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Goat farming is still in its infancy in Ireland. The purpose of this study was to survey goat farmers on their knowledge and current control practices of parasites as well as some information on the specific gastrointestinal parasite fauna of goats in Ireland. The main farming enterprise was cheese (46%) production and the majority of farms (55%) practising a pasture-based grazing system. Anthelmintics was regularly used on all farms with visual appraisal most commonly used (73%) to calculate animal weight. Anthelmintic dose given ranged from the recommended sheep dose (22%) to twice the recommended sheep dose (33%). A variety of different nematode, trematode, cestode and protozoa parasites were detected in the pooled samples. The prevalence of Eimeria spp. was the highest ranging from 79 and 100% in the adult goats and kids, respectively. Prevalence of trematodes ranges from 3 to 17% in adult goats, while none were detected in the kids. Of the gastrointestinal nematodes, strongyle eggs were most prevalent in adult (69%) and kids (42%). This first report on endoparasitic infections of goats in Ireland reveals a high prevalence of endoparasites and that farmers still uses inappropriate and unsustainable parasite control practices, highlighting the need for further education.
School of Veterinary Medicine, University College Dublin, Dublin, Ireland
Laura Rinaldi
Department of Veterinary Medicine and Animal Production, University of Naples Federico II, Naples, Italy
*Address all correspondence to: theo.dewaal@ucd.ie
1. Introduction
Goat farming in Ireland is still very much in its infancy, with only a small number of goat farms with the main emphasis on milk and cheese production. During the latter years of the last decade, there was a major influx of new entrants into the goat industry in Ireland, and data from the 2021 National Sheep and Goat Census (https://assets.gov.ie/226265/1598c686-c575-414d-9719-33ff1ff3c72d.pdf) recorded 735 registered herds representing a total number of 8077 goats. However, by far the majority (66.5%; n = 489) of the herds are very small, <5 goats/herd, and only 13 herds (1.8%) reported herd sizes of >100, representing 3051 (37.8%) goats.
Gastrointestinal helminths (GIH) affect the health and productivity of goats on pasture worldwide, and the frequent use of anthelmintic drugs is still the most common method used for controlling infection. This has led to the widespread development of anthelmintic resistance [1]. Resistance to two or more anthelmintic drug classes has been extensively documented in goat nematodes worldwide [1].
Although the GIH infecting sheep and goats are very similar the majority of studies on host–parasite interactions have been conducted in sheep and extrapolated to goats, but this may not reflect the true picture [2]. For example, the immune response in goats against GIH is less efficient in limiting helminth populations [2, 3, 4, 5]. Studies have also shown that, in dairy goats in particular, there are similar GIH infection levels between adult and young animals, whereas in sheep adult ewes are usually much less heavily infected [5, 6]. Also, goats seem to be less able to control challenge infections compared to sheep, probably because the “immune memory” in goats does not last as long as in sheep [3, 7]. Therefore, an understanding of the GIH infecting goats is essential to improve the productivity and control of GIH in grazing goats. Furthermore, in many jurisdictions, the registration of anthelmintic drugs does not discriminate between sheep and goats, but several studies have shown that goats, due to their higher metabolic rate, require a higher drug dose to maintain the same plasma concentrations and half-life as other similar species [6, 7, 8, 9]. The route by which anthelmintic may also be important in goats as one study have shown that macrocyclic lactone drugs are most effective in goats when administered orally [10].
Several surveys on helminths and anthelmintic resistance have been conducted in sheep flocks in Ireland [11, 12, 13]. However, very little is known about helminths and anthelmintic usage in Irish goat farms. This study aimed to survey goat farmers on their knowledge and current control practices of parasites as well as some information on the specific GI parasite fauna of goats in Ireland.
Goat owners that were members of Teagasc, the Agriculture and Food Development Authority of Ireland, were invited to participate in this study. Owners who confirmed their participation were sent a questionnaire and faecal collection kit with instructions on how to collect and send samples to the University College Dublin Parasitology laboratory (UCD PL). The questionnaire consisted of 27 questions and was divided into four parts, (i) basic farm information, (ii) goat herd demographics such as the main breeds, flock size, and kidding season, (iii) grazing management including pasture characteristics, grazing and housing periods, supplementary feeding and co-grazing species, and (iv) antiparasitic treatment and control practices, that included the frequency of treatments, dose determination, anthelmintic products used, perceived efficacy of the treatments and the source of information regarding worm control.
2.2 Sampling and laboratory procedure
Owners were instructed to collect faecal samples from 15 randomly selected adult (>18 months of age) and 5 kids (<12 months of age). Individual samples were placed in faecal pots (Sarsted, Germany), marked and placed into zip-seal bags before being posted to UCD PL.
When the samples were received at UCD PL, they were immediately vacuum packed [14] and couriered to CREMOPAR, Italy, where the parasitological analysis was performed [15]. Briefly, equal amounts (approx. 2 g) of faeces from each sample were pooled into four pools of five individuals (where possible). Faecal egg counts were performed on each pooled sample using the FLOTAC dual procedure [16, 17] with a detection limit of six eggs per gram of faeces (EPG). To detect and count gastrointestinal nematodes, cestodes and protozoa, a sodium chloride flotation solution (NaCl, specific gravity = 1.200) was used, while a zinc sulphate-based flotation solution (ZnSO4 specific gravity = 1.350) was used to detect and count trematode eggs and lungworm larvae. All data were recorded in an Excel spreadsheet.
