Next Article in Journal
Association of Staphylococcal Populations on Teatcups of Milking Parlours with Vaccination against Staphylococcal Mastitis in Sheep and Goat Farms
Next Article in Special Issue
Exploratory Space–Time Analyses of Reported Lyme Borreliosis Cases in France, 2016–2019
Previous Article in Journal
WGS of Commensal Neisseria Reveals Acquisition of a New Ribosomal Protection Protein (MsrD) as a Possible Explanation for High Level Azithromycin Resistance in Belgium
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Pathogens
Volume 10
Issue 4
10.3390/pathogens10040386
pathogens-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

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

Circulation of Babesia Species and Their Exposure to Humans through Ixodes ricinus

by
Tal Azagi
1,*,
Ryanne I. Jaarsma
1,
Arieke Docters van Leeuwen
1,
Manoj Fonville
1,
Miriam Maas
1,
Frits F. J. Franssen
1,
Marja Kik
2,
Jolianne M. Rijks
2,
Margriet G. Montizaan
2,
Margit Groenevelt
3,
Mark Hoyer
4,
Helen J. Esser
5,
Aleksandra I. Krawczyk
1,6,
David Modrý
7,8,9,
Hein Sprong
1,6 and
Samiye Demir
1
1
Centre for Infectious Disease Control, National Institute for Public Health and the Environment, 3720 BA Bilthoven, The Netherlands
2
Dutch Wildlife Health Centre, Utrecht University, 3584 CL Utrecht, The Netherlands
3
Diergeneeskundig Centrum Zuid-Oost Drenthe, 7741 EE Coevorden, The Netherlands
4
Veterinair en Immobilisatie Adviesbureau, 1697 KW Schellinkhout, The Netherlands
5
Wildlife Ecology & Conservation Group, Wageningen University, 6708 PB Wageningen, The Netherlands
6
Laboratory of Entomology, Wageningen University, 6708 PB Wageningen, The Netherlands
7
Institute of Parasitology, Biology Centre CAS, 370 05 Ceske Budejovice, Czech Republic
8
Department of Botany and Zoology, Faculty of Science, Masaryk University, 611 37 Brno, Czech Republic
9
Department of Veterinary Sciences/CINeZ, Faculty of Agrobiology, Food and Natural Resources, Czech University of Life Sciences Prague, 165 00 Prague, Czech Republic
*
Author to whom correspondence should be addressed.
Pathogens 2021, 10(4), 386; https://doi.org/10.3390/pathogens10040386
Submission received: 21 February 2021 / Revised: 15 March 2021 / Accepted: 19 March 2021 / Published: 24 March 2021
(This article belongs to the Special Issue Current Research on Hard Tick-Borne Diseases)
Browse Figures
Review Reports Versions Notes

Abstract

:
Human babesiosis in Europe has been attributed to infection with Babesia divergens and, to a lesser extent, with Babesia venatorum and Babesia microti, which are all transmitted to humans through a bite of Ixodes ricinus. These Babesia species circulate in the Netherlands, but autochthonous human babesiosis cases have not been reported so far. To gain more insight into the natural sources of these Babesia species, their presence in reservoir hosts and in I. ricinus was examined. Moreover, part of the ticks were tested for co-infections with other tick borne pathogens. In a cross-sectional study, qPCR-detection was used to determine the presence of Babesia species in 4611 tissue samples from 27 mammalian species and 13 bird species. Reverse line blotting (RLB) and qPCR detection of Babesia species were used to test 25,849 questing I. ricinus. Fragments of the 18S rDNA and cytochrome c oxidase subunit I (COI) gene from PCR-positive isolates were sequenced for confirmation and species identification and species-specific PCR reactions were performed on samples with suspected mixed infections. Babesia microti was found in two widespread rodent species: Myodes glareolus and Apodemus sylvaticus, whereas B. divergens was detected in the geographically restricted Cervus elaphus and Bison bonasus, and occasionally in free-ranging Ovis aries. B. venatorum was detected in the ubiquitous Capreolus capreolus, and occasionally in free-ranging O. aries. Species-specific PCR revealed co-infections in C. capreolus and C. elaphus, resulting in higher prevalence of B. venatorum and B. divergens than disclosed by qPCR detection, followed by 18S rDNA and COI sequencing. The non-zoonotic Babesia species found were Babesia capreoli, Babesia vulpes, Babesia sp. deer clade, and badger-associated Babesia species. The infection rate of zoonotic Babesia species in questing I. ricinus ticks was higher for Babesia clade I (2.6%) than Babesia clade X (1.9%). Co-infection of B. microti with Borrelia burgdorferi sensu lato and Neoehrlichia mikurensis in questing nymphs occurred more than expected, which reflects their mutual reservoir hosts, and suggests the possibility of co-transmission of these three pathogens to humans during a tick bite. The ubiquitous spread and abundance of B. microti and B. venatorum in their reservoir hosts and questing ticks imply some level of human exposure through tick bites. The restricted distribution of the wild reservoir hosts for B. divergens and its low infection rate in ticks might contribute to the absence of reported autochthonous cases of human babesiosis in the Netherlands.

1. Introduction

Babesiosis in humans and domesticated animals is caused by infection with tick-borne apicomplexan parasites of the genus Babesia [1]. These parasites are also called piroplasms due to their pear-shaped appearance when replicating in erythrocytes. Initially, three lineages of piroplasms were defined, based on character-state definitions, but more recent phylogenic analyses showed that Piroplasmida comprises of approximately six lineages, of which Babesia species represent at least three distinct clades [2,3,4]. The clade terminology throughout this paper follows the most recent study [3]. The B. microti-like lineage, clade I, includes Babesia microti, Babesia vulpes, Babesia felis, and Babesia species recently identified in badgers [5,6]. Babesia species from clade X, originally designated as Babesia sensu stricto, includes Babesia divergens, Babesia venatorum (formerly called EU1–3), Babesia capreoli, Babesia canis, and Babesia sp. deer clade, sometimes referred to as the European variant of Babesia odocoilei [7]. Particularly Babesia species from clade X are of economic importance in the livestock industry [8,9]. In addition, both clades comprise some species that are considered zoonotic and can affect human health [10,11]. However, it should be noted that within the designated Babesia species, genetic variants exist that might confer their zoonotic potential [12,13,14,15]. In the Netherlands, B. microti, B. divergens, B. venatorum, B. capreoli, Babesia motasi, Babesia caballi, and B. canis have been found in wildlife, domestic animals or zoo animals [16,17,18,19,20,21]. Whether zoonotic variants are present, and what other Babesia species circulate in the Netherlands is not known.
The majority of human cases of babesiosis have been caused by B. microti in the United States, with an (under)estimated incidence of 2000 cases per year [22], followed by Canada and China [11]. In contrast, the number of reported cases in Europe has remained low [23]. To date, approximately fifty cases have been reported in Europe [24,25,26,27]. The majority of these cases have been attributed to B. divergens, three to B. venatorum and two cases to B. microti [28,29]. Persistent infections with B. divergens are mostly confined to asplenic patients, and are characterized by sepsis, severe anemia, hemoglobinuria, and jaundice due to hemolysis. Persistent infections with the other two Babesia species appear to be less severe, although the reported cases were also of asplenic or immunocompromised patients [13,30]. Despite the low clinical incidence, significant seroprevalence rates in humans have been recorded in many parts of Europe, indicating that exposure or (endured) infection with Babesia species is not a rare event [31,32,33,34,35,36]. A first possible explanation for these findings is that the European Babesia species do not give rise to persistent infections in immunocompetent humans, although they may cause seroconversion [37]. A second possible explanation may be that human babesiosis is underdiagnosed due to a milder disease course and non-characteristic symptoms in immunocompetent patients, lack of awareness about the disease, and the difficulty in making a diagnosis [27,38]. Although molecular evidence of infection with B. divergens in humans has been found in the Netherlands, no clinical cases of autochthonous babesiosis have been reported thus far [39].
Many hard ticks (Ixodidae) have been identified as vectors of Babesia species in the literature, but most of these are based on a perceived association with the disease rather than on an objective demonstration of transmission [40,41]. In Europe, Ixodes ricinus ticks serve as the principal vectors of various emerging human tick-borne pathogens, and are also considered to transmit B. microti, B. divergens and B. venatorum to humans [24,42]. Ixodes ricinus feed only once per life stage, and Babesia species have developed the ability to persist through successive tick developmental stages, referred to as transstadial transmission. An additional perpetuation strategy of Babesia species from clade X is transovarial transmission, allowing for the spread of the parasite from a single maternal tick to thousands of offspring [43]. Though several studies already investigated the presence and distribution of Babesia species in I. ricinus in the Netherlands, the accuracy of these studies has been limited due to the relatively low number of ticks tested in combination with the low infection rates of Babesia species [16,44]. A third explanation for the lack of reported cases of babesiosis in the Netherlands might therefore be the low infection rate of zoonotic Babesia species in I. ricinus, hence a low human exposure to Babesia species through tick bites.
The infection rate of Babesia species in I. ricinus ticks depends on the abundance and spread of competent vertebrate hosts [45,46,47]. A competent reservoir host is a vertebrate from which I. ricinus can become infected with Babesia species and vice versa [48]. The identification of competent reservoir hosts for zoonotic Babesia species is key to determine the disease hazard [49,50]. Well-documented hosts for zoonotic Babesia species in Europe are cattle (B. divergens), roe deer (B. venatorum), and small mammals (B. microti). Although some deer species have been incriminated by PCR as hosts of B. divergens in several studies, their role in the transmission of zoonotic B. divergens remains poorly understood [12,13]. To further complicate the matter, Babesia species might have partly overlapping competent hosts [51,52,53], and other tick species, such as Ixodes trianguliceps, might be involved in sylvatic cycles as well [24,54,55,56]. Moreover, the abundance of non-competent hosts may reduce the pathogen prevalence in ticks via a dilution effect [57,58]. A fourth explanation for the lack of reported cases of babesiosis in the Netherlands might be a limited distribution of competent reservoir hosts, which would result in a low or only localized exposure to zoonotic Babesia species [59].
In this cross-sectional study, we tested for the presence of different Babesia species in different vertebrates to determine their role as potential reservoir hosts in the Netherlands. Furthermore, we evaluated the relative abundance of Babesia species in questing ticks and analyzed the probability of co-infections with other tick-borne pathogens (TBPs) as a basis to understand the hazard and exposure of each species known to cause disease.

