal, V.; López, C.; Panadero, R.; Díez-Baños, P.; et al. Gastrointestinal nematode species shared by Roe Deer (Capreolus capreolus) and Grazing Cattle from Galicia. In Proceedings of the 13th Conference on Animal Production, Zaragoza, Spain, 12–13 May 2009; pp. 176–178. [Google Scholar]
  69. Vázquez, L.; Painceira, A.; Dacal, V.; Pato, F.J.; Panadero, R.; López, C.; Díaz, P.; Arias, M.S.; Francisco, I.; Díez-Baños, P.; et al. Long-Term Study of Internal Parasitic Infections in Free-Ranging Roe Deer (Capreolus capreolus) from NW Spain. Rev. Ibero-Latinoam. Parasitol. 2010, 69, 172–177. [Google Scholar]
  70. Stock, T.M.; Barrett, M.W. Helminth Parasites of the Gastrointestinal Tracts and Lung of Moose (Alces alces) and Wapiti (Cervus elaphus) from Cypress Hills, Alberta, Canada. Proc. Helminthol. Soc. Wash. 1983, 50, 246–251. [Google Scholar]
  71. Hoeve, J.; Joachim, D.G.; Addison, E.M. Parasites of Moose (Alces alces) from an Agricultural Area of Eastern Ontario. J. Wildl. Dis. 1988, 24, 371–374. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  72. 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]
  73. 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-Gaspesie Caribou. IJP: Parasites Wildl. 2018, 7, 90–94. [Google Scholar] [CrossRef]
  74. Meshram, M.D.; Shirale, Y.S.; Khillare, K.P. Incidence of Helminth infection in Axis deer. Vet. World 2008, 1, 10. [Google Scholar]
  75. McKenzie, M.E.; Davidson, W.R. Helminth Parasites on Intermingling Axis Deer, Wild Swine and Domestic Cattle from the Island of Molokai, Hawaii. J. Wildl. Dis. 1989, 25, 252–257. [Google Scholar] [CrossRef] [Green Version]
  76. Swain, K.; Vohra, S.; Gupta, S.; Routray, A.; Panigrahi, S.; Sahoo, S.; Ganguly, S. Trichuris Infection in the Spotted Deer (Axis axis) at Hispar (Haryana). J. Immunol. Immunopathol. 2017, 19, 107–109. [Google Scholar] [CrossRef]
  77. Achhami, B.; Sharma, H.P.; Bam, A.B. Gastrointestinal Parasites of Musk Deer (Moschus chrysogaster Hodgson, 1839) in Langtang National park, Nepal. JIST 2016, 2, 71–75. [Google Scholar]
  78. Hu, X.-L.; Liu, G.; Wei, Y.-T.; Wang, Y.-H.; Zhang, T.-X.; Yang, S.; Hu, D.-F.; Liu, S.-Q. Regional and Seasonal Effect of the Gastrointestinal Parasitism in Captive Forest Musk Deer. Acta Trop. 2018, 117, 1–8. [Google Scholar] [CrossRef] [PubMed]
  79. Tanjung, M.; Sibarani, H.L. Species and Prevalence of Endoparasites on the Feces of Sambar Deer (Cervus unicolor) and Spotted Deer (Axis axis) in Conservation Universitas Sumatera Utara. J. Phys. Conf. Ser. 2018, 1116. [Google Scholar] [CrossRef]
  80. Mc Kenna, P.B. Checklist of Helminth Parasites of Terrestrial Mammals in New Zealand. N. Z. J. Zool. 1997, 24, 277–290. [Google Scholar] [CrossRef]
  81. Salaba, O.; Rylkova, K.; Vadlejch, J.; Petrtyl, M.; Schankova, S.; Brozova, A.; Jebavy, L.; Langrova, I. The First Determination of Trichuris sp. from Roe Deer by Amplification and Sequenation of the ITS-5.8S-ITS2 Segment of Ribosomal DNA. Parasitol. Res. 2013, 112, 955–960. [Google Scholar] [CrossRef] [PubMed]
  82. Patra, G.; Efimova, M.A.; Sahara, A.; Borthakur, S.K.; Ghosh, S.; Behera, G.P.; Polley, S.; Debbarma, A. Incidence of Ecto- and Endo-Parasitic Fauna in Small Wild Ruminants from North Eastern Region of India. Biol. Rhythm. 2019. [Google Scholar] [CrossRef]
  83. Mir, A.Q.; Dua, K.; Singla, L.D.; Sharma, S.; Singh, M.P. Prevalence of Parasitic Infection in Captive Wild Animals in Bir Moti Bagh Mini Zoo (Deer Park), Patiala, Punjab. Vet. World 2016, 9, 540–543. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  84. Rana, M.A.; Jabeen, F.; Shabnab, M.; Ahmad, I.; Hassan, M.M. Comparasitve Study of Endoparasites in Captive Hog Deer (Axis porcinus). Int. J. Biosci. 2015, 6, 162–170. [Google Scholar]
  85. Abdybekova, A.M.; Sultanov, A.; Dzhusupbekova, G.D.; Torgerson, P.R. Parasites of Farmed Marals in Kazakhstan. Small Rumin. Res. 2017, 153, 142–145. [Google Scholar] [CrossRef]
  86. Vejl, P.; Nechybova, S.; Perinkova, P.; Melounova, M.; Sedlakova, V.; Vasek, J.; Cilova, D.; Rylkova, K.; Jankovska, I.; Vadlejch, J.; et al. Reliable Molecular Differentiation of Trichuris ovis and Trichuris discolor from Sheep (Ovis orientalis aries) and Roe Deer (Capreolus capreolus) and Morphological Characterization of Their Females: Morphology Does Not Work Sufficiently. Parasitol. Res 2017, 10.1007/s00436-017-5524-9. [Google Scholar] [CrossRef] [PubMed]
  87. Nechybova, S.; Vejl, P.; Hart, V.; Melounova, M.; Cilova, D.; Vasek, J.; Jankovska, I.; Vadlejch, J.; Langrova, I. Long-Term Occurrence of Trichuris Species in Wild Ruminants in the Czech Republic. Parasitol. Res. 2018, 117, 1699–1708. [Google Scholar] [CrossRef] [PubMed]
Table
Table 1. Prevalence of Trichuris in neo-tropical deer species.