2.3 Data analysis
Data from the questionnaire and faecal analysis were entered on Microsoft Excel (Microsoft 365, Version 2301) and analysed using Epi Info™ (2021 Version 7.2.5.0). Descriptive statistics for each question were analysed. To determine the percentage responses to each question, missing data were excluded from the total number of responses. In the results, the total number of responses (n) are indicated for each percentage.
Parasitological data were summarised as proportion or averages, and 95% confidence intervals (CI) were calculated using Epi Info™ (2021 Version 7.2.5.0). The mean abundance of parasite infections was assessed from the pool faecal samples examined. In this survey, the nomenclature described by Bush et al. [18] was used.
A total of 64 goat owners were invited to participate in the survey and 11 (17%) agreed and returned questionnaires with accompanying samples. The flock size on the majority of farms was less than 100 (n = 5; 50%) with a mean of 23 (range 6–38) adult goats and 20 (range 3–39) kids (Table 1) and included a mixture of different breeds (Table 2, Figures 1–3).
Number of adult goats
Number of farms
Mean number of adult goats (min-max)
Mean number of kids (min-max)
<100
5
22.6 (6–38)
19.8 (3–39)
101–200
3
168.7 (150–200)
200.0 (40–360)
>201
2
245.0 (240–250)
80.0 (40–120)
Table 1.
The number of goats kept on the farms participating in the survey.
Breed
Number of farms
Saanen
5
Toggenburg
3
Old Irish Goat
1
British Alpine/Alpine cross
4
French Alpine cross
1
Anglo-Nubian/Nubian cross
2
Boer Goats Cross
1
Table 2.
Goat breeds kept by farms participating in the survey.
The South African Boer goat – One of the largest goat breeds. Photo by Die_Berlinerin from Pixabay.
Figure 3.
The critically endangered Old Irish goat. Photo by Angela from Pixabay.
Although the main kidding season was spring (n = 10; 91%), kidding also occurred during other seasons (Figure 4). The main farming enterprise was cheese (n = 5; 46%) or milk production (n = 4; 36%) (Figure 5). Five farms (45%) kept animals indoors year-round, while three farms (27%) had year-round grazing, and the remaining farms (n = 3; 27%) turned goats out onto pasture in early spring (March or April) until late autumn (October to November).
Figure 4.
The season when kidding occurred on the participating farms.
Figure 5.
Main enterprise of Irish goat farms.
Of the farms that turn out animals onto pasture, half (n = 3) also co-graze with other ruminants (Figures 6 and 7). On all farms, goats were fed concentrates but also received several other feedstuffs and on four farms (37%) goats did not graze any grass (Figure 6).
Figure 6.
Nutrition of goats on the Irish farms participating in the survey.
Figure 7.
Grazing practices of goat farms in Ireland.
The main parasite control practices and anthelmintic use are summarised in Table 3 and Figure 5. In general, the majority of respondents (n = 9) indicated that they treat all the animals, but only a few (n = 3) treat animals following a set programme (Table 3). More than half of the respondents (n = 5; 55.5%) calculate the dose for treatment based on the heaviest adult animal, generally based on visual inspection (n = 4) (Table 3).
Adults n
Kids n
Dose according to
Treated following a set programme
3
3
Treated at sign of disease
5
4
Time when goats treated
At any other time
2
1
Treated at dry-off
5
—
Treated at housing
4
2
Treated at sign of disease
5
4
Treated at turnout
2
1
Treated at weaning
—
2
Weight basis for dosing
Treat to average weight (visual estimation)
4
4
Treat to heaviest (actual weight)
1
2
Treat to heaviest (visual estimation)
4
2
Anthelmintic dose compared to prescribed sheep dose
Sheep dose
2
1.25x sheep dose
2
1.5x sheep dose
2
2x sheep dose
3
Annual treatment frequency (average)
Nematodes
2.2
2.0
Lungworm
2.2
2.0
Liver fluke
1.8
1.5
Rumen fluke
1.0
1.0
Table 3.
Treatment practices for gastrointestinal parasites.
The choice of anthelmintic drugs varied amongst the farms, but the most common drug class used to treat nematodes was the macrocyclic lactones (ML) 50% (n = 3) followed by benzimidazoles (n = 2; 33%) and levamisole (n = 1; 17%). Only three respondents indicated that they treat the goats for Fasciola hepatica or rumen flukes. Only two respondents indicated the drug used was either benzimidazole (n = 1) or oxyclozanide (n = 1) to treat both these parasites. The actual anthelmintic dose given to goats ranged from the recommended sheep dose (n = 2; 22%) to twice the recommended sheep dose (n = 3; 33%) (Table 3). Past experience and veterinary advice were the most common criteria used when selecting an anthelmintic product (Figure 8).
Figure 8.
Sources used for information on parasite control by Irish goat farmers.
3.2 Gastrointestinal parasite fauna
Pooled samples from 29 adults and 12 goat kids were examined. Different nematode, trematode, cestode and protozoa parasites were detected in the pooled samples from the goats. The mean abundance and prevalence of these parasites in adult goats and kids are summarised in Table 4. Animals on all five farms that reported zero grazing were infected with Eimeria spp. and on the two farms that reported feeding cut grass to the animals were also infected with Strongyloides, strongyle nematodes, Nematodirus and Calicophoron daubneyi and Fasciola hepatica.
Parasite
Mean Abundance
95% CI#
Prevalence %
Mean Abundance
95% CI
Prevalence %
Adults
Kids
Calicophoron daubneyi
13.45
−5.40–32.30
17.2
0
0.0
Eimeria spp.