2. Results

2.1. Babesia in Wildlife

2.1.1. Babesia Sensu Stricto (Clade X)

A total of 4611 vertebrate samples were tested by qPCR for the presence of Babesia DNA. Using qPCR for the specific detection of the Babesia sensu stricto-clade (clade X), infection was found in the cloven-hooved mammals Bison bonasus (21%; n = 19), Capreolus capreolus (85%; n = 608), Cervus elaphus (64%; n = 147), Dama dama (9%; n = 100), Ovis aries (2%; n = 634), but not in Bos taurus (n = 116), Capra aegagrus (n = 5) and Sus scrofa (n = 111) (Table 1). Babesia sensu stricto-clade was not detected in any of the birds (n = 99), carnivores (n = 812), lagomorphs (n = 238), rodents (n = 1550), horses (n = 15), hedgehogs (32) or moles (n = 125) tested (Table 1). The qPCR-positive samples were typed to the species level based on a fragment of the 18 S rRNA gene obtained from conventional PCR (Table 1). The 18 S rRNA fragment of B. capreoli (FJ944828) is only two nucleotides different from B. divergens (AY046576), namely at positions 631 and 663 [51]. Typing was successful only for part of the samples: out of the 635 qPCR-positive Babesia sensu stricto samples, 366 DNA sequences were retrieved. Babesia capreoli was identified in 291 C. capreolus and in four D. dama samples, B. venatorum in three C. capreolus and three O. aries, and B. divergens was successfully typed from 26 C. elaphus, two B. bonasus, and two O. aries tissue samples (Table 1).
Remarkably, 65 of the 18 S rRNA sequences from C. capreolus (roe deer) contained double peaks in the sequence trace files, resulting in ambiguous nucleotides (not shown). Fifteen samples were probably mixed infections with B. capreoli and B. venatorum and in eight sequences B. capreoli and B. divergens could not be differentiated because of the presence of one or two ambiguous nucleotides at positions 631 or 663 (Table 1, designated as 23 ambiguous 18 S rRNA sequences). The 518 qPCR-positive C. capreolus samples were therefore also typed to the species level using DNA sequences obtained from PCR on a fragment of the cytochrome c oxidase subunit I (COI) gene. The genetic diversity of the COI fragment is higher than that of the 18 S rRNA fragment, and can therefore unequivocally distinguish between the different Babesia species. However, the success rate of COI typing is lower than that of 18S rRNA. With 18 S rRNA typing, 316 samples yielded a Babesia sequence compared to 158 samples with cytochrome c oxidase subunit I (COI) sequences. From these COI sequences, 134 were designated as B. capreoli, eight as B. venatorum, and 16 sequences could not be assigned because of the large number of ambiguous nucleotides (not shown).
Capreolus capreolus derived sequences that presented double peaks in some of the sequence trace files were suspected of mixed infections. To be able to estimate their occurrence and to identify the Babesia species involved, these samples were subjected to four separate PCRs with species-specific primers. Each PCR was designed for the specific detection of B. venatorum, B. divergens, B. capreoli or Babesia sp. deer clade. A mixed infection in a C. capreolus sample was identified when two or three of the specific PCRs yielded a product of the right size. From all the C. capreolus samples, 290 were randomly selected and tested. Infection with B. capreoli was found in 89% of the samples (n = 258), B. venatorum was detected in 46% of the samples (n = 132) and B. divergens was not detected in any of the tested samples. Infections with both B. capreoli and B. venatorum were found in 44% of the 290 samples (n = 128).
As mixed infections were also suspected in C. elaphus (red deer), 84 randomly selected samples were subjected to the species-specific PCRs. Out of 77 positive samples, B. divergens was detected in 80% of the samples (n = 67), Babesia sp. deer clade was detected in 74% of the samples (n = 62), B. venatorum was not detected in any of the tested samples. Infection with both B. divergens and Babesia sp. deer clade was found in 71% of the samples (n = 60).

2.1.2. Babesia Clade I

Using qPCR detection of DNA from the B. microti-like-clade, infection was found in Meles meles (84%; n = 128), Nyctereutes procyonoides (29%; n = 7), Vulpes vulpes (72%; n = 173), Apodemus sylvaticus (0.8%; n = 634), Microtus arvalis (3%, n = 100) and in Microtus glareolus (6%; n = 405) (Table 1). DNA from the B. microti-like-clade was not detected in any of the ungulates (n = 1740), birds (n = 99), lagomorphs (n = 238), horses (n = 15) hedgehogs (n = 32), or moles (n = 125) tested, neither in several other mustelid or rodent species (Table 1). qPCR-positive samples were typed to the species level using DNA sequences obtained from a conventional PCR fragment of the 18 S rRNA gene (Table 1), a fraction of which were confirmed by sequencing of the COI gene (Supplementary dataset I). Based on the 18 S rRNA sequences, which were successfully retrieved for part of the samples, B. microti was identified in M. glareolus (n = 12), A. sylvaticus (n = 2), and B. vulpes was found in V. vulpes (n = 58) and N. procyonoides (n = 2), whereas three different Babesia badger-type sequences were identified in M. meles (n = 53).

2.2. Geographic Distribution of Wildlife

The reservoir hosts for B. divergens, C. elaphus (140 blocks) and B. bonasus (9 blocks), were restricted to less than 10% of the surface area of the country (Figure 1A,B). Capreolus capreolus (1483 blocks) is the main reservoir host for B. capreoli and B. venatorum, and has been observed in more than 90% of the surface area of the country (Figure 1C). The distribution of D. dama (279 blocks), which carried B. capreoli and Babesia sp. deer clade-EU is also dispersed, but only in ±20% of the country (Figure 1D). The reservoir hosts for B. microti, namely A. sylvaticus (1549 blocks) and M. glareolus (1291 blocks) (Figure 2A,B) are widely spread and their distribution covers almost the whole of the Netherlands. V. vulpes (1509 blocks) and N. procyonoides (145 blocks) (Figure 2C,D), the reservoir hosts for B. vulpes, have been observed in more than 90% of the surface area of the country. Lastly, M. meles (760 blocks) (Figure 2E), which are reservoir hosts for the Babesia badger-types, were observed in ~50% of the country, mostly in the eastern half of the country. It appears to be absent in areas with sea clay soil and dunes (Figure 1). Babesia clade I was also found in the widely spread M. arvalis (Figure 2F); however, these sequences could not be typed to the species level.

2.3. Babesia Species in Ixodes ricinus

Questing ticks from various field studies in the Netherlands and one Belgian study were screened for Babesia clade I and X by reverse line blot hybridization assay (RLB) until 2012, and thenceforth by qPCR (Table 2 and Table 3).
Babesia sensu stricto-clade DNA was detected in 1.9% (n = 25,849) of the questing I. ricinus nymphs collected in the Netherlands (Table 2). All qPCR-positive samples (n = 489) were typed to the species level by using PCR and sequencing a fragment of the 18 S rRNA gene. Based on the retrieved 18 S rRNA sequences (n = 226), B. venatorum was identified in 0.8% (n = 210), B. capreoli in 0.04% (n = 11), B. divergens in 0.01% (n = 4), and the European variant of Babesia sp. deer clade in <0.01% (n = 1) of the ticks (Table 2). The four B. divergens-positive ticks were from areas where either B. bonasus or C. elaphus were present (not shown). Babesia microti-like-clade DNA was detected in 2.6% (n = 18,626) of the questing I. ricinus nymphs collected in the Netherlands (Table 3). Typing by 18 S rRNA sequencing was successful for a fraction of the qPCR-positive samples (n = 45) identifying all of them as B. microti.
In addition to detection of Babesia, 8831 nymphs collected from the Nijverdal and Rodent studies (Table 2 and Table 3) were analyzed for the presence of Borrelia burgdorferi sensu lato, Neoehrlichia mikurensis, Borrelia miyamotoi, and Anaplasma phagocytophilum. Co-infection of Babesia clade X with A. phagocytophilum occurred significantly less than expected (Table 4). Co-infection of Babesia clade I with B. burgdorferi sl and N. mikurensis occurred significantly more than expected randomly (Table 4).

3. Discussion

We have described the presence of different Babesia species in over 4000 samples from various vertebrate hosts in order to determine their role as potential reservoirs for zoonotic Babesia species in the Netherlands. Moreover, we have shown that some ungulate species may host more than one Babesia species concomitantly.
Babesia sensu stricto species B. capreoli, B. venatorum, B. divergens, and Babesia sp. deer clade were detected in cloven-hooved mammals as has been reported in other European countries [64,65,66,67,68,69,70]. Of the 635 Babesia sensu stricto positive samples, 366 were successfully typed to the species level. B. venatorum was found in a fraction of clade X positive C. capreolus samples (5/518) and of O. aries positive samples (3/10). B. venatorum has been detected in European mammals [64,65,66,67], mainly in C. capreolus, with a prevalence ranging from 0.4% in Italy [66] and 1.6% in the Czech Republic [7] to 26.0% in Germany [65]. Babesia divergens was found in C. elaphus (26/94) and B. bonasus (2/4), and to a lesser extent in O. aries (2/10). Other studies have reported a prevalence in C. elaphus ranging from 2.6% in Switzerland [53] to 33% in Poland [71], where the prevalence in B. bonasus has also been found to be above 30% [72]. However, the numbers in the current study are an underestimation, as typing to the species level using Sanger sequencing was only successful for a fraction of qPCR positive samples. This can be corroborated by the results of the species-specific PCR reactions that were performed for a selection of the samples. Co-infections in C. capreolus and C. elaphus samples revealed B. venatorum and B. divergens sequences were masked behind B. capreoli and Babesia sp. deer clade sequences, respectively. Thus, a higher prevalence of the zoonotic Babesia was uncovered (B. venatorum in 46% of C. capreolus samples and B. divergens in 80% of C. elaphus samples). Evidence showing one host can harbor more than one Babesia species, has been reported previously, in Czech, Norwegian, Swiss, and Austrian deer [7,53,73,74]. These results highlight the limitation of relying upon a single qPCR with successive PCR and Sanger sequencing, as this approach cannot detect concomitant microorganisms of the same genus as accurately as species-specific PCR does. Moreover, these results should encourage the re-evaluation of past surveys and the approach of future ones when dealing with TBPs of various zoonotic species.
Despite the high prevalence of B. divergens in C. elaphus in the Netherlands, the prevalence in questing nymphs was low (<0.01%) in accordance with previous studies from the Netherlands and other European countries [16,75,76,77,78,79,80,81]. Although B. divergens has been detected in I. ricinus nymphs and larvae before and has been experimentally acquired and transmitted by them [82,83], other studies have detected B. divergens predominantly in adult ticks [84]. Moreover, in vivo studies showed only adult ticks can successfully acquire the piroplasm from infected cattle [85] and observations suggesting large mammals are mostly parasitized by adult ticks [37] show these may play a greater role in the acquisition of Babesia clade X. As B. divergens is transmitted both transovarially and transstadially [85,86], piroplasm loads may be higher in larvae than nymphs. These variations could be significant and confound molecular detection methods of a certain sensitivity when loads are low, and it has been suggested that testing salivary glands and not whole ticks may lead to more accurate results [83]. Moreover, Babesia reproduction is induced by feeding [37] and most studies screen questing ticks. Lastly, the low prevalence in ticks may reflect that neither acquisition by adults, nor transovarial and transstadial transmission have been found to be 100% efficient [83,85].
Surprisingly, an analysis of co-infections in a subset of the questing ticks revealed a negative correlation between Clade X and A. phagocytophilum, the etiologic agent of human granulocytic anaplasmosis (HGA), which has been found as a co-infection with B. divergens in reservoir mammals [67,73,87]. Anaplasma phagocytophilum may induce immunosuppression in hosts [88], perhaps facilitating co-infections in the animal reservoirs not found in the tick due to a decrease in fitness of the tick vector when infected with both TBPs. Further investigations are necessary to unravel the underlying mechanism of this remarkable finding.
Although clade X Babesia, mainly B. divergens, has been implied as the cause of most European human babesiosis [12,89], no Dutch cases have been reported to date. Based on the findings of this study, this could be partially explained by the limited spread of the reservoir hosts C. elaphus and B. bonasus, meaning human exposure to infected ticks and consequently disease risk are low. Another possible explanation is that B. divergens strains from cattle and deer differ in their zoonotic potential [12]. It is relevant to note that just as tick populations have been expanding in recent decades [90,91], so has the deer population in Europe, with an increase of 54% in C. elaphus population between 1984 and 2000 [92]. Therefore, surveillance of Babesia in ticks and free ranging vertebrates remains useful for monitoring its future distribution.
Babesia clade I species B. microti, B. vulpes and three different Babesia badger-type haplotypes were found in this study. Babesia vulpes was detected in N. procyonoides and in the widely distributed V. vulpes. Although not zoonotic, this Babesia species has been known to cause disease in dogs [9,56,93,94,95] and has not been previously described in the Netherlands [9]. Likewise, Babesia badger-type haplotypes, which were found in this study in M. meles in accordance with previous publications [96,97], have been implied in symptomatic babesiosis in dogs [6].
Based on 18 S rRNA sequences B. microti was detected in bank voles (M. glareolus) (7%) and wood mice (A. sylvaticus) (0.7%), which supports their involvement in the natural life cycle of B. microti [98,99]. Its presence has been reported in A. sylvaticus in England with a prevalence of 8.8% [100]. However, A. sylvaticus was found to be negative for this parasite in most of the other studies from Europe [98,99,101]. The absence or low parasite load of A. sylvaticus may be due to the preference of this species for an open habitat leading to a lower tick load [101]. In contrast B. microti has been repeatedly detected across Europe in M. glareolus with a prevalence ranging from 0.03% to 40% in Germany and England respectively [99,102].
According to data obtained from the NDFF (the Dutch national database of flora and fauna), both bank voles and wood mice are widely distributed throughout the Netherlands. However, in our study the prevalence in a large cohort of questing ticks was relatively low (<3%). The rate of infected ticks in other European countries has a wide range, from 0.1% to 50.8% in Germany and Poland respectively [103,104]. This large variation in prevalence has been attributed to seasonal differences, patchy distribution of Babesia spp., the short lifespan of the hosts, the differences in the sensitivity of the detection methods and the selection of collection technique (from an animal host or the vegetation) [79,105,106].
The seroprevalence of B. microti in humans was reported to be 1% in eastern Croatia [107], 1.5% in Switzerland [32], 9% in Belgium [35], and 13% in Germany [108], inferring some level of human exposure in European countries [98]. Nonetheless, babesiosis cases caused by B. microti are rare in Europe, as opposed to North America where it is the causative agent of most human piroplasmosis cases [106], possibly due to the lower pathogenicity of the various European strains [12,32,101].
Another layer of complexity is added by the shared sylvatic cycle of B. microti with some B. burgdorferi sl genospecies and N. mikurensis [61]. Our study shows these TBPs co-occurred in questing nymphs significantly more than expected by chance, reflecting their ecological overlap. Co-infections between two or more of these TBPs have previously been reported both in questing ticks [109,110,111] and in ticks feeding on humans [39]. Such co-infections have been hypothesized to affect the course of disease [109,112,113]. Moreover, co-infections of other TBPs with B. burgdorferi sl might go undetected under the diagnosis of Lyme disease.
This and many other studies assess the presence of Babesia DNA, not its viability or infectivity. However, the inability of DNA-based detection methods to asseverate infectiousness does not render the approach irrelevant for surveillance of Babesia, especially when implemented in screening I. ricinus ticks, which have been widely implicated as the primary vectors for Babesia in Europe [24,42].
Routinely monitoring the potential wildlife hosts of zoonotic pathogens and their vectors together with insights into the genetic diversity and enzootic cycles of European strains may help effective and timely response to emerging zoonoses [58,114].