Table 1. Prevalence of Trichuris in neo-tropical deer species.
HostParasiteMethod of
Identification		HabitatPrevalence% (x/y)Refs.
Odocoileus virginianus (White-tailed deer)Trichuris ovisMorphology of adult and faecal flotationFree range3.3 (4/120)[6]
Odocoileus virginianus (White-tailed deer)Trichuris ovisMorphology of adult and faecal flotationFree range3.4 (4/117)[7]
Odocoileus virginainus (White-tailed deer)Trichuris ovisMorphology of adultFree range16 (5/31)[8]
Odocoileus virginianus (White-tailed deer)Trichuris spp.Morphology of adultFree range9.1 (4/44)[19]
Mazama gouazoubira (Grey Brocket deer)Trichuris ovisFaecal flotationFree range9.1 (1/11)[23]
Alces alces (Moose)Trichuris spp.Morphology of adultFree range4.83 (3/62)[35]
Mazama amerciana (Red Brocket deer)Trichuris spp.Faecal flotationFree range31.3 (5/16)[36]
Fallow deer
(Dama dama)		Trichuris spp.Faecal flotationFree range1/68 (1.15)[37]
Odocoileus hemionus (Mule deer)Trichuris discolorMorphology of adultFree range4 (1/25)[30]
Odocoileus virginianus (White-tailed deer)Trichuris spp.Faecal flotationFree range4 (7/28)[29]
Rusa unicolor
(Samba deer)		Trichuris spp.Faecal flotationFree range20 (2/10)[38]
Odocoileus virginianus (White-tailed deer)Trichuris spp.Faecal flotationFree range17 (1/6)
Odocoileus virginianus (White-tailed deer)Trichuris ovisFaecal flotationFree range6.49 (5/77)[39]
Odocoileus virginianus (White-tailed deer)Trichuris spp.Faecal flotationFree range5.26 (4/76)[40]
Morphology of adult1.32 (1/76)
Odocoileus virginianus (White-tailed deer)Trichuris ovisFaecal flotationFree range3.82 (26/681)[41]
Morphology of adult7.14 (1/14)
Table
Table 2. Prevalence of Trichuris in European deer species.
Table 2. Prevalence of Trichuris in European deer species.
HostParasiteMethod of
Identification		HabitatPrevalence% (x/y)Refs.