71.59
37.98–105.20
79.3
862.5
−270.23–1995.23
100.0
Fasciola hepatica
0.21
−0.23–0.63
3.4
0
0.0
“Strongyle” eggs
421.03
80.35–761.73
69.0
223
−23.97–469.97
41.7
Lungworm larvae
6.41
−3.15–15.98
10.3
12.5
−0.14–25.14
33.3
Moniezia spp.*
6.9
16.7
Nematodirus spp.
0.62
−0.09–1.34
10.3
1.5
−0.87–3.87
16.7
Strongyloides spp.
0.21
−0.23–0.63
3.4
1
−1.20 to 3.20
8.3
Trichuris spp.
1.66
0.05–3.26
13.8
8.5
−4.24–21.24
25.0
Table 4.
Mean abundance and prevalence of parasites detected in pooled faecal samples of adult and goat kids in Ireland.
No survey has been done before to investigate parasite and parasite control practices in Irish goat farms. Despite the small number of survey respondents, it still provides some insights into the structure of goat farming in Irland and endoparasite control practices in general. The majority of goat farms were small enterprises with a mean flock size of 23 goats per farm, similar to the situation in the rest of the EU (https://www.europarl.europa.eu/thinktank/en/document/EPRS_BRI(2017)608663). The main breeds kept on farms were Saanen (n = 5; 46%) and British Alpine or Alpine crosses (n = 4; 36%), with diary and cheese production the main enterprises, as is the trend in other European countries [19]. One farm was involved in the conservation of the Old Irish goat breed (Figure 3). This hardy goat breed, capable of thriving in cold, harsh and damp conditions, was introduced to Ireland approximately 5000 years ago. Today it is on the brink of extinction, with only a few hundred individuals left. The Old Irish Goat Society aims to preserve the breed through the help of volunteers (https://backyardgoats.iamcountryside.com/goat-breeds/breed-profile-old-irish-goats/).
As in the rest of Europe, goat farms in this survey were divided into two systems, nearly half of farms (n = 5; 46%) followed a zero-grazing system where animals were permanently confined. Animals were fed on concentrates and other fed forages, provided directly in the feeding trough, which on two farms also included cut grass. Zero grazing allowed farmers to increase milk production [20, 21]. It is, however, important always to be vigilant for possible parasite infections, especially when the grass is fed to animals, as shown in this study where the animals on two farms that reported feeding cut grass were infected with a number of GIH. The majority of farms (n = 6; 55%) in this survey, however, practised pasture-based grazing systems with animals turned onto pasture from early spring to late autumn. Half of these farms (n = 3; 50%) also co-grazed goats with either cattle and/or sheep. This is similar to previous observations that co-grazing is relatively uncommon in goat farming [22, 23]. Mixed species grazing places goats at a higher risk of exposure to infective parasite stages, as goats are susceptible to many of the same parasites as sheep and cattle [2]. Grazing, rather than browsing, also puts goats at greater risk of parasite infection, as several studies have also shown that when goats are forced to graze, they are far more susceptible to infection than sheep [2]. Co-grazing may also play an important role in the transfer of anthelmintic-resistant helminths between host species. This is supported by the detection of early cases of moxidectin and monepantel-resistant nematodes from goats in Australia and New Zealand [24]. Anthelmintic resistance to the three commonly used drug classes BZ, levamisole, and ML is widespread in Irish sheep [11, 13].
Anthelmintics were used on all farms to control gastrointestinal parasites in both adult goats and goat kids, even on the farms that reported following a zero-grazing regiment. The annual treatment frequency for nematodes ranged from 1 to 4 for adults and 1–3 for kids. This is lower than the four or more treatments given to lambs by Irish sheep farmers [25]. It is also lower than the treatment frequencies reported for goats in Europe [23, 26, 27, 28] and New Zealand [29, 30, 31]. The development of anthelmintic resistance has been associated with the frequency of anthelmintic administration, as the selection pressure for resistance increases when the interval between treatments decreases [32, 33]. At least the lower treatment frequency reported in this survey should result in more worms in refugia and therefore put less selection pressure for anthelmintic resistance, but as discussed later, other management factors implement may select for resistance.
ML was the most frequently used anthelmintic class (n = 3; 50%), which was also the trend reported in many other countries [1, 34, 35, 36]. The majority of farmers (8/11; 73%) calculate dose rates based on visual estimation of weight. Similar trends was also reported in other surveys in Europe [23, 27].
With regard to the specific anthelmintic dose rate used in goats, 22% of farms used the prescribed sheep dose, three farms (22%) used twice the recommended sheep dose, and the remaining used between 1.25x and 1.5x the sheep dose. Using an accurate dose to treat helminths is essential to ensure good parasite control efficacy. Reliance on the estimation of weights using visual estimation has been shown to lead to underdosing and increases the selection pressure for resistance in sheep [22, 32, 33, 37]. Accurate dose calculation is especially important in goats since it has been known for more than 30 years that the pharmacokinetics of anthelmintic drugs in goats is different to that of sheep, and goats require higher dose rates to maintain the same plasma concentration and half-life [6, 8, 9, 38, 39]. This is further complicated because, in most countries, including Ireland, anthelmintic drugs are not specifically registered for use in goats. The only exception is eprinomectin which is licenced for use in goats in Ireland in both injectable and pour-on formulations; however, no distinction is made between sheep and goat doses (https://www.hpra.ie/). Only six of the respondents expressed an opinion on whether or not the anthelmintic treatments work as well as it did in previous years, with two-thirds (66%) believed that drugs are less efficacious than before. However, a FECRT was never performed on any of the farms. Goat farmers mostly rely on past experience and veterinary advice when selecting a drug which is similar to sheep farmers in Ireland and the UK [25, 40] and goat farmers in Denmark [23].