4. Materials and Methods

4.1. Collection of Field Samples

The collection of questing I. ricinus relied predominantly on convenience sampling from previous and ongoing studies in the Netherlands [61,62,115,116]. Spleen samples from wildlife were collected from culled or deceased animals and were either sent to the Dutch National Institute for Public Health and the Environment (RIVM) directly or via the Dutch Wildlife Health Centre (DWHC). The collection of spleen samples from mustelids and Sciurus vulgaris were described in previous studies [117,118]. Spleens and livers of hunted red foxes (V. vulpes) were opportunistically collected in a study for detection of fox tape worm [119]. Spleens and livers of hunted and road-killed raccoon dogs (N. procyonoides) were opportunistically collected in a study on zoonotic pathogens of raccoon dogs [120]. Moles (Talpa europaea) were culled by the employees of the Water Board as part of their control task, and were tested at the RIVM. Blood samples of wild boar were collected by hunters in 2014 and serum was sent to the RIVM as part of an ongoing surveillance of animal diseases, which was coordinated by GD Animal Health [121]. Spleens were collected from 99 birds that were found dead or were euthanized and sent to the DWHC for postmortem examination. These birds were identified as Chloris chloris (n = 7), Coccothraustes coccothraustes (n = 2), Coloeus monedula (n = 6), Fringilla coelebs (n = 3), Garrulus glandarius (n = 1), Parus major (n = 5), Phylloscopus trochilus (n = 1), Pica pica (n = 4), Pyrrhula pyrrhula (n = 1), Sturnus vulgaris (n = 7), Turdus iliacus (n = 5), Turdus merula (n = 49), and Turdus philomelos (n = 8). The collection of EDTA-blood from C. capreolus was described in a previous study [122]. EDTA-blood from livestock (B. bonasus, Bos taurus, Equus ferus caballus, and O. aries) grazing in or adjacent to nature areas was collected by qualified veterinarians for medical purposes. Spleens from rodents were collected in different studies on the ecology of tick-borne pathogens in the Netherlands from 13 locations between August–October 2018 and March–June 2019. Rodent trapping, anesthetization, euthanasia, and all other aspects of the animal experiments were approved by the Central Committee Animal Experimentation in the Netherlands (AVD1040020173624), the Animal Welfare Body of Wageningen University (2017.W-0049), and the Netherlands Ministry of Economic Affairs (FF/75A/2015/014).
Tick species and stages were identified morphologically using stereo-microscope and identification keys [123,124]. All samples were kept frozen (−20 °C or −80 °C) until further processing. Most ticks and tissue samples were collected as convenience sampling. This means there is no congruous tempo-spatial context to the data. This allowed the study to comprise a vast amount of data for the evaluation of the research question. Moreover, ticks from Antwerpen collected as part of an urban tick study in Belgium [62] were included and deemed relevant for this study because of their geographic closeness.

4.2. DNA Extraction, qPCR, and RLB Protocols

The sampling approach meant that for questing nymphs two different molecular detection methods were employed: RLB and qPCR. RLB provides higher specificity, and the ability to detect mixed infections [125,126,127], but overtime was substituted by a more efficient and robust qPCR assay. Because of their differences, this study does not compare the infection prevalence between surveys carried out with different molecular techniques (See Table 2 and Table 3). DNA from questing ticks was extracted by alkaline lysis [128]. DNA from engorged ticks, blood and spleen samples was extracted using the DNeasy® Blood & Tissue kit (QIAGEN, Hilden, Germany) as per the manufacturer’s instructions. To detect potential cross-contamination, negative controls were included in each batch of extraction. DNA lysates were screened for the presence of Babesia species from the B. sensu stricto clade with a qPCR targeting a 62-bp portion of the 18 S rRNA gene [129]. Babesia species from the B. microti-like clade were detected using a qPCR, which targets a 104-bp fragment of the internal transcribed spacer (ITS) region using primers 5’-CTCACACAACGATGAAGGACGCA-3’ (Bmicr_ITS_F), 5’-AACAGAGGCAGTGTGTACAATACATTCAGA-3’ (Bmicr_ITS_R), and the probe 5-Atto 520-GCA +GAATTTAG+CAAAT+CAACAGG-BHQ-1-3’ (Bmicr_ITS_px1). These qPCRs were carried out on a LightCycler 480 (Roche Diagnostics Nederland B.V, Almere, the Netherlands) in a final volume of 20 μL with iQ multiplex Powermix, 3 μL of sample and 0.2 μM for all primers and different concentrations for probes [130]. The presence of Babesia spp. in tick lysates from some studies was determined by PCR followed by RLB as described before [16,60]. Plasmid and negative water controls were used on every plate tested. To minimize contamination, and false-positive samples, the DNA extraction, PCR mix preparation, sample addition, and qPCR analyses were performed in separated air locked dedicated labs.

4.3. DNA Sequencing

Samples that were found positive by qPCR were amplified by conventional PCR, targeting a 400 to 440-bp fragment of the 18 S rRNA gene [16] and a conventional PCR targeting a fragment of the COI gene (Table 5). Moreover, 18 S rDNA is the most common marker used for classification at higher taxonomic levels due to its high sensitivity and a wide range of target species [2]; however, it has some limitations for discrimination at species (or lower) level of piroplasmids and for studies on intraspecific variability because of insufficient sequence variation among closely related species and the presence of several different gene copies [131,132].
The fragment of the mitochondrial COI gene shows high interspecific variability and discrimination accuracy, on the other hand it has some drawbacks such as having higher A + T content [131,133] and lower efficiency of amplification. Thus, both approaches were used in order to type the samples as accurately as possible.
Positive controls of DNA samples from B. divergens and B. microti were kindly supplied by Dr. Simone Caccio (Istituto Superiore di Sanità, Rome). The COI fragments of Babesia from the B. microti-clade were amplified as described previously [134], whereas the Babesia sensu stricto clade sequences were obtained by using 5’-ATWGGATTYTAT ATGAGTAT-3’ (Cox1_Bab_For1) and 5’-ATAATCWGGWATYCTCCTTGG-3’ (Cox1_Bab_Rev1) as primers [7]. PCR products were sequenced by dideoxy-dye termination sequencing of both strands, and compared with sequences in GenBank (http://www.ncbi.nlm.nih.gov/ accessed on 3 January 2019), using BLAST [135].

4.4. Species Identification Based on DNA Sequencing

The sequences were stored and analyzed in BioNumerics (Version 7.1, Applied Math, Sint-Martens-Latem, Belgium) after subtraction of the primer sequences for species identification. The DNA sequences that were generated in this study can be found in Supplementary dataset I.
The collected sequences were aligned with those from related organisms in GenBank, and pairwise alignments were generated using unweighted pair group method with arithmetic mean (UPGMA). A species was assigned to a sequence, which was identical or more than 99% similar to the reference sequences from GenBank. Extra attention was given to the 18 S rRNA fragment, where B. capreoli (FJ944828) is only two nucleotides different from B. divergens (AY046576). Sequences with double peaks in some of the sequence trace files (ambiguous nucleotides) at critical positions were not assigned to the species level.

4.5. Species Specific PCR

Infections with more than one Babesia from the Babesia sensu stricto clade were suspected in C. capreolus and C. elaphus samples on the basis of double peaks in some of the sequence trace files. To be able to quantify the occurrence of mixed infections and to identify the Babesia species involved, species-specific primers were designed for B. venatorum, B. divergens, Babesia sp. deer clade, and B. capreoli. These primers (100 nM each) specifically amplified small fragments of the COI gene (Table 5) using the following PCR program: 15 min at 95 °C, 35 cycles each consisting of 30 s at 95 °C, 30 s at 60 °C, and 1 min at 72 °C, and ending with 10 min at 72 °C. The HotStarTaq Polymerase Kit (Qiagen, Venlo, the Netherlands) was used for all PCR experiments. PCR products were detected by electrophoresis in a 1.5% agarose gel stained with SYBR gold (Invitrogen, Leiden, the Netherlands).

4.6. Data Collection on Geographic Spread of Vertebrates in The Netherlands

The geographic distribution of reservoir hosts for the identified Babesia species was retrieved from https://www.verspreidingsatlas.nl on 4 September 2020 [137], setting the time frame between the year 2000 and 2020. The geographic distribution of these vertebrates is based on field observations made by trained volunteers from the Dutch Mammal Society and are registered in 1 km2 blocks, out of 41,543 km² in the Netherlands.

4.7. Statistical Analyses

In order to test the relation between Babesia clade I or Babesia clade X with other TBPs detected in questing ticks, the significance of the observed (O) co-infections versus the randomly expected (E = (O−E)2/E) co-infections was assessed by the Fisher’s exact test and p-values (p < 0.05) were corrected using the Bonferroni procedure for multiple testing. The analyses were conducted in R [138] using the base package, tidyverse and dplyr [139,140].

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/pathogens10040386/s1, Sup-plementary dataset I.

Author Contributions

Conceptualization, T.A., R.I.J., H.S.; methodology, R.I.J., A.D.v.L., M.F., D.M.; validation R.I.J., M.F., A.D.v.L., A.I.K.; formal Analysis, T.A., S.D., H.S.; investigation, M.G., H.J.E., M.M., D.M., M.K., F.F.J.F., J.M.R., A.I.K., M.H., M.G.M.; writing—original draft preparation, T.A., S.D., H.S.; writing—review and editing, M.G., H.J.E., M.M., D.M., M.G.M, M.K., F.F.J.F., J.M.R., M.H., A.I.K.; supervision, H.S., S.D.; project Administration, A.I.K., H.J.E., A.D.v.L., M.F., H.S.; funding Acquisition, H.S. All authors have read and agreed to the published version of the manuscript.

Funding

This project is supported by the Dutch Ministry of Health, Welfare and Sports. T.A., and H.S. were also supported by a grant from ZonMw (project number 522003007, Ticking on Pandora’s box) and a grant from the EU Interreg North Sea Region program, as part of the NorthTick project. The funders had no role in the study design and interpretation, or the decision to submit the work for publication.

Institutional Review Board Statement

Not applicable; approval of the work by an ethics committee is not required for the pathogen surveillance of dead wildlife in the Netherlands. No materials were specifically collected for this study. Studies from which the original materials were collected were carried out according to the national animal welfare regulations.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available within the manuscript and supplementary files.

Acknowledgments

We thank Katsuhisa Takumi (RIVM) for the data extraction and the preparation of the figures.

Conflicts of Interest

The authors declare no conflict of interest.