Capreolus capreolus (Roe deer)Trichuris capreolusMorphology of adult and faecal flotationFree range (Czech)8.9% (10/112)[42]
Trichuris globulosa21.4% (24/112)
Rangifer tarandus (Reindeer)Trichuris spp.Morphology of adult and faecal flotationCaptive reared (Belgium)25% (1/4)[47]
Cervus elaphus (Red deer)Trichuris spp.Faecal flotationFree range (Northern Poland)1.8% (9/500)[49]
Dama dama (Fallow deer)3.6% (16/440)
Alces alces (Moose)Trichuris ovisFaecal flotationFree range (Belarus)33.3% (6/18)[48]
Capreolus capreolus (Roe deer)37.5% (6/16)
Cervus elaphus (Red deer)31.3% (5/16)
Dama dama (Fallow deer)Trichuris capreoliFaecal flotationFarmed (Slovenia)2.3% (10/43)[46]
Trichuris ovis2.3% (10/43)
Cervus elaphus (Red deer)Trichuris spp.Morphology of adult and faecal flotationFree range (Croatia)2.4% (10/41)[51]
Capreolus capreolus (Roe deer)0% (0/25)
Capreolus capreolus (Roe deer)Trichuris ovisMorphology of adultFree range (Turkey)13.3% (2/15)[50]
Capreolus capreolus (Roe deer)Trichuris capreoliMorphology of adultFree range (Iberian Peninsula)53.1% (116/218)[53]
Trichuris ovis10.5% (23/218)
Dama damaTrichuris globulosaMorphology of adultFree range (Austria)14.3% (1/7)[52]
Alces alces (Moose)Trichuris spp.Faecal flotationFree range (Norway)33% (8/24)[56]
Cervus elaphus (Red deer)Trichuris globulosaMorphology of adult and faecal flotation30.8% (4/13)[55]
Alces alces (Moose)Trichuris spp.Faecal flotation2.2% (1/45)[57]
Alces alces (Moose)Trichuris spp.Morphology of adultFree range (Sweden)Caecum [10% (10/50)]
Rectum [2% (1/50)]		[58]
Faecal flotation3.8% (2/50)
Capreolus capreolus (Roe deer)Trichuris globulosaMorphology of adultFree range (Russia)38.9% (7/18)[60]
Capreolus capreolus (Roe deer)Trichuris globulosaMorphology of adultFree range (Poland)9.43% (5/53)[61]
Capreolus capreolus (Roe deer) and Dama dama (Fallow deer)Trichuris spp.Faecal flotationFree range (Poland)10.48% (11/105)[62]
Capreolus capreolus (Roe deer)Trichuris spp.Faecal flotationFree range (German)7.8% (5/64)[63]
Trichuris globulosaMorphology of adult67.2% (43/64)
Trichuris ovisMorphology of adult4.7% (3/64)
Alces alces (Moose)Tricuris spp.Morphology of adultFree range (Poland)83.3% (10/12)[64]
Faecal flotation68.9% (137/199)
Capreolus capreolus (Roe deer)Trichuris spp.Faecal flotationFree range (Southwest Poland)9.2% (12/131)[65]
Dama dama (Fallow deer)Triichuris capreoliMorphology of adultFree range (German)20% (10/49)[66]
Trichuris ovis2% (1/49)
Capreolus capreolus (Roe deer)Trichuris spp.Morphology of adultFree range (Galacia, Spain)59.4% (111/187) male[67]
22.6% (7/31) female
Capreolus capreolus (Roe deer)Trichuris spp.Faecal flotationFree range (Galacia, Spain)47.2% (173/367)[68]
Capreolus capreolus (Roe deer)Trichuris spp.Faecal flotationFree range (North-western Spain)3% (4/128)[69]
4% (15/367)
Table
Table 3. Prevalence of Trichuris in Canadian deer species.
Table 3. Prevalence of Trichuris in Canadian deer species.
HostParasiteMethod of IdentificationHabitatPrevalence% (x/y)Refs.
Alces alces (Moose)Trichuris spp.Morphology of adultFree range34% (48/140)[70]
Cervus elaphus (Red deer)20% (37/186)
Alces alces (Moose)Trichuris ovisMorphology of adultFree range13% (2/16)[71]
Trichuris discolor25% (4/16)
Rangifer tarandus (Reindeer)Trichuris ovisMorphology of adult and Faecal flotationFree range40% (2/5)[72]
Rangifer tarandus caribou (Caribou)Trichuris spp.Faecal flotationFree range6% (2/32)[73]
Table
Table 4. Prevalence of Trichuris in Axis, spotted, Samba and musk Deer.
Table 4. Prevalence of Trichuris in Axis, spotted, Samba and musk Deer.
HostParasiteMethod of
Identification		HabitatPrevalence% (x/y)Refs.
Cervus axis (Spotted deer)Trichuris spp.Morphology of adultFree range30 (3/10)[75]
Cervus axis (Spotted deer)Trichuris spp.Faecal flotationFree range8.5 (17/200)[75]
Moschus chrysogaster (Alpine musk deer)Trichuris spp.Faecal flotationFree range66.7 (6/9)[77]
Cervus unicolor (Sambar deer)Trichuris spp.Faecal sedimentationFree range0 (0/15)[78]
Axis chrysobactin (Spotted deer)14.28 (1/7)
Moschus chrysogaster (Alpine musk deer)Trichuris spp.Faecal flotationFree rangeSpring 13.6 (6/44)
Summer 24.5 (53/216)
Winter 5.7 (4/70)		[79]
Axis chrysobactin (Hog deer)Trichuris globulosaFaecal flotationCaptive reared74.4 (64/86)[84]
Cervus elaphus (Moral)Trichuris skrjabiniFaecal flotationCaptive1.98 (10/505)[85]
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Jones, K.R. Trichuriasis in Selected Deer (Cervidae) Species: A Geographical Perspective. Ruminants 2021, 1, 178-190. https://doi.org/10.3390/ruminants1020013

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Jones KR. Trichuriasis in Selected Deer (Cervidae) Species: A Geographical Perspective. Ruminants. 2021; 1(2):178-190. https://doi.org/10.3390/ruminants1020013

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Jones, Kegan Romelle. 2021. "Trichuriasis in Selected Deer (Cervidae) Species: A Geographical Perspective" Ruminants 1, no. 2: 178-190. https://doi.org/10.3390/ruminants1020013

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