The results indicated that 100% of goats were infected with a variety gastrointestinal and lungworm parasite species, also commonly recorded in previous surveys of sheep in Ireland [11, 41]. Eimeria spp. was the most prevalent parasite detected in 100% and 79% of kids and adult goats, respectively. A high prevalence of Eimeria spp. infection has also been reported in Lithuania [42] and Italy [28, 43]. Infection with strongyle nematodes was also highlighted as prevalent in the goats (69% adult goats; 42% goat kids) in this study, similar to what was recorded in Italy [28, 36] and Lithuania [44]. Parasitism with these endoparasites is normally associated with significant production losses, with a decrease in milk yield and quality, reduced growth rate, discarded organs at slaughter and death [36, 45, 46].
This survey also found lungworm infection in 10% of farms in adult goats and 33% of goat kids. A higher lungworm prevalence was found in dairy goats in France (95.5%) [47], northern Italy (44–78%) [36], and Czech Republic (93.1%) [48]; however, lower infection was detected in Germany (35.1%) [49] and Norway (31.2%) [50]. In general, infection with lungworm parasites seldom causes clinical disease in sheep and goats [51, 52].
The prevalence of Strongyloides papillosus (8.3% and 3.4%, goat kids and adults, respectively) was lower than found in other surveys, which generally vary between age groups, management systems, areas and climates (12.8%) [53], Slovakia (14.05%) [54] and Somalia (25.58%) [55]. Strongyloides infections are generally only sporadically associated with clinical signs in herbivores [56]. An experimental infection with S. papillosus in goats showed clinical signs, including transient diarrhoea, elongated faecal pellets terminally, dehydration, anorexia and anaemia [57]. Nematodirus battus prevalence was also generally low (10.3 & 16.7% adult goats and kids, respectively), but still an interesting finding considering that the faecal samples were collected in late summer (August–October). Nematodirosis is generally regarded as a disease of young lambs in the spring and adult animals play a negligible role in its epidemiology [58]. However, recent studies have found that substantial proportions of Nematodirus eggs will hatch without the need for chilling [59, 60]. This is evidenced by recent reports of outbreaks of nematodirosis later in the grazing season (or indeed in the autumn) in older lambs [61, 62]. The prevalence of Trichuris spp. of 10.3% and 25% in adult goats and goat kids, respectively, was higher than reported in other surveys, which range from 0.7–17% [53, 54, 63]. The overall low prevalence reported in the literature with this parasite may be related to difficulty in detecting light infection of this nematode in faecal flotation techniques [64]. This parasite of the cecum is very seldom associated with clinical disease in ruminants [65, 66].
Trematode infection ranged from 3% for F. hepatica and 17% for Calicophoron spp. in adult goats only. The low Fasciola prevalence was surprising as it is quite common in sheep and cattle throughout Ireland [67, 68] and is responsible for considerable production losses across Europe [69, 70]. There are only a limited number of studies that reported rumen fluke infections in European goats [71]; however, Calicophoron spp. is emerging as a prominent parasite in cattle and sheep in many parts of Europe, including Ireland [71, 72, 73, 74, 75, 76]. A decline in Fasciola prevalence has been noted in a previous survey in Ireland [73]. The immature stages of Calicophoron can cause severe disease and have even been associated with outbreaks in cattle and sheep [77, 78, 79], while the adult ruminal phase is usually clinically inapparent.
This is the first survey of goat parasite control and GIH infection goats in Ireland. The present study does have some limitations, (i) only a very small number of farms were surveyed; and (ii) since the farmers volunteered participation, that could potentially have biased the response as the farmers probably had a particular interest in parasites; (iii) only pooled samples were used to estimate prevalence abundance. However, the authors do believe that it still provides some valuable information on general farming practices and helminth prevalence. Overall, the present results highlight several animal husbandry and parasite control practices on goat farms that may favour the spread or development of anthelmintic resistance in nematode populations. There is a clear need to improve the communication of information on new techniques and methods of parasite control to the farmers. The results also point to the need to develop new commercial drugs for the goat industry in order to avoid the excessive dependence on one class of anthelmintics, with probable consequences on long-term efficacy.
Further studies on caprine gastrointestinal parasites using other techniques, for instance, molecular tools, would be indicated. Additionally, future studies should be more longitudinal in nature to study the effect of season, grazing patterns and goats breeds on the prevalence of gastrointestinal parasites.
The authors wish to thank Cian Condon from Teagasc and the farmers for their participation in the study and all the staff at CREMOPAR for their technical support. The research leading to these results has received funding from the European Union Seventh Framework Programme FP7-KBBE-2011-5 under grant agreement no 288975.