Abbreviations

18S ribosomal RNA (18S rRNA), Borrelia burgdorferi sensu lato (B. burgdorferi sl), cytochrome c oxidase subunit I (COI), internal transcribed spacer (ITS), polymerase chain reaction (PCR), quantitative polymerase chain reaction (qPCR), reverse line blotting (RLB), tick-borne pathogen (TBP).

References

  1. Homer, M.J.; Aguilar-Delfin, I.; Telford, S.R., 3rd; Krause, P.J.; Persing, D.H. Babesiosis. Clin. Microbiol Rev. 2000, 13, 451–469. [Google Scholar] [CrossRef] [PubMed]
  2. Schnittger, L.; Rodriguez, A.E.; Florin-Christensen, M.; Morrison, D.A. Babesia: A world emerging. Infect. Genet. Evol. 2012, 12, 1788–1809. [Google Scholar] [CrossRef] [PubMed]
  3. Jalovecka, M.; Sojka, D.; Ascencio, M.; Schnittger, L. Babesia Life Cycle—When Phylogeny Meets Biology. Trends Parasitol. 2019, 35, 356–368. [Google Scholar] [CrossRef] [PubMed]
  4. Schreeg, M.E.; Marr, H.S.; Tarigo, J.L.; Cohn, L.A.; Bird, D.M.; Scholl, E.H.; Levy, M.G.; Wiegmann, B.M.; Birkenheuer, A.J. Mitochondrial Genome Sequences and Structures Aid in the Resolution of Piroplasmida phylogeny. PLoS ONE 2016, 11, e0165702. [Google Scholar] [CrossRef]
  5. Baneth, G.; Florin-Christensen, M.; Cardoso, L.; Schnittger, L. Reclassification of Theileria annae as Babesia vulpes sp. nov. Parasit. Vectors 2015, 8, 207. [Google Scholar] [CrossRef] [Green Version]
  6. Hornok, S.; Horvath, G.; Takacs, N.; Kontschan, J.; Szoke, K.; Farkas, R. Molecular identification of badger-associated Babesia sp. DNA in dogs: Updated phylogeny of piroplasms infecting Caniformia. Parasit. Vectors 2018, 11, 235. [Google Scholar] [CrossRef]
  7. Hrazdilova, K.; Rybarova, M.; Siroky, P.; Votypka, J.; Zintl, A.; Burgess, H.; Steinbauer, V.; Zakovcik, V.; Modry, D. Diversity of Babesia spp. in cervid ungulates based on the 18S rDNA and cytochrome c oxidase subunit I phylogenies. Infect. Genet. Evol. 2020, 77, 104060. [Google Scholar] [CrossRef]
  8. Nathaly Wieser, S.; Schnittger, L.; Florin-Christensen, M.; Delbecq, S.; Schetters, T. Vaccination against babesiosis using recombinant GPI-anchored proteins. Int. J. Parasitol. 2019, 49, 175–181. [Google Scholar] [CrossRef]
  9. Solano-Gallego, L.; Sainz, A.; Roura, X.; Estrada-Pena, A.; Miro, G. A review of canine babesiosis: The European perspective. Parasit. Vectors 2016, 9, 336. [Google Scholar] [CrossRef] [Green Version]
  10. Vannier, E.; Krause, P.J. Human babesiosis. N. Engl. J. Med. 2012, 366, 2397–2407. [Google Scholar] [CrossRef] [Green Version]
  11. Yang, Y.; Christie, J.; Koster, L.; Du, A.; Yao, C. Emerging Human Babesiosis with “Ground Zero” in North America. Microorganisms 2021, 9, 440. [Google Scholar] [CrossRef]
  12. Gray, J.S. Identity of the causal agents of human babesiosis in Europe. Int. J. Med. Microbiol 2006, 296 (Suppl. 40), 131–136. [Google Scholar] [CrossRef]
  13. Gray, J.; Zintl, A.; Hildebrandt, A.; Hunfeld, K.P.; Weiss, L. Zoonotic babesiosis: Overview of the disease and novel aspects of pathogen identity. Ticks Tick Borne Dis. 2010, 1, 3–10. [Google Scholar] [CrossRef]
  14. Hamsikova, Z.; Kazimirova, M.; Harustiakova, D.; Mahrikova, L.; Slovak, M.; Berthova, L.; Kocianova, E.; Schnittger, L. Babesia spp. in ticks and wildlife in different habitat types of Slovakia. Parasit. Vectors 2016, 9, 292. [Google Scholar] [CrossRef] [Green Version]
  15. Goethert, H.K.; Telford, S.R., 3rd. What is Babesia microti? Parasitology 2003, 127, 301–309. [Google Scholar] [CrossRef] [Green Version]
  16. Wielinga, P.R.; Fonville, M.; Sprong, H.; Gaasenbeek, C.; Borgsteede, F.; van der Giessen, J.W. Persistent detection of Babesia EU1 and Babesia microti in Ixodes ricinus in the Netherlands during a 5-year surveillance: 2003–2007. Vector Borne Zoonotic Dis. 2009, 9, 119–122. [Google Scholar] [CrossRef]
  17. Bos, J.H.; Klip, F.C.; Sprong, H.; Broens, E.M.; Kik, M.J.L. Clinical outbreak of babesiosis caused by Babesia capreoli in captive reindeer (Rangifer tarandus tarandus) in the Netherlands. Ticks Tick Borne Dis. 2017, 8, 799–801. [Google Scholar] [CrossRef]
  18. Uilenberg, G.; Rombach, M.C.; Perie, N.M.; Zwart, D. Blood parasites of sheep in the Netherlands. II. Babesia motasi (Sporozoa, Babesiidae). Vet. Q 1980, 2, 3–14. [Google Scholar] [CrossRef]
  19. Uilenberg, G.; Top, P.D.; Arends, P.J.; Kool, P.J.; van Dijk, J.E.; van Schieveen, P.B.; Zwart, D. [Autochthonous babesiosis in dogs in the Netherlands?]. Tijdschr Diergeneeskd 1985, 110, 93–98. [Google Scholar]
  20. Matjila, T.P.; Nijhof, A.M.; Taoufik, A.; Houwers, D.; Teske, E.; Penzhorn, B.L.; de Lange, T.; Jongejan, F. Autochthonous canine babesiosis in the Netherlands. Vet. Parasitol. 2005, 131, 23–29. [Google Scholar] [CrossRef]
  21. Jongejan, F.; Ringenier, M.; Putting, M.; Berger, L.; Burgers, S.; Kortekaas, R.; Lenssen, J.; van Roessel, M.; Wijnveld, M.; Madder, M. Novel foci of Dermacentor reticulatus ticks infected with Babesia canis and Babesia caballi in the Netherlands and in Belgium. Parasit. Vectors 2015, 8, 232. [Google Scholar] [CrossRef] [Green Version]
  22. Carpi, G.; Walter, K.S.; Mamoun, C.B.; Krause, P.J.; Kitchen, A.; Lepore, T.J.; Dwivedi, A.; Cornillot, E.; Caccone, A.; Diuk-Wasser, M.A. Babesia microti from humans and ticks hold a genomic signature of strong population structure in the United States. BMC Genom. 2016, 17, 888. [Google Scholar] [CrossRef] [Green Version]
  23. Krause, P.J. Human babesiosis. Int. J. Parasitol. 2019, 49, 165–174. [Google Scholar] [CrossRef]
  24. Hildebrandt, A.; Gray, J.S.; Hunfeld, K.P. Human babesiosis in Europe: What clinicians need to know. Infection 2013, 41, 1057–1072. [Google Scholar] [CrossRef]
  25. Gonzalez, L.M.; Castro, E.; Lobo, C.A.; Richart, A.; Ramiro, R.; Gonzalez-Camacho, F.; Luque, D.; Velasco, A.C.; Montero, E. First report of Babesia divergens infection in an HIV patient. Int. J. Infect. Dis. 2015, 33, 202–204. [Google Scholar] [CrossRef] [Green Version]
  26. Morch, K.; Holmaas, G.; Frolander, P.S.; Kristoffersen, E.K. Severe human Babesia divergens infection in Norway. Int. J. Infect. Dis. 2015, 33, 37–38. [Google Scholar] [CrossRef] [Green Version]
  27. Paleau, A.; Candolfi, E.; Souply, L.; De Briel, D.; Delarbre, J.M.; Lipsker, D.; Jouglin, M.; Malandrin, L.; Hansmann, Y.; Martinot, M. Human babesiosis in Alsace. Med. Mal. Infect. 2020, 50, 486–491. [Google Scholar] [CrossRef]
  28. Hildebrandt, A.; Hunfeld, K.P.; Baier, M.; Krumbholz, A.; Sachse, S.; Lorenzen, T.; Kiehntopf, M.; Fricke, H.J.; Straube, E. First confirmed autochthonous case of human Babesia microti infection in Europe. Eur. J. Clin. Microbiol. Infect. Dis 2007, 26, 595–601. [Google Scholar] [CrossRef]
  29. Arsuaga, M.; Gonzalez, L.M.; Lobo, C.A.; de la Calle, F.; Bautista, J.M.; Azcarate, I.G.; Puente, S.; Montero, E. First Report of Babesia microti-Caused Babesiosis in Spain. Vector Borne Zoonotic Dis. 2016, 16, 677–679. [Google Scholar] [CrossRef] [Green Version]
  30. Haselbarth, K.; Tenter, A.M.; Brade, V.; Krieger, G.; Hunfeld, K.P. First case of human babesiosis in Germany—Clinical presentation and molecular characterisation of the pathogen. Int. J. Med. Microbiol. 2007, 297, 197–204. [Google Scholar] [CrossRef] [PubMed]
  31. Gorenflot, A.; Moubri, K.; Precigout, E.; Carcy, B.; Schetters, T.P. Human babesiosis. Ann. Trop. Med. Parasitol. 1998, 92, 489–501. [Google Scholar] [CrossRef] [PubMed]
  32. Foppa, I.M.; Krause, P.J.; Spielman, A.; Goethert, H.; Gern, L.; Brand, B.; Telford, S.R., 3rd. Entomologic and serologic evidence of zoonotic transmission of Babesia microti, eastern Switzerland. Emerg. Infect. Dis. 2002, 8, 722–726. [Google Scholar] [CrossRef] [PubMed]
  33. Hunfeld, K.P.; Lambert, A.; Kampen, H.; Albert, S.; Epe, C.; Brade, V.; Tenter, A.M. Seroprevalence of Babesia infections in humans exposed to ticks in midwestern Germany. J. Clin. Microbiol. 2002, 40, 2431–2436. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Gabrielli, S.; Calderini, P.; Cassini, R.; Galuppi, R.; Tampieri, M.P.; Pietrobelli, M.; Cancrini, G. Human exposure to piroplasms in Central and Northern Italy. Vet. Ital. 2014, 50, 41–47. [Google Scholar] [CrossRef]
  35. Lempereur, L.; Shiels, B.; Heyman, P.; Moreau, E.; Saegerman, C.; Losson, B.; Malandrin, L. A retrospective serological survey on human babesiosis in Belgium. Clin. Microbiol. Infect. 2015, 21, 96.e1–96.e7. [Google Scholar] [CrossRef] [Green Version]
  36. Zukiewicz-Sobczak, W.; Zwolinski, J.; Chmielewska-Badora, J.; Galinska, E.M.; Cholewa, G.; Krasowska, E.; Zagorski, J.; Wojtyla, A.; Tomasiewicz, K.; Klapec, T. Prevalence of antibodies against selected zoonotic agents in forestry workers from eastern and southern Poland. Ann. Agric. Environ. Med. 2014, 21, 767–770. [Google Scholar] [CrossRef] [Green Version]
  37. Zintl, A.; Mulcahy, G.; Skerrett, H.E.; Taylor, S.M.; Gray, J.S. Babesia divergens, a bovine blood parasite of veterinary and zoonotic importance. Clin. Microbiol. Rev. 2003, 16, 622–636. [Google Scholar] [CrossRef] [Green Version]
  38. Tijsse-Klasen, E.; Koopmans, M.P.; Sprong, H. Tick-borne pathogen—reversed and conventional discovery of disease. Front. Public Health 2014, 2, 73. [Google Scholar] [CrossRef] [Green Version]
  39. Jahfari, S.; Hofhuis, A.; Fonville, M.; van der Giessen, J.; van Pelt, W.; Sprong, H. Molecular Detection of Tick-Borne Pathogens in Humans with Tick Bites and Erythema Migrans, in the Netherlands. PLoS Negl. Trop. Dis. 2016, 10, e0005042. [Google Scholar] [CrossRef] [Green Version]
  40. Gray, J.S.; Estrada-Pena, A.; Zintl, A. Vectors of Babesiosis. Annu. Rev. Entomol. 2019, 64, 149–165. [Google Scholar] [CrossRef]
  41. Friedhoff, K.T. Transmission of Babesia. In Babesiosis of Domestic Animals and Man; CRC Press: Boca Raton, FL, USA, 1988; pp. 23–52. [Google Scholar]
  42. Sprong, H.; Azagi, T.; Hoornstra, D.; Nijhof, A.M.; Knorr, S.; Baarsma, M.E.; Hovius, J.W. Control of Lyme borreliosis and other Ixodes ricinus-borne diseases. Parasit. Vectors 2018, 11, 145. [Google Scholar] [CrossRef] [Green Version]
  43. Mehlhorn, H.; Shein, E. The piroplasms: Life cycle and sexual stages. Adv. Parasitol. 1984, 23, 37–103. [Google Scholar] [CrossRef]
  44. Coipan, E.C.; Jahfari, S.; Fonville, M.; Maassen, C.B.; van der Giessen, J.; Takken, W.; Takumi, K.; Sprong, H. Spatiotemporal dynamics of emerging pathogens in questing Ixodes ricinus. Front. Cell Infect. Microbiol. 2013, 3, 36. [Google Scholar] [CrossRef] [Green Version]
  45. Takumi, K.; Sprong, H.; Hofmeester, T.R. Impact of vertebrate communities on Ixodes ricinus-borne disease risk in forest areas. Parasit. Vectors 2019, 12, 434. [Google Scholar] [CrossRef] [Green Version]
  46. Hofmeester, T.; Coipan, E.; Van Wieren, S.; Prins, H.; Takken, W.; Sprong, H. Few vertebrate species dominate the Borrelia burgdorferi sl life cycle. Environ. Res. Lett. 2016, 11, 043001. [Google Scholar] [CrossRef] [Green Version]
  47. Mihalca, A.D.; Sandor, A.D. The role of rodents in the ecology of Ixodes ricinus and associated pathogens in Central and Eastern Europe. Front. Cell Infect. Microbiol. 2013, 3, 56. [Google Scholar] [CrossRef] [Green Version]
  48. Martin, L.B.; Burgan, S.C.; Adelman, J.S.; Gervasi, S.S. Host Competence: An Organismal Trait to Integrate Immunology and Epidemiology. Integr. Comp. Biol. 2016, 56, 1225–1237. [Google Scholar] [CrossRef] [Green Version]
  49. Keesing, F.; Brunner, J.; Duerr, S.; Killilea, M.; Logiudice, K.; Schmidt, K.; Vuong, H.; Ostfeld, R.S. Hosts as ecological traps for the vector of Lyme disease. Proc. Biol. Sci. 2009, 276, 3911–3919. [Google Scholar] [CrossRef] [Green Version]