1.Baudinette E, O'Handley R, Trengove C. Anthelmintic resistance of gastrointestinal nematodes in goats: A systematic review and meta-analysis. Veterinary Parasitology. 2022;312:109809. DOI: 10.1016/j.vetpar.2022.109809
2.Hoste H, Sotiraki S, Landau SY, Jackson F, Beveridge I. Goat–Nematode interactions: think differently. Trends in Parasitology. 2010;26(8):376-381. DOI: 10.1016/j.pt.2010.04.007
3.Huntley JF, Patterson M, Mackellar A, Jackson F, Stevenson LM, Coop RL. A comparison of the mast cell and eosinophil responses of sheep and goats to gastrointestinal nematode infections. Research in Veterinary Science. 1995;58(1):5-10. DOI: 10.1016/0034-5288(95)90080-2
4.Hoste H, Torres-Acosta JFJ, Aguilar-Caballero AJ. Nutrition–parasite interactions in goats: is immunoregulation involved in the control of gastrointestinal nematodes? Parasite Immunology. 2008;30(2):79-88. DOI: 10.1111/j.1365-3024.2007.00987.x
5.Pérez J, Zafra R, Buffoni L, Hernández S, Cámara S, Martínez-Moreno A. Cellular phenotypes in the abomasal mucosa and abomasal lymph nodes of goats infected with Haemonchus contortus. Journal of Comparative Pathology. 2008;138(2-3):102-107. DOI: 10.1016/j.jcpa.2007.11.007
6.Várady M, Papadopoulos E, Dolinská M, Königová A. Anthelmintic resistance in parasites of small ruminants: Sheep versus goats. Helminthologia. 2011;48(3):137. DOI: 10.2478/s11687-011-0021-7
7.Hoste H, Chartier C. Response to challenge infection with Haemonchus contortus and Trichostrongylus colubriformis in dairy goats. Consequences on milk production. Veterinary Parasitology. 1998;74(1):43-54. DOI: 10.1016/s0304-4017(97)00130-1
8.Gokbulut C, Yalinkilinc HS, Aksit D, Veneziano V. Comparative pharmacokinetics of levamisole-oxyclozanide combination in sheep and goats following per os administration. Canadian Journal of Veterinary Research. 2014;78(4):316-320
9.Singh P, Scott I, Jacob A, Storillo VM, Pomroy WE. Pharmacokinetics of abamectin in sheep, goat and deer. Small Ruminant Research. 2018;165:30-33. DOI: 10.1016/j.smallrumres.2018.06.009
10.Lanusse CE, Alvarez LI, Lifschitz AL. Gaining insights into the pharmacology of anthelmintics using Haemonchus contortus as a model nematode. In: Gasser RB, Von Samson-Himmelstjerna G, editors. Advances in Parasitology. Vol. 93. Amsterdam: Academic Press; 2016. pp. 465-518
11.Good B, Hanrahan JP, de Waal DT, Patten T, Kinsella A, Lynch CO. Anthelmintic-resistant nematodes in Irish commercial sheep flocks- the state of play. Irish Veterinary Journal. 2012;65(1):21. DOI: 10.1186/2046-0481-65-21
12.Keane OM, Keegan JD, Good B, de Waal T, Fanning J, Gottstein M, et al. High level of treatment failure with commonly used anthelmintics on Irish sheep farms. Irish Veterinary Journal. 2014;67(1):1-7. DOI: 10.1186/2046-0481-67-16
13.Keagan JD, Keane OM, Good B, DeWaal T, Denny M, Hanrahan JP, et al. A nationwide survey of anthelmintic treatment failure on sheep farms in Ireland. Irish. Veterinary Journal. 2017;2017:70. DOI: 10.1186/s13620-017-0086-9
14.Rinaldi L, Coles GC, Maurelli MP, Musella V, Cringoli G. Calibration and diagnostic accuracy of simple flotation, McMaster and FLOTAC for parasite egg counts in sheep. Veterinary Parasitology. 2011;177(3-4):345-352. DOI: 10.1016/j.vetpar.2010.12.010
15.Rinaldi L, Catelan D, Musella V, Cecconi L, Hertzberg H, Torgerson PR, et al. Haemonchus contortus: Spatial risk distribution for infection in sheep in Europe. Geospatial Health. 2015;9(2):325-331. DOI: 10.4081/gh.2015.355
16.Rinaldi L, Cringoli G. Parasitological and pathophysiological methods for selective application of anthelmintic treatments in goats. Small Ruminant Research. 2012;103(1):18-22. DOI: 10.1016/j.smallrumres.2011.10.014
17.Cringoli G, Rinaldi L, Maurelli MP, Utzinger J. FLOTAC: New multivalent techniques for qualitative and quantitative copromicroscopic diagnosis of parasites in animals and humans. Nature Protocols. 2010;5(3):503-515. DOI: 10.1038/nprot.2009.235
18.Bush AO, Lafferty KD, Lotz JM, Shostak AW. Parasitology meets ecology on its own terms: Margolis et al. revisited. The Journal of Parasitology. 1997;83(4):575-583. DOI: 10.2307/3284227
19.Ruiz Morales FA, Castel Genís JM, Guerrero YM. Current status, challenges and the way forward for dairy goat production in Europe. Asian-Australas Journal of Animal Science. 2019;32(8):1256-1265. DOI: 10.5713/ajas.19.0327
20.Goetsch AL. Recent advances in the feeding and nutrition of dairy goats. Asian-Australas Journal of Animal Science. 2019;32(8):1296-1305. DOI: 10.5713/ajas.19.0255
21.Granados-Rivera LD, Jáquez JAM, Bautista Y. Effect of the feeding system (grazing vs. zero grazing) on the production, composition, and fatty acid profile in milk of creole goats in northern Mexico. Emirates Journal of Food and Agriculture. 2022;2022:34. DOI: 10.9755/ejfa.2022.v34.i6.2885
22.Coles GC, Roush RT. Slowing the spread of anthelmintic resistant nematodes of sheep and goats in the United Kingdom. The Veterinary Record. 1992;1992:130. DOI: 10.1136/vr.130.23.505
23.Maingi N, Bjørn H, Thamsborg SM, Dangolla A, Kyvsgaard NC. A questionnaire survey of nematode parasite control practices on goat farms in Denmark. Veterinary Parasitology. 1996;66(1-2):25-37. DOI: 10.1016/S0304-4017(96)01004-7
24.Scott I, Pomroy WE, Kenyon PR, Smith G, Adlington B, Moss A. Lack of efficacy of monepantel against Teladorsagia circumcincta and Trichostrongylus colubriformis. Veterinary Parasitology. 2013;198(1-2):166-171. DOI: 10.1016/j.vetpar.2013.07.037
25.Patten T, Good B, Hanrahan JP, Mulcahy G, de Waal T. Gastrointestinal nematode control practices on lowland sheep farms in Ireland with reference to selection for anthelmintic resistance. Irish Veterinary Journal. 2011;64(1):4. DOI: 10.1186/2046-0481-64-4
26.Domke AVM, Chartier C, Gjerde B, Höglund J, Leine N, Vatn S, et al. Prevalence of anthelmintic resistance in gastrointestinal nematodes of sheep and goats in Norway. Parasitology Research. 2012;111(1):185-193. DOI: 10.1007/s00436-012-2817-x
27.Hoste H, Chartier C, Etter E, Goudeau C, Soubirac F, Lefrileux Y. A questionnaire survey on the practices adopted to control gastrointestinal nematode parasitism in dairy goat farms in France. Veterinary Research Communications. 2000;24(7):459-469. DOI: 10.1023/A:1006499522154
28.Maurizio A, Stancampiano L, Tessarin C, Pertile A, Pedrini G, Asti C, et al. Survey on endoparasites of dairy goats in North-Eastern Italy using a farm-tailored monitoring approach. Veterinary Sciences. 2021;8(5):69. DOI: 10.3390/vetsci8050069
29.Kettle PR, Vlassoff A, Reid TC, Horton CT, et al. A survey of nematode control measures used by milking goat farmers and of anthelmintic resistance on their farms. New Zealand Veterinary Journal. 1983;31(8):139-143. DOI: 10.1080/00480169.1983.34999
30.Pearson AB, MacKenzie R. Parasite control in fibre goats — Results of a postal questionnaire. New Zealand Veterinary Journal. 1986;34(11):198-199. DOI: 10.1080/00480169.1986.35348
31.Scherrer AM, Pomroy WE, Charleston WAG. A survey of anthelmintic resistance on ten goat farms in the Manawatu region in 1988. New Zealand Veterinary Journal. 1989;37(4):148-149. DOI: 10.1080/00480169.1989.35590
32.Prichard RK. Anthelmintic resistance in nematodes: Extent, recent understanding and future directions for control and research. International Journal for Parasitology. 1990;20(4):515-523
33.Woodgate RG, Besier RB. Sustainable use of anthelmintics in an integrated parasite management program for sheep nematodes. Animal Production Science. 2010;50(6):440-443. DOI: 10.1071/AN10022
34.Domke AVM, Chartier C, Gjerde B, Leine N, Vatn S, Østerås O, et al. Worm control practice against gastro-intestinal parasites in Norwegian sheep and goat flocks. Acta Veterinaria Scandinavica. 2011;53(1):29. DOI: 10.1186/1751-0147-53-29
35.Lespine A, Chartier C, Hoste H, Alvinerie M. Endectocides in goats: Pharmacology, efficacy and use conditions in the context of anthelmintics resistance. Small Ruminant Research. 2012;103(1):10-17. DOI: 10.1016/j.smallrumres.2011.10.013
36.Lambertz C, Poulopoulou I, Wuthijaree K, Gauly M. Endoparasitic infections and prevention measures in sheep and goats under mountain farming conditions in Northern Italy. Small Ruminant Research. 2018;164:94-101.https://doi.org/10.1016/j.smallrumres.2018.05.007
37.Edwards JR, Wroth R, de Chaneet GC, Besier RB, Karlsson J, Morcombe PW, et al. Survey of anthelmintic resistance in Western Australian sheep flocks. 1. Prevalence. Australian Veterinary Journal. 1986;63(5):135-138
38.Lanusse C, Alvarez L, Lifschitz A. Pharmacological knowledge and sustainable anthelmintic therapy in ruminants. Veterinary Parasitology. 2014;204(1):18-33. DOI: 10.1016/j.vetpar.2013.11.003
39.Hennessy DR, Sangster NC, Steel JW, Collins GH. Comparative pharmacokinetic behaviour of albendazole in sheep and goats. International Journal for Parasitology. 1993;23(3):321-325. DOI: 10.1016/0020-7519(93)90006-k
40.Morgan ER, Hosking BC, Burston S, Carder KM, Hyslop AC, Pritchard LJ, et al. A survey of helminth control practices on sheep farms in Great Britain and Ireland. The Veterinary Journal. 2012;192(3):390-397. DOI: 10.1016/j.tvjl.2011.08.004