  50. LoGiudice, K.; Ostfeld, R.S.; Schmidt, K.A.; Keesing, F. The ecology of infectious disease: Effects of host diversity and community composition on Lyme disease risk. Proc. Natl. Acad. Sci. USA 2003, 100, 567–571. [Google Scholar] [CrossRef] [Green Version]
  51. Malandrin, L.; Jouglin, M.; Sun, Y.; Brisseau, N.; Chauvin, A. Redescription of Babesia capreoli (Enigk and Friedhoff, 1962) from roe deer (Capreolus capreolus): Isolation, cultivation, host specificity, molecular characterisation and differentiation from Babesia divergens. Int. J. Parasitol. 2010, 40, 277–284. [Google Scholar] [CrossRef]
  52. Overzier, E.; Pfister, K.; Herb, I.; Mahling, M.; Bock, G., Jr.; Silaghi, C. Detection of tick-borne pathogens in roe deer (Capreolus capreolus), in questing ticks (Ixodes ricinus), and in ticks infesting roe deer in southern Germany. Ticks Tick Borne Dis. 2013, 4, 320–328. [Google Scholar] [CrossRef] [PubMed]
  53. Michel, A.O.; Mathis, A.; Ryser-Degiorgis, M.P. Babesia spp. in European wild ruminant species: Parasite diversity and risk factors for infection. Vet. Res. 2014, 45, 65. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Bown, K.J.; Lambin, X.; Telford, G.R.; Ogden, N.H.; Telfer, S.; Woldehiwet, Z.; Birtles, R.J. Relative importance of Ixodes ricinus and Ixodes trianguliceps as vectors for Anaplasma phagocytophilum and Babesia microti in field vole (Microtus agrestis) populations. Appl. Environ. Microbiol. 2008, 74, 7118–7125. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Cayol, C.; Jaaskelainen, A.; Koskela, E.; Kyrolainen, S.; Mappes, T.; Siukkola, A.; Kallio, E.R. Sympatric Ixodes-tick species: Pattern of distribution and pathogen transmission within wild rodent populations. Sci. Rep. 2018, 8, 16660. [Google Scholar] [CrossRef]
  56. Najm, N.A.; Meyer-Kayser, E.; Hoffmann, L.; Herb, I.; Fensterer, V.; Pfister, K.; Silaghi, C. A molecular survey of Babesia spp. and Theileria spp. in red foxes (Vulpes vulpes) and their ticks from Thuringia, Germany. Ticks Tick Borne Dis. 2014, 5, 386–391. [Google Scholar] [CrossRef]
  57. Dudek, K. Impact of biodiversity on tick-borne diseases. Przegl Epidemiol. 2014, 68, 681–684. [Google Scholar]
  58. Tomassone, L.; Berriatua, E.; De Sousa, R.; Duscher, G.G.; Mihalca, A.D.; Silaghi, C.; Sprong, H.; Zintl, A. Neglected vector-borne zoonoses in Europe: Into the wild. Vet. Parasitol. 2018, 251, 17–26. [Google Scholar] [CrossRef] [Green Version]
  59. Lambin, E.F.; Tran, A.; Vanwambeke, S.O.; Linard, C.; Soti, V. Pathogenic landscapes: Interactions between land, people, disease vectors, and their animal hosts. Int. J. Health Geogr. 2010, 9, 54. [Google Scholar] [CrossRef] [Green Version]
  60. Tijsse-Klasen, E.; Fonville, M.; Reimerink, J.H.; Spitzen-van der Sluijs, A.; Sprong, H. Role of sand lizards in the ecology of Lyme and other tick-borne diseases in the Netherlands. Parasit. Vectors 2010, 3, 42. [Google Scholar] [CrossRef] [Green Version]
  61. Krawczyk, A.I.; van Duijvendijk, G.L.A.; Swart, A.; Heylen, D.; Jaarsma, R.I.; Jacobs, F.H.H.; Fonville, M.; Sprong, H.; Takken, W. Effect of rodent density on tick and tick-borne pathogen populations: Consequences for infectious disease risk. Parasit. Vectors 2020, 13, 34. [Google Scholar] [CrossRef] [Green Version]
  62. Heylen, D.; Lasters, R.; Adriaensen, F.; Fonville, M.; Sprong, H.; Matthysen, E. Ticks and tick-borne diseases in the city: Role of landscape connectivity and green space characteristics in a metropolitan area. Sci. Total Environ. 2019, 670, 941–949. [Google Scholar] [CrossRef]
  63. Sprong, H.; Moonen, S.; van Wieren, S.E.; Hofmeester, T.R. Effects of cattle grazing on Ixodes ricinus-borne disease risk in forest areas of the Netherlands. Ticks Tick Borne Dis. 2020, 11, 101355. [Google Scholar] [CrossRef]
  64. Bonnet, S.; Jouglin, M.; L’Hostis, M.; Chauvin, A. Babesia sp. EU1 from roe deer and transmission within Ixodes ricinus. Emerg. Infect. Dis. 2007, 13, 1208–1210. [Google Scholar] [CrossRef]
  65. Kauffmann, M.; Rehbein, S.; Hamel, D.; Lutz, W.; Heddergott, M.; Pfister, K.; Silaghi, C. Anaplasma phagocytophilum and Babesia spp. in roe deer (Capreolus capreolus), fallow deer (Dama dama) and mouflon (Ovis musimon) in Germany. Mol. Cell Probes 2017, 31, 46–54. [Google Scholar] [CrossRef]
  66. Zanet, S.; Trisciuoglio, A.; Bottero, E.; de Mera, I.G.; Gortazar, C.; Carpignano, M.G.; Ferroglio, E. Piroplasmosis in wildlife: Babesia and Theileria affecting free-ranging ungulates and carnivores in the Italian Alps. Parasit. Vectors 2014, 7, 70. [Google Scholar] [CrossRef] [Green Version]
  67. Welc-Faleciak, R.; Werszko, J.; Cydzik, K.; Bajer, A.; Michalik, J.; Behnke, J.M. Co-infection and genetic diversity of tick-borne pathogens in roe deer from Poland. Vector Borne Zoonotic Dis. 2013, 13, 277–288. [Google Scholar] [CrossRef] [Green Version]
  68. Duh, D.; Petrovec, M.; Bidovec, A.; Avsic-Zupanc, T. Cervids as Babesiae hosts, Slovenia. Emerg. Infect. Dis. 2005, 11, 1121–1123. [Google Scholar] [CrossRef]
  69. Garcia-Sanmartin, J.; Aurtenetxe, O.; Barral, M.; Marco, I.; Lavin, S.; Garcia-Perez, A.L.; Hurtado, A. Molecular detection and characterization of piroplasms infecting cervids and chamois in Northern Spain. Parasitology 2007, 134, 391–398. [Google Scholar] [CrossRef]
  70. Cezanne, R.; Mrowietz, N.; Eigner, B.; Duscher, G.G.; Glawischnig, W.; Fuehrer, H.P. Molecular analysis of Anaplasma phagocytophilum and Babesia divergens in red deer (Cervus elaphus) in Western Austria. Mol. Cell Probes 2017, 31, 55–58. [Google Scholar] [CrossRef]
  71. Sawczuk, M.; Maciejewska, A.; Adamska, M.; Skotarczak, B. [Roe deer (Capreolus capreolus) and red deer (Cervus elaphus) as a reservoir of protozoans from Babesia and Theileria genus in north-western Poland]. Wiad Parazytol. 2005, 51, 243–247. [Google Scholar]
  72. Karbowiak, G.; Demiaszkiewicz, A.W.; Pyziel, A.M.; Wita, I.; Moskwa, B.; Werszko, J.; Bien, J.; Gozdzik, K.; Lachowicz, J.; Cabaj, W. The parasitic fauna of the European bison (Bison bonasus) (Linnaeus, 1758) and their impact on the conservation. Part 1. The summarising list of parasites noted. Acta Parasitol. 2014, 59, 363–371. [Google Scholar] [CrossRef]
  73. Razanske, I.; Rosef, O.; Radzijevskaja, J.; Bratchikov, M.; Griciuviene, L.; Paulauskas, A. Prevalence and co-infection with tick-borne Anaplasma phagocytophilum and Babesia spp. in red deer (Cervus elaphus) and roe deer (Capreolus capreolus) in Southern Norway. Int. J. Parasitol. Parasites Wildl. 2019, 8, 127–134. [Google Scholar] [CrossRef]
  74. Silaghi, C.; Hamel, D.; Pfister, K.; Rehbein, S. Babesia species and co-infection with Anaplasma phagocytophilum in free-ranging ungulates from Tyrol (Austria). Tierärztliche Mschr. Vet. Med. Austria 2011, 98, 268–274. [Google Scholar]
  75. Michelet, L.; Delannoy, S.; Devillers, E.; Umhang, G.; Aspan, A.; Juremalm, M.; Chirico, J.; van der Wal, F.J.; Sprong, H.; Boye Pihl, T.P.; et al. High-throughput screening of tick-borne pathogens in Europe. Front. Cell Infect. Microbiol. 2014, 4, 103. [Google Scholar] [CrossRef] [PubMed]
  76. Lempereur, L.; Wirtgen, M.; Nahayo, A.; Caron, Y.; Shiels, B.; Saegerman, C.; Losson, B.; Linden, A. Wild cervids are host for tick vectors of babesia species with zoonotic capability in Belgium. Vector Borne Zoonotic Dis. 2012, 12, 275–280. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  77. Schorn, S.; Pfister, K.; Reulen, H.; Mahling, M.; Silaghi, C. Occurrence of Babesia spp., Rickettsia spp. and Bartonella spp. in Ixodes ricinus in Bavarian public parks, Germany. Parasit. Vectors 2011, 4, 135. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  78. Adamska, M.; Skotarczak, B. Molecular detecting of piroplasms in feeding and questing Ixodes ricinus ticks. Ann. Parasitol. 2017, 63, 21–26. [Google Scholar] [CrossRef] [PubMed]
  79. Karlsson, M.E.; Andersson, M.O. Babesia species in questing Ixodes ricinus, Sweden. Ticks Tick Borne Dis. 2016, 7, 10–12. [Google Scholar] [CrossRef]
  80. Schotta, A.M.; Wijnveld, M.; Stockinger, H.; Stanek, G. Approaches for Reverse Line Blot-Based Detection of Microbial Pathogens in Ixodes ricinus Ticks Collected in Austria and Impact of the Chosen Method. Appl. Environ. Microbiol. 2017, 83. [Google Scholar] [CrossRef] [Green Version]
  81. Egyed, L.; Elo, P.; Sreter-Lancz, Z.; Szell, Z.; Balogh, Z.; Sreter, T. Seasonal activity and tick-borne pathogen infection rates of Ixodes ricinus ticks in Hungary. Ticks Tick Borne Dis. 2012, 3, 90–94. [Google Scholar] [CrossRef]
  82. Joyner, L.P.; Davies, S.F.; Kendall, S.B. The experimental transmission of Babesia divergens by Ixodes Ricinus. Exp. Parasitol. 1963, 14, 367–373. [Google Scholar] [CrossRef]
  83. Bonnet, S.; Jouglin, M.; Malandrin, L.; Becker, C.; Agoulon, A.; L’Hostis, M.; Chauvin, A. Transstadial and transovarial persistence of Babesia divergens DNA in Ixodes ricinus ticks fed on infected blood in a new skin-feeding technique. Parasitology 2007, 134, 197–207. [Google Scholar] [CrossRef]
  84. Radzijevskaja, J.; Paulauskas, A.; Rosef, O. Prevalence of Anaplasma phagocytophilum and Babesia divergens in Ixodes ricinus ticks from Lithuania and Norway. Int. J. Med Microbiol. 2008, 298, 218–221. [Google Scholar] [CrossRef] [Green Version]
  85. Donnelly, J.; Peirce, M.A. Experiments on the transmission of Babesia divergens to cattle by the tick Ixodes ricinus. Int. J. Parasitol. 1975, 5, 363–367. [Google Scholar] [CrossRef]
  86. Mackenstedt, U.; Gauer, M.; Mehlhorn, H.; Schein, E.; Hauschild, S. Sexual cycle of Babesia divergens confirmed by DNA measurements. Parasitol. Res. 1990, 76, 199–206. [Google Scholar] [CrossRef]
  87. Andersson, M.O.; Vichova, B.; Tolf, C.; Krzyzanowska, S.; Waldenstrom, J.; Karlsson, M.E. Co-infection with Babesia divergens and Anaplasma phagocytophilum in cattle (Bos taurus), Sweden. Ticks Tick Borne Dis. 2017, 8, 933–935. [Google Scholar] [CrossRef]
  88. Lommano, E.; Bertaiola, L.; Dupasquier, C.; Gern, L. Infections and coinfections of questing Ixodes ricinus ticks by emerging zoonotic pathogens in Western Switzerland. Appl. Environ. Microbiol. 2012, 78, 4606–4612. [Google Scholar] [CrossRef] [Green Version]
  89. Meer-Scherrer, L.; Adelson, M.; Mordechai, E.; Lottaz, B.; Tilton, R. Babesia microti infection in Europe. Curr. Microbiol. 2004, 48, 435–437. [Google Scholar] [CrossRef]
  90. Hvidsten, D.; Frafjord, K.; Gray, J.S.; Henningsson, A.J.; Jenkins, A.; Kristiansen, B.E.; Lager, M.; Rognerud, B.; Slatsve, A.M.; Stordal, F.; et al. The distribution limit of the common tick, Ixodes ricinus, and some associated pathogens in north-western Europe. Ticks Tick Borne Dis. 2020, 11, 101388. [Google Scholar] [CrossRef]
  91. Gray, J.S.; Dautel, H.; Estrada-Pena, A.; Kahl, O.; Lindgren, E. Effects of climate change on ticks and tick-borne diseases in europe. Interdiscip. Perspect. Infect. Dis. 2009, 2009, 593232. [Google Scholar] [CrossRef]
  92. Burbaitė, L.; Csányi, S. Red deer population and harvest changes in Europe. Acta Zool. Litu. 2010, 20, 179–188. [Google Scholar] [CrossRef] [Green Version]
  93. Checa, R.; Lopez-Beceiro, A.M.; Montoya, A.; Barrera, J.P.; Ortega, N.; Galvez, R.; Marino, V.; Gonzalez, J.; Olmeda, A.S.; Fidalgo, L.E.; et al. Babesia microti-like piroplasm (syn. Babesia vulpes) infection in red foxes (Vulpes vulpes) in NW Spain (Galicia) and its relationship with Ixodes hexagonus. Vet. Parasitol. 2018, 252, 22–28. [Google Scholar] [CrossRef]
  94. Falkeno, U.; Tasker, S.; Osterman-Lind, E.; Tvedten, H.W. Theileria annae in a young Swedish dog. Acta Vet. Scand. 2013, 55, 50. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Rene-Martellet, M.; Moro, C.V.; Chene, J.; Bourdoiseau, G.; Chabanne, L.; Mavingui, P. Update on epidemiology of canine babesiosis in Southern France. BMC Vet. Res. 2015, 11, 223. [Google Scholar] [CrossRef] [PubMed]
  96. Bartley, P.M.; Wilson, C.; Innes, E.A.; Katzer, F. Detection of Babesia DNA in blood and spleen samples from Eurasian badgers (Meles meles) in Scotland. Parasitology 2017, 144, 1203–1210. [Google Scholar] [CrossRef] [PubMed]
  97. Hornok, S.; Trauttwein, K.; Takacs, N.; Hodzic, A.; Duscher, G.G.; Kontschan, J. Molecular analysis of Ixodes rugicollis, Candidatus Neoehrlichia sp. (FU98) and a novel Babesia genotype from a European badger (Meles meles). Ticks Tick Borne Dis. 2017, 8, 41–44. [Google Scholar] [CrossRef]
  98. Beck, R.; Vojta, L.; Curkovic, S.; Mrljak, V.; Margaletic, J.; Habrun, B. Molecular survey of Babesia microti in wild rodents in central Croatia. Vector Borne Zoonotic Dis. 2011, 11, 81–83. [Google Scholar] [CrossRef]
  99. Obiegala, A.; Pfeffer, M.; Pfister, K.; Karnath, C.; Silaghi, C. Molecular examinations of Babesia microti in rodents and rodent-attached ticks from urban and sylvatic habitats in Germany. Ticks Tick Borne Dis. 2015, 6, 445–449. [Google Scholar] [CrossRef]
  100. Turner, C.M. Seasonal and age distributions of Babesia, Hepatozoon, Trypanosoma and Grahamella species in Clethrionomys glareolus and Apodemus sylvaticus populations. Parasitology 1986, 93 (Pt. 2), 279–289. [Google Scholar] [CrossRef]
  101. Duh, D.; Petrovec, M.; Trilar, T.; Avsic-Zupanc, T. The molecular evidence of Babesia microti infection in small mammals collected in Slovenia. Parasitology 2003, 126, 113–117. [Google Scholar] [CrossRef]
  102. Kallio, E.R.; Begon, M.; Birtles, R.J.; Bown, K.J.; Koskela, E.; Mappes, T.; Watts, P.C. First report of Anaplasma phagocytophilum and Babesia microti in rodents in Finland. Vector Borne Zoonotic Dis. 2014, 14, 389–393. [Google Scholar] [CrossRef] [Green Version]
  103. Hartelt, K.; Oehme, R.; Frank, H.; Brockmann, S.O.; Hassler, D.; Kimmig, P. Pathogens and symbionts in ticks: Prevalence of Anaplasma phagocytophilum (Ehrlichia sp.), Wolbachia sp., Rickettsia sp., and Babesia sp. in Southern Germany. Int. J. Med. Microbiol. 2004, 293 (Suppl. 37), 86–92. [Google Scholar] [CrossRef]
  104. Asman, M.; Solarz, K.; Cuber, P.; Gasior, T.; Szilman, P.; Szilman, E.; Tondas, E.; Matzullok, A.; Kusion, N.; Florek, K. Detection of protozoans Babesia microti and Toxoplasma gondii and their co-existence in ticks (Acari: Ixodida) collected in Tarnogorski district (Upper Silesia, Poland). Ann. Agric. Environ. Med. 2015, 22, 80–83. [Google Scholar] [CrossRef] [Green Version]
  105. Lempereur, L.; De Cat, A.; Caron, Y.; Madder, M.; Claerebout, E.; Saegerman, C.; Losson, B. First molecular evidence of potentially zoonotic Babesia microti and Babesia sp. EU1 in Ixodes ricinus ticks in Belgium. Vector Borne Zoonotic Dis. 2011, 11, 125–130. [Google Scholar] [CrossRef] [Green Version]
  106. Karbowiak, G.; Biernat, B.; Stanczak, J.; Werszko, J.; Szewczyk, T.; Sytykiewicz, H. The role of particular ticks developmental stages in the circulation of tick-borne pathogens in Central Europe. 6. Babesia. Ann. Parasitol. 2018, 64, 265–284. [Google Scholar] [CrossRef]
  107. Topolovec, J.; Puntaric, D.; Antolovic-Pozgain, A.; Vukovic, D.; Topolovec, Z.; Milas, J.; Drusko-Barisic, V.; Venus, M. Serologically detected "new" tick-borne zoonoses in eastern Croatia. Croat. Med. J. 2003, 44, 626–629. [Google Scholar]
  108. Hunfeld, K.P.; Allwinn, R.; Peters, S.; Kraiczy, P.; Brade, V. Serologic evidence for tick-borne pathogens other than Borrelia burgdorferi (TOBB) in Lyme borreliosis patients from midwestern Germany. Wien. Klin Wochenschr. 1998, 110, 901–908. [Google Scholar]
  109. Belongia, E.A. Epidemiology and impact of coinfections acquired from Ixodes ticks. Vector Borne Zoonotic Dis. 2002, 2, 265–273. [Google Scholar] [CrossRef]
  110. Raileanu, C.; Moutailler, S.; Pavel, I.; Porea, D.; Mihalca, A.D.; Savuta, G.; Vayssier-Taussat, M. Borrelia Diversity and Co-infection with Other Tick Borne Pathogens in Ticks. Front. Cell Infect. Microbiol. 2017, 7, 36. [Google Scholar] [CrossRef] [Green Version]
  111. Moutailler, S.; Valiente Moro, C.; Vaumourin, E.; Michelet, L.; Tran, F.H.; Devillers, E.; Cosson, J.F.; Gasqui, P.; Van, V.T.; Mavingui, P.; et al. Co-infection of Ticks: The Rule Rather Than the Exception. PLoS Negl. Trop. Dis. 2016, 10, e0004539. [Google Scholar] [CrossRef] [Green Version]
  112. Wormser, G.P.; Dattwyler, R.J.; Shapiro, E.D.; Halperin, J.J.; Steere, A.C.; Klempner, M.S.; Krause, P.J.; Bakken, J.S.; Strle, F.; Stanek, G.; et al. The clinical assessment, treatment, and prevention of lyme disease, human granulocytic anaplasmosis, and babesiosis: Clinical practice guidelines by the Infectious Diseases Society of America. Clin. Infect. Dis. 2006, 43, 1089–1134. [Google Scholar] [CrossRef]
  113. Krause, P.J.; Telford, S.R., 3rd; Spielman, A.; Sikand, V.; Ryan, R.; Christianson, D.; Burke, G.; Brassard, P.; Pollack, R.; Peck, J.; et al. Concurrent Lyme disease and babesiosis. Evidence for increased severity and duration of illness. JAMA 1996, 275, 1657–1660. [Google Scholar] [CrossRef]
  114. Morner, T.; Obendorf, D.L.; Artois, M.; Woodford, M.H. Surveillance and monitoring of wildlife diseases. Rev. Sci. Tech. 2002, 21, 67–76. [Google Scholar] [CrossRef]
  115. Coipan, C.E.; van Duijvendijk, G.L.A.; Hofmeester, T.R.; Takumi, K.; Sprong, H. The genetic diversity of Borrelia afzelii is not maintained by the diversity of the rodent hosts. Parasit. Vectors 2018, 11, 454. [Google Scholar] [CrossRef] [Green Version]
  116. Hofmeester, T.R.; Sprong, H.; Jansen, P.A.; Prins, H.H.T.; van Wieren, S.E. Deer presence rather than abundance determines the population density of the sheep tick, Ixodes ricinus, in Dutch forests. Parasit. Vectors 2017, 10, 433. [Google Scholar] [CrossRef]
  117. Hofmeester, T.R.; Krawczyk, A.I.; van Leeuwen, A.D.; Fonville, M.; Montizaan, M.G.E.; van den Berge, K.; Gouwy, J.; Ruyts, S.C.; Verheyen, K.; Sprong, H. Role of mustelids in the life-cycle of ixodid ticks and transmission cycles of four tick-borne pathogens. Parasit. Vectors 2018, 11, 600. [Google Scholar] [CrossRef]
  118. Ruyts, S.C.; Frazer-Mendelewska, E.; Van Den Berge, K.; Verheyen, K.; Sprong, H. Molecular detection of tick-borne pathogens Borrelia afzelii, Borrelia miyamotoi and Anaplasma phagocytophilum in Eurasian red squirrels (Sciurus vulgaris). Eur. J. Wildl. Res. 2017, 63, 43. [Google Scholar] [CrossRef]
  119. Maas, M.; van Roon, A.; Dam-Deisz, W.; van der Giessen, J. Geringe Verspreiding Van Vossenlintworm in Groningen; RIVM Briefrapport: De Bilt, The Netherlands, 2018. [Google Scholar]
  120. Maas, M.; Mulder, J.; Montizaan, M.; Dam-Deisz, W.; Jaarsma, R.; Takumi, K.; van Roon, A.; Franssen, F.; van der Giessen, J. Zoönotische Pathogenen bij de Wasbeerhond en Wasbeer in Nederland; RIVM Briefrapport: De Bilt, The Netherlands, 2018; pp. 1–28. [Google Scholar]
  121. Guldemond, A.; Dijkman, W.; Keuper, D. Wilde Zwijnen op Weg in Nederland; CLM Onderzoek en Advies: Culemborg, The Netherlands, 2015; pp. 1–48. [Google Scholar]
  122. Rijks, J.M.; Montizaan, M.G.E.; Bakker, N.; de Vries, A.; Van Gucht, S.; Swaan, C.; van den Broek, J.; Grone, A.; Sprong, H. Tick-Borne Encephalitis Virus Antibodies in Roe Deer, the Netherlands. Emerg. Infect. Dis. 2019, 25, 342–345. [Google Scholar] [CrossRef]
  123. Estrada-Pena, A.; D’Amico, G.; Palomar, A.M.; Dupraz, M.; Fonville, M.; Heylen, D.; Habela, M.A.; Hornok, S.; Lempereur, L.; Madder, M.; et al. A comparative test of ixodid tick identification by a network of European researchers. Ticks Tick Borne Dis. 2017, 8, 540–546. [Google Scholar] [CrossRef] [Green Version]
  124. Estrada-Peña, A.; Mihalca, A.D.; Petney, T.N. Ticks of Europe and North Africa: A Guide to Species Identification; Springer: Berlin/Heidelberg, Germany, 2018. [Google Scholar]
  125. Nagore, D.; Garcia-Sanmartin, J.; Garcia-Perez, A.L.; Juste, R.A.; Hurtado, A. Detection and identification of equine Theileria and Babesia species by reverse line blotting: Epidemiological survey and phylogenetic analysis. Vet. Parasitol. 2004, 123, 41–54. [Google Scholar] [CrossRef]
  126. Schnittger, L.; Yin, H.; Qi, B.; Gubbels, M.J.; Beyer, D.; Niemann, S.; Jongejan, F.; Ahmed, J.S. Simultaneous detection and differentiation of Theileria and Babesia parasites infecting small ruminants by reverse line blotting. Parasitol. Res. 2004, 92, 189–196. [Google Scholar] [CrossRef] [PubMed]
  127. Altay, K.; Dumanli, N.; Aktas, M. Molecular identification, genetic diversity and distribution of Theileria and Babesia species infecting small ruminants. Vet. Parasitol. 2007, 147, 161–165. [Google Scholar] [CrossRef] [PubMed]
  128. Schouls, L.M.; Van De Pol, I.; Rijpkema, S.G.; Schot, C.S. Detection and identification of Ehrlichia, Borrelia burgdorferi sensu lato, and Bartonella species in Dutch Ixodes ricinus ticks. J. Clin. Microbiol. 1999, 37, 2215–2222. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  129. Oines, O.; Radzijevskaja, J.; Paulauskas, A.; Rosef, O. Prevalence and diversity of Babesia spp. in questing Ixodes ricinus ticks from Norway. Parasit. Vectors 2012, 5, 156. [Google Scholar] [CrossRef] [Green Version]
  130. Kazimirova, M.; Hamsikova, Z.; Spitalska, E.; Minichova, L.; Mahrikova, L.; Caban, R.; Sprong, H.; Fonville, M.; Schnittger, L.; Kocianova, E. Diverse tick-borne microorganisms identified in free-living ungulates in Slovakia. Parasit. Vectors 2018, 11, 495. [Google Scholar] [CrossRef]
  131. Hrazdilova, K.; Mysliwy, I.; Hildebrand, J.; Bunkowska-Gawlik, K.; Janaczyk, B.; Perec-Matysiak, A.; Modry, D. Paralogs vs. genotypes? Variability of Babesia canis assessed by 18S rDNA and two mitochondrial markers. Vet. Parasitol. 2019, 266, 103–110. [Google Scholar] [CrossRef]
  132. Gou, H.; Guan, G.; Liu, A.; Ma, M.; Xu, Z.; Liu, Z.; Ren, Q.; Li, Y.; Yang, J.; Chen, Z.; et al. A DNA barcode for Piroplasmea. Acta Trop. 2012, 124, 92–97. [Google Scholar] [CrossRef]
  133. Pan, W.; Byrne-Steele, M.; Wang, C.; Lu, S.; Clemmons, S.; Zahorchak, R.J.; Han, J. DNA polymerase preference determines PCR priming efficiency. BMC Biotechnol. 2014, 14, 10. [Google Scholar] [CrossRef] [Green Version]
  134. Tuvshintulga, B.; Sivakumar, T.; Battsetseg, B.; Narantsatsaral, S.O.; Enkhtaivan, B.; Battur, B.; Hayashida, K.; Okubo, K.; Ishizaki, T.; Inoue, N.; et al. The PCR detection and phylogenetic characterization of Babesia microti in questing ticks in Mongolia. Parasitol. Int. 2015, 64, 527–532. [Google Scholar] [CrossRef] [Green Version]
  135. Altschul, S.F.; Gish, W.; Miller, W.; Myers, E.W.; Lipman, D.J. Basic local alignment search tool. J. Mol. Biol. 1990, 215, 403–410. [Google Scholar] [CrossRef]
  136. Coimbra-Dores, M.J.; Jaarsma, R.I.; Carmo, A.O.; Maia-Silva, M.; Fonville, M.; da Costa, D.F.F.; Brandao, R.M.L.; Azevedo, F.; Casero, M.; Oliveira, A.C.; et al. Mitochondrial sequences of Rhipicephalus and Coxiella endosymbiont reveal evidence of lineages co-cladogenesis. FEMS Microbiol. Ecol. 2020, 96. [Google Scholar] [CrossRef]
  137. NDFF. NDFF Dissemination Atlas. Available online: http://verspreidingsatlas.nl (accessed on 4 September 2020).