41.Lee AM, Wolfe A, Cassidy JP, Messam LL, Moriarty JP, O’Neill R, et al. First confirmation by PCR of Jaagsiekte sheep retrovirus in Ireland and prevalence of ovine pulmonary adenocarcinoma in adult sheep at slaughter. Irish Veterinary Journal. 2017;70(1):33. DOI: 10.1186/s13620-017-0111-z
42.Stadalienė I, Petkevičius S, Šarkūnas M. The impact of grazing management on seasonal activity of gastrointestinal parasites in goats. Helminthologia. 2014;51(2):103-111. DOI: 10.2478/s11687-014-0217-8
43.Gazzonis AL, Zanzani SA, Giuliani G, Alberti E, Villa L, Manfredi MT. Comparison of naturally occurring eimeria infections in Alpine and Nera di Verzasca goat breeds reared in a sub-alpine environment. The Journal of Parasitology. 2021;107(3):463-471. DOI: 10.1645/20-153
44.Stadalienė I, Höglund J, Petkevičius S. Seasonal patterns of gastrointestinal nematode infection in goats on two Lithuanian farms. Acta Veterinaria Scandinavica. 2015;57(1):16. DOI: 10.1186/s13028-015-0105-3
45.Fthenakis GC, Papadopoulos E. Impact of parasitism in goat production. Small Ruminant Research. 2018;163:21-23. DOI: 10.1016/j.smallrumres.2017.04.001
46.Rinaldi L, Veneziano V, Cringoli G. Dairy goat production and the importance of gastrointestinal strongyle parasitism. Transactions of the Royal Society of Tropical Medicine and Hygiene. 2007;101(8):745-746. DOI: 10.1016/j.trstmh.2007.03.010
47.Chartier C, Reche B. Gastrointestinal helminthes and lungworms of French dairy goats: Prevalence and geographical distribution in Poitou-Charentes. Veterinary Research Communication. 1992;16:327-355
48.Kyriánová I, Vadlejch J, Langrova I. Comparison of lungworm infection in a herd of young and dairy goats at an organic farm. Scientia Agriculturae Bohemica. 2019;50:23-28. DOI: 10.2478/sab-2019-0004
49.Voigt K, Sieber PL, Sauter-Louis C, Knubben-Schweizer G, Scheuerle M. Prevalence of pasture-associated metazoal endoparasites in Bavarian dairy goat herds and farmers' approaches to parasite control. Berliner und Münchener Tierärztliche Wochenschrift. 2016;129(7-8):323-332
50.Domke AVM, Chartier C, Gjerde B, Leine N, Vatn S, Stuen S. Prevalence of gastrointestinal helminths, lungworms and liver fluke in sheep and goats in Norway. Veterinary Parasitology. 2013;194(1):40-48. DOI: 10.1016/j.vetpar.2012.12.023
51.Vadlejch J, Makovický P, Čadková Z, Langrová I. Efficacy and persistent activity of moxidectin against natural Muellerius capillaris infection in goats and pathological consequences of muelleriosis. Veterinary Parasitology. 2016;218:98-101. DOI: 10.1016/j.vetpar.2016.01.009
52.Asmare K, Sibhat B, Haile A, Sheferaw D, Aragaw K, Abera M, et al. Lungworm infection in small ruminants in Ethiopia: Systematic review and meta-analysis. Veterinary Parasitology Reg Study Reports. 2018;14:63-70. DOI: 10.1016/j.vprsr.2018.08.003
53.Tumusiime M, Ndayisenga F, Ntampaka P. Prevalence of gastrointestinal nematodes, cestodes, and protozoans of goats in Nyagatare District, Rwanda. Veterinary Medicine (Auckland, NZ). 2022;13:339-349. DOI: 10.2147/VMRR.S389336
54.Babják M, Königová A, Urda-Dolinská M, Várady M. Gastrointestinal helminth infections of dairy goats in Slovakia. Helminthologia. 2017;54(3):211-217. DOI: 10.1515/helm-2017-0027
55.Abdi-Soojeede MI. Common gastro-intestinal parasites of goats (Capra aegagrus hircus) from Mogadishu, Somalia. Open Journal of Veterinary Medicine. 2018;08(12):232-240. DOI: 10.4236/ojvm.2018.812020
56.Thamsborg SM, Ketzis J, Horii Y, Matthews JB. Strongyloides spp. infections of veterinary importance. Parasitology. 2017;144(3):274-284. DOI: 10.1017/s0031182016001116
57.Pienaar JG, Basson PA, du Plessis JL, Collins HM, Naude TW, Boyazoglu PA, et al. Experimental studies with Stronglyloides papillosus in goats. The Onderstepoort Journal of Veterinary Research. 1999;66(3):191-235
58.Sutherland I, Scott I. Gastrointestinal Nematodes of Sheep and Cattle: Biology and Control. Wiley-Blackwell; 2010
59.van Dijk J, Morgan ER. The influence of temperature on the development, hatching and survival of Nematodirus battus larvae. Parasitology. 2008;135(2):269-283. DOI: 10.1017/S0031182007003812
60.van Dijk J, Morgan ER. Variation in the hatching behaviour of Nematodirus battus: Polymorphic bet hedging? International Journal for Parasitology. 2010;40(6):675-681. DOI: 10.1016/j.ijpara.2009.11.002
61.McMahon C, Gordon AW, Edgar HWJ, Hanna REB, Brennan GP, Fairweather I. The effects of climate change on ovine parasitic gastroenteritis determined using veterinary surveillance and meteorological data for Northern Ireland over the period 1999-2009. Veterinary Parasitology. 2012;190(1):167-177. DOI: 10.1016/j.vetpar.2012.06.016