  138. Team, R. Core. R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing; R Foundation for Statistical Computing: Vienna, Austria, 2012; Available online: https://www.R-project.org (accessed on 4 September 2019).
  139. Wickham, H.; Averick, M.; Bryan, J.; Chang, W.; McGowan, L.D.A.; François, R.; Grolemund, G.; Hayes, A.; Henry, L.; Hester, J. Welcome to the Tidyverse. J. Open Source Softw. 2019, 4, 1686. [Google Scholar] [CrossRef]
  140. Wickham, H.; François, R.; Henry, L.; Müller, K. Dplyr: A Grammar of Data Manipulation. R Package Version 1.0.2. 2020. Available online: https://CRAN.R-project.org/package=dplyr (accessed on 4 September 2020).
Pathogens 10 00386 g001 550
Figure 1. Distribution of wild competent wildlife species for Babesia clade X species found in the Netherlands. (A) Cervus elaphus (Red deer), (B) Bison bonasus (European bison), (C) Capreolus capreolus (Roe deer), (D) Dama dama (Fallow deer). Source: https://www.verspreidingsatlas.nl (accessed on 4 September 2020).
Figure 1. Distribution of wild competent wildlife species for Babesia clade X species found in the Netherlands. (A) Cervus elaphus (Red deer), (B) Bison bonasus (European bison), (C) Capreolus capreolus (Roe deer), (D) Dama dama (Fallow deer). Source: https://www.verspreidingsatlas.nl (accessed on 4 September 2020).
Pathogens 10 00386 g001
Pathogens 10 00386 g002 550
Figure 2. Distribution of competent wildlife species for Babesia clade I species found in the Netherlands. (A) Apodemus sylvaticus (Woodmouse), (B) Myodes glareolus (Bank vole), (C) Vulpes vulpes (Red fox), (D) Nyctereutes procyonoides (Raccoon dog), (E) Meles meles (European badger), (F) Microtus arvalis (Common vole). Source: https://www.verspreidingsatlas.nl (accessed on 4 September 2020).
Figure 2. Distribution of competent wildlife species for Babesia clade I species found in the Netherlands. (A) Apodemus sylvaticus (Woodmouse), (B) Myodes glareolus (Bank vole), (C) Vulpes vulpes (Red fox), (D) Nyctereutes procyonoides (Raccoon dog), (E) Meles meles (European badger), (F) Microtus arvalis (Common vole). Source: https://www.verspreidingsatlas.nl (accessed on 4 September 2020).
Pathogens 10 00386 g002
Table
Table 1. Presence of Babesia species in vertebrate tissue samples.
Table 1. Presence of Babesia species in vertebrate tissue samples.
Vertebrate (Order)Vertebrate
(Species)		Tested
(n)		Clade I
Positive (%)		Clade X
Positive (%)		Samples Successfully Typed
(18 S rRNA Typing)		>99% Identity
(GenBank)
ArtiodactylaBos taurus11600-
Bison bonasus1904 (21%)B. divergens (2)AY046576
Capra aegagrus500-
Capreolus capreolus6080518 (85%)B. capreoli (291)
B. venatorum (3)
Ambiguous (23) **		AY726009
FJ215873
Cervus elaphus147094 (64%)B. divergens (26)
Babesia sp. deer clade (32)		AY046576
HQ638138
Dama dama10009 (9%)B. capreoli (4)
Babesia sp. deer clade-EU (1)		AY726009
HQ638138
Ovis aries634010 (2%)B. divergens (2)
B. venatorum (3)		AY046576
FJ215873
Sus scrofa11100-
Aves13 species9900 -
CarnivoraMartes foina13400-
Martes martes12800-
Meles meles128108 (84%)0B. badger-type A (48)
B. badger-type B (4)
B. badger-type C (1)		KT223484
KT223485
MG799847
Mustela putorius24200-
Nyctereutes procyonoides72 (29%)0B. vulpes (2)AF188001
Vulpes vulpes173124 (72%)0B. vulpes (58)AF188001
EulipotyphlaTalpa europaea12500-
ErinaceidaeErinaceus europaeus3200-
LagomorphaLepus europaeus15000-
Oryctolagus cuniculus8800-
PerissodactylaEquus ferus caballus150 0-
RodentiaApodemus flavicollis2900-
Apodemus sylvaticus6345 (0.8%)0B. microti (2)KX161765
Castor fiber800-
Microtus arvalis1003 (3%)0-
Myodes glareolus40525 (6%)0B. microti (12)KX161765
Ondatra zibethicus21000-
Rattus rattus4900-
Sciurus vulgaris11500-
DNA extracts from spleen or EDTA-blood of 4611 wildlife and free ranging domesticated animals were tested by two qPCRs (Clade I and Clade X). Typing was performed on qPCR-positive samples, and was based on sequencing a fragment of the 18 S rRNA (see methods). Typing of qPCR-positive samples by 18 S rRNA PCR and sequencing was not always successful. ** not typeable due to double peaks at discriminatory nucleotide positions in trace files.
Table
Table 2. Presence of Babesia Clade X in Questing I. Ricinus.
Table 2. Presence of Babesia Clade X in Questing I. Ricinus.
Study AcronymI. ricinusMethodTestedPositiveSamples Successfully Typed/SequencedPeriodProvince
(Reference)stage nn%B. capreoliB. divergensBabesia sp. deer cladeB. venatorumyearssites (n)
National Survey [44]N + ARLB857130.015 132000–2010Gelderland (7)
N + ARLB23270.03 72006–2010Limburg (1)
N + ARLB1995110.006 1 102000–2010Noord-Holland (4)
N + ARLB24210.004 12006–2010Brabant (1)
N + ARLB39340.01 42006–2010Drenthe (2)
N + ARLB23200 2006–2010Overijssel (2)
N + ARLB16220.012 22006–2010Zuid-Holland (1)
NRLB7930.038 32007–2010Friesland (1)
NRLB600 2007Groningen (1)
NRLB4020.05 22007–2010Utrecht (1)
Duin-Kruidberg [16]L + N + ARLB1.488140.009 1 132003–2007Noord-Holland (1)
Lizard study [60]N + ARLB49120.004 22007–2009Gelderland (8)
Drenthe *N + ARLB172730.002 32010–2012Drenthe (32)
Rodent study [61] *NqPCR7637730.01521552012–2014Gelderland (2)
Urban ticks [62]NqPCR1780190.0112 162014–2016Antwerpen, Belgium (13)
AqPCR26830.011 32014–2016Antwerpen, Belgium (11)
Cattle study [63]NqPCR11200 2015Drenthe (1)
NqPCR18120.011 12015Gelderland (1)
NqPCR19110.005 12015Noord-Brabant (1)
NqPCR73510.001 2015Noord-Holland (4)
NqPCR16520.012 12015Overijssel (1)
NqPCR783120.015 82015Utrecht (4)
NqPCR18510.005 12015Zuid-Holland (1)
De Groote Peel * NqPCR7940.051 32017Brabant (1)
Sheep study *N + AqPCR1555240.0151 162017Drenthe (3)
Microbiome *NqPCR560130.023 112018Gelderland (1)
NqPCR52020.004 12018Noord-Holland (1)
NqPCR1960470.0243 332018Utrecht (1)
Nijverdal * NqPCR11942230.187 2019Overijssel (1)
Total 258494891.9%11412102000–2019
Ticks from eleven studies were tested for the presence of Babesia clade I with either RLB or qPCR. Typing was done by RLB directly (n = 62) or conventional PCR and sequencing on a fragment of the 18 S rRNA gene on part of the qPCR-positive samples (n = 164). Stage (N/A) refers to the tick life stage, i.e., adult (A) or nymph (N). * Molecular detection of Babesia clade X was performed for this study. (n) Data of field sites from one study were combined to one data point per province.
Table
Table 3. Presence of Babesia clade I in questing I. ricinus.
Table 3. Presence of Babesia clade I in questing I. ricinus.
Study AcronymI. ricinusMethodTestedPositiveTypingPeriodProvince
(Reference)stage nn%Sequencingyearssites (n)
National Survey [44] N + ARLB85760.7-2000–2010Gelderland (7)
N + ARLB199520.1-2000–2010Noord-Holland (4)
N + ARLB24200-2006–2010Brabant (1)
N + ARLB39300-2006–2010Drenthe (2)
N + ARLB23200-2006–2010Limburg (1)
N + ARLB23283.4-2006–2010Overijssel (2)
N + ARLB16200-2006–2010Zuid-Holland (1)
NRLB7911.3-2007–2010Friesland (1)
NRLB600-2007Groningen (1)
NRLB4000-2007–2010Utrecht (1)
Duin-Kruidberg [16]N + ARLB90820.20B. microti (2)2003–2007Noord-Holland (1)
Lizard study [60]N + ARLB49161.20-2007–2009Gelderland (8)
Drenthe (this study)N + ARLB172700-2010–2012Drenthe (32)
Rodent study [61]NqPCR76373935.1B. microti (39)2012–2014Gelderland (2)
Cattle study [63] *NqPCR11200-2015Drenthe (1)
NqPCR181147.7B. microti (4)2015Gelderland (1)
NqPCR19100-2015Noord-Brabant (1)
NqPCR73500-2015Noord-Holland (4)
NqPCR16500-2015Overijssel (1)
NqPCR78300-2015Utrecht (4)
NqPCR18500-2015Zuid-Holland (1)
De Groote Peel *NqPCR7900-2017Brabant (1)
Nijverdal * NqPCR1194504.20-2019Overijssel (1)
Total 186264822.6B. microti (45)2000–2019
Ticks from eight studies were tested for the presence of Babesia clade I with either RLB or qPCR. Typing was done by conventional PCR and sequencing on a fragment of the 18 S rRNA gene on part of the positive samples (n = 60) and 45 isolates yielded a DNA sequence identified as B. microti. Stage (N/A) refers to the tick life stage, i.e., adult (A) or nymph (N). * Molecular detection of Babesia clade I was performed for this study. (n)Data of field sites from one study were combined to one data point per province.
Table
Table 4. Coinfection of tick-borne pathogens with Babesia species.
Table 4. Coinfection of tick-borne pathogens with Babesia species.
Babesia CladePathogenPositive ObservedExpectedP-ValueOdds Ratio
Babesia Clade IB. burgdorferi sl 1292273651.10 × 10−1511.72
B. miyamotoi261141311.08
N. mikurensis939129471.10 × 10−153.86
A. phagocytophilum64221321.80 × 10−10.62
Babesia Clade X1237611.15
Babesia Clade XB. burgdorferi sl 1292171810.93
B. miyamotoi261640.901.69
N. mikurensis9396130.180.42
A. phagocytophilum642290.040.2
Babesia Clade I4427611.15
A total of 8831 I. ricinus nymphs from Nijverdal and the rodent study (Table 3 and Table 4) were tested for other tick-borne pathogens. A Fisher’s exact test was used in order to evaluate the significance of the observed number of co-infections between either Babesia clade I or Babesia clade X and each tick-borne pathogen, and what would be randomly expected. This was assessed by calculating the Odds Ratio and their 95% confidence intervals (not shown). p-values were corrected using the Bonferroni test.
Table
Table 5. Nucleotide sequences of primers and probes used in this study.
Table 5. Nucleotide sequences of primers and probes used in this study.
BabesiaTargetPrimer (Name)Primer (Sequence)PurposeSize (bp)Reference
Clade IITSBmicr_ITS_F5’-CTCACACAACGATGAAGGACGCA-3’qPCR103 bp[136]
Bmicr_ITS_R5’-AACAGAGGCAGTGTGTACAATACATTCAGA-3’
Bmicr_ITS_Px15′- Atto520-GCA+GAATTTAG+CAAAT+CAACAGG- BHQ1-3′
Clade X18SrRNABab_18SrRNA-F5’-CAGCTTGACGGTAGGGTATTGG-3’qPCR62 bp[129]
Bab_18SrRNA-R5’-TCGAACCCTAATTCCCCGTTA-3’
Bab_18SrRNA-P5’-Atto647N-CGAGGCAGCAACGG-MGB-BHQ2-3’
Babesia spp.18SrRNABath-Fn5’-TAAGAATTTCACCTCTGACAGTTA-3’PCR/SEQ±420 bp[16]
Bath-Rn5’-ACACAGGGAGGTAGTGACAAG-3’
Clade ICOICox1F133GGAGAGCTAGGTAGTAGTGGAGATAGGPCR/SEQ1023 bp[134]
Cox1R1130GTGGAAGTGAGCTACCACATACGCTG
Clade XCOICox1_Bab_For15′-ATWGGATTYTATATGAGTAT-3′PCR/SEQ±1250 bp[7]
Cox1_Bab_Rev15′-ATAATCWGGWATYCTCCTTGG-3′
B. venatorumCOIBven-F1595’-ATTGGAAGTGGTACTGGTTGGACTT-3’PCR538 bpThis study
Bven-R6965’-GACATCATTACGATTCCTATGC-3’
B. divergensCOIBdiv-F1655’-AGTGGAACTGGGTGGACATTGTAC-3’PCR234 bpThis study
Bdiv-R3985’-TACCGGCAATGACAAAAGTAG-3’
B. capreoliCOIBcap-F1655’-AGTGGAACAGGATGGACGCTATAT-3’PCR443 bpThis study
Bcap-R6075’-GTCTGATTACCGAACACTTCC-3’
Babesia sp. deer cladeCOIBodo-F3605’-CTTTGACTGCTTTCTTGTTG-3’PCR434 bpThis study
Bodo-R7935’-ATCATAACAATTCCTATGCTC-3’
The two qPCRs were used for the screening of wildlife and tick samples for the presence of Babesia spp. qPCR-positive samples were further analyzed by conventional PCRs and sequencing for species identification (PCR/SEQ). For the detection of multiple Babesia sensu stricto species (Clade X) in one sample (mixed infections), four conventional PCRs, each specifically targeting one Babesia species, were performed and analyzed by TBE-agarose gel electrophoresis. Abbreviations: Atto520 fluorescent dye; BHQ1 (blackholequencher) quenches the fluorescence of Atto520; “+” stands for Locked Nucleic Acid (LNA), this is used to raise the annealing-temperature of the probe; MGB stands for Minor groove binder, which is also used for raising the annealing-temperature.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Azagi, T.; Jaarsma, R.I.; Docters van Leeuwen, A.; Fonville, M.; Maas, M.; Franssen, F.F.J.; Kik, M.; Rijks, J.M.; Montizaan, M.G.; Groenevelt, M.; et al. Circulation of Babesia Species and Their Exposure to Humans through Ixodes ricinus. Pathogens 2021, 10, 386. https://doi.org/10.3390/pathogens10040386

AMA Style

Azagi T, Jaarsma RI, Docters van Leeuwen A, Fonville M, Maas M, Franssen FFJ, Kik M, Rijks JM, Montizaan MG, Groenevelt M, et al. Circulation of Babesia Species and Their Exposure to Humans through Ixodes ricinus. Pathogens. 2021; 10(4):386. https://doi.org/10.3390/pathogens10040386

Chicago/Turabian Style

Azagi, Tal, Ryanne I. Jaarsma, Arieke Docters van Leeuwen, Manoj Fonville, Miriam Maas, Frits F. J. Franssen, Marja Kik, Jolianne M. Rijks, Margriet G. Montizaan, Margit Groenevelt, and et al. 2021. "Circulation of Babesia Species and Their Exposure to Humans through Ixodes ricinus" Pathogens 10, no. 4: 386. https://doi.org/10.3390/pathogens10040386

APA Style

Azagi, T., Jaarsma, R. I., Docters van Leeuwen, A., Fonville, M., Maas, M., Franssen, F. F. J., Kik, M., Rijks, J. M., Montizaan, M. G., Groenevelt, M., Hoyer, M., Esser, H. J., Krawczyk, A. I., Modrý, D., Sprong, H., & Demir, S. (2021). Circulation of Babesia Species and Their Exposure to Humans through Ixodes ricinus. Pathogens, 10(4), 386. https://doi.org/10.3390/pathogens10040386

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

Article Metrics

Back to TopTop