62.Sargison ND, Wilson DJ, Scott PR. Observations on the epidemiology of autumn nematodirosis in weaned lambs in a Scottish sheep flock. The Veterinary Record. 2012;170(15):391. DOI: 10.1136/vr.100145
63.Ruhoollah KW, Al-Jabr OA, Khan T, Khan A, El-Ghareeb WR, et al. Prevalence of gastrointestinal parasite in small ruminants of district Dir upper Khyber Pakhtunkhwa Province of Pakistan. Brazilian Journal of Biology. 2021;83:e248978. DOI: 10.1590/1519-6984.248978
64.Zvinorova PI, Halimani TE, Muchadeyi FC, Matika O, Riggio V, Dzama K. Breeding for resistance to gastrointestinal nematodes – The potential in low-input/output small ruminant production systems. Veterinary Parasitology. 2016;225:19-28. DOI: 10.1016/j.vetpar.2016.05.015
65.Taylor MA, Coop RL, Wall RL. Parasites of sheep and goats. In: Taylor MA, Coop RL, Wall RL, editors. Veterinary Parasitology. Wiley Blackwell; 2015. pp. 436-523
66.Bulbul KH, Akand AH, Hussain J, Parbin S, Hasin D. A brief understanding of Trichuris ovis in ruminants. International Journal of Veterinary Sciences and Animal Husbandry. 2020;5(3):72-74
67.Munita MP, Rea R, Martinez-Ibeas AM, Byrne N, McGrath G, Munita-Corbalan LE, et al. Liver fluke in Irish sheep: Prevalence and associations with management practices and co-infection with rumen fluke. Parasites & Vectors. 2019;12(1):525. DOI: 10.1186/s13071-019-3779-y
68.Selemetas N, de Waal T. Detection of major climatic and environmental predictors of liver fluke exposure risk in Ireland using spatial cluster analysis. Veterinary Parasitology. 2015;209(3-4):242-253. DOI: 10.1016/j.vetpar.2015.02.029
69.Charlier J, Hoglund J, Morgan ER, Geldhof P, Vercruysse J, Claerebout E. Biology and epidemiology of gastrointestinal nematodes in cattle. The Veterinary Clinics of North America. Food Animal Practice. 2020;36(1):1-15. DOI: 10.1016/j.cvfa.2019.11.001
70.Kahl A, von Samson-Himmelstjerna G, Krücken J, Ganter M. Chronic wasting due to liver and rumen flukes in sheep. Animals. 2021;11(2):549. DOI: 10.3390/ani11020549
71.Alstedt U, Voigt K, Jager MC, Knubben-Schweizer G, Zablotski Y, Strube C, et al. Rumen and liver fluke infections in sheep and goats in northern and southern Germany. Animals. 2022;12(7):876. DOI: 10.3390/ani12070876
72.Toolan DP, Mitchell G, Searle K, Sheehan M, Skuce PJ, Zadoks RN. Bovine and ovine rumen fluke in Ireland—Prevalence, risk factors and species identity based on passive veterinary surveillance and abattoir findings. Veterinary Parasitology. 2015;212(3-4):168-174. DOI: 10.1016/j.vetpar.2015.07.040
73.Naranjo-Lucena A, Munita Corbalán MP, Martínez-Ibeas AM, McGrath G, Murray G, Casey M, et al. Spatial patterns of Fasciola hepatica and Calicophoron daubneyi infections in ruminants in Ireland and modelling of C. daubneyi infection. Parasites & Vectors. 2018;11(1):531. DOI: 10.1186/s13071-018-3114-z
74.Jones RA, Williams HW, Dalesman S, Ayodeji S, Thomas RK, Brophy PM. The prevalence and development of digenean parasites within their intermediate snail host, Galba truncatula, in a geographic area where the presence of Calicophoron daubneyi has recently been confirmed. Veterinary Parasitology. 2017;240:68-74. DOI: 10.1016/j.vetpar.2017.03.021
75.Jones RA, Williams HW, Mitchell S, Robertson S, Macrelli M. Exploration of factors associated with spatial−temporal veterinary surveillance diagnoses of rumen fluke (Calicophoron daubneyi) infections in ruminants using zero-inflated mixed modelling. Parasitology. 2022;149(2):253-260. DOI: 10.1017/S0031182021001761
76.Huson KM, Wild C, Fenn C, Robinson MW. Optimized conditions for the in vitro excystment of Calicophoron daubneyi metacercariae. Parasitology. 2017;2017:1-5. DOI: 10.1017/S0031182017002220
77.O'Shaughnessy J, Garcia-Campos A, McAloon CG, Fagan S, de Waal T, McElroy M, et al. Epidemiological investigation of a severe rumen fluke outbreak on an Irish dairy farm. Parasitology. 2018;145(7):948-952. DOI: 10.1017/s0031182017002086
78.Mason C, Stevenson H, Cox A, Dick I. Disease associated with immature paramphistome infection in sheep. The Veterinary Record. 2012;170(13):343-344. DOI: 10.1136/vr.e2368
79.Pavan Kumar C, Syaama Sundar N, Devi PV. Outbreak of immature paramphistomosis in Nellore Jodipi sheep. Journal of Parasitic Diseases: Official Organ of the Indian Society for Parasitology. 2016;40(2):533-535. DOI: 10.1007/s12639-014-0541-4
Written By
Theo de Waal and Laura Rinaldi
Submitted: 14 February 2023Reviewed: 15 February 2023Published: 28 March 2023
Open access peer-reviewed chapter
