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Review

Experimental Infection of Ticks: An Essential Tool for the Analysis of Babesia Species Biology and Transmission

by
Sarah I. Bonnet
1,2,* and
Clémence Nadal
3,4
1
Animal Health Department, INRAE, 37380 Nouzilly, France
2
Functional Genetics of Infectious Diseases Unit, Institut Pasteur, CNRS UMR 2000, Université de Paris, 75015 Paris, France
3
Epidemiology Unit, Laboratory for Animal Health, University Paris Est, 94700 Maisons-Alfort, France
4
Anses, INRAE, Ecole Nationale Vétérinaire d’Alfort, UMR BIPAR, Laboratoire de Santé Animale, 94700 Maisons-Alfort, France
*
Author to whom correspondence should be addressed.
Pathogens 2021, 10(11), 1403; https://doi.org/10.3390/pathogens10111403
Submission received: 9 August 2021 / Revised: 22 October 2021 / Accepted: 26 October 2021 / Published: 29 October 2021
(This article belongs to the Special Issue Babesia and Human Babesiosis)
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Abstract

:
Babesiosis is one of the most important tick-borne diseases in veterinary health, impacting mainly cattle, equidae, and canidae, and limiting the development of livestock industries worldwide. In humans, babesiosis is considered to be an emerging disease mostly due to Babesia divergens in Europe and Babesia microti in America. Despite this importance, our knowledge of Babesia sp. transmission by ticks is incomplete. The complexity of vectorial systems involving the vector, vertebrate host, and pathogen, as well as the complex feeding biology of ticks, may be part of the reason for the existing gaps in our knowledge. Indeed, this complexity renders the implementation of experimental systems that are as close as possible to natural conditions and allowing the study of tick-host-parasite interactions, quite difficult. However, it is unlikely that the development of more effective and sustainable control measures against babesiosis will emerge unless significant progress can be made in understanding this tripartite relationship. The various methods used to date to achieve tick transmission of Babesia spp. of medical and veterinary importance under experimental conditions are reviewed and discussed here.

1. Introduction

Babesiosis remains prevalent worldwide and represents an important threat for both humans and animals [1,2]. The disease, impacting mainly cattle, sheep, goat, equidae, canidaecanidae, and accidentally humans, is caused by apicomplexan parasites belonging to the Babesia genus that exclusively infect erythrocytes of their vertebrate hosts [3]. To date, more than 100 Babesia species have been identified [1]. Babesia spp. are transmitted by hard ticks—occasionally by blood transfusion—and require both a competent vertebrate and invertebrate host to maintain the transmission cycle [3].
Human babesiosis is caused by Babesia microti, a Babesia crassa-like pathogen, Babesia divergens, Babesia duncani, and Babesia venatorum, as well as other parasites closely genetically related to these pathogens, such as B. divergens-like, B. duncani-like, and B. microti-like [4]. Infections in otherwise healthy individuals is usually mild to moderate and most cases of severe disease occur in immunocompromised individuals. B. microti is endemic in the northeastern and upper midwestern regions of the United States, while B. duncani is present on the west coast of the country [4]. In Europe, most of the human cases are due to B. divergens, whereas in Asia they are due to B. venatorum, B. crassa-like, and B. microti [4]. In cattle, Babesia spp. have a significant worldwide economic, social, and epidemiological impact and include, among the most important species, B. bovis, B. bigemina, Babesia major, and B. divergens [5]. B. bovis and B. bigemina are present in many countries in Africa, Asia, Australia, Central and South America, and Southern Europe between 40° N and 32° S. B. major is present in Europe, Northwest Africa and Asia, and B. divergens is present in northern Europe [5]. Ovine babesiosis due to Babesia ovis and Babesia motasi is considered as the most critical blood-borne parasitic disease of small ruminants in tropical and non-tropical regions (occurring in South-eastern Europe, North Africa, and Asia) [6]. In equids, Babesia caballi is (with Theileria equi and Theileria haneyi) the agent of equine piroplasmosis known to be endemic in several countries of Africa, Asia, the Americas, and mainly in the Mediterranean basin for Europe [7,8,9]. The disease represents a significant animal health issue and causes notable economic losses for the equine industry. Finally, babesiosis is one of the most important globally extended and quickly spreading tick-borne diseases in dogs worldwide. Babesia canis is the main cause of canine babesiosis in Europe and is only sporadically found around the world, whereas Babesia gibsoni, the most prevalent species, and Babesia vogeli have a global distribution. Babesia rossi, one of the most pathogenic species, is endemic in southern Africa [10].
The current approaches available for babesiosis control have many important limitations, including increased resistance to acaricides by ticks, as well as the numerous drawbacks of these acaricides and of the current vaccines and babesicidal drugs (e.g., efficacy, toxicity, environmental effects) [11]. The development of improved control measures against babesiosis is limited by the numerous and significant gaps in our understanding of the biology of Babesia spp., especially regarding molecular interaction between parasites, vectors, and vertebrate hosts, as well as the factors that may influence both the development and the transmission of the parasite [12]. To fill these gaps, it is essential to be able to reproduce the life cycle of Babesia species in controlled experimental conditions, including transmission by ticks [13,14]. In addition, the validation of new methods of interruption of the cycle requires that it can be first carried out entirely under such conditions. Finally, it is important to mention that despite the promise of in vitro culture systems [15], maintenance by in vitro culture [16] or needle-passage in the vertebrate host [17] in the absence of tick passage may generate significant changes in the parasite population, potentially creating a bias in research results.
Several laboratory studies that aimed to understand babesiosis pathogenesis have focused their interest on infecting laboratory animals through artificial parasite injection [18,19]. Regarding tick-parasite interaction, the great majority of studies carried out concern mainly epidemiological studies focusing on the detection of parasites in ticks collected into the field [20]. Some in vitro studies have also been performed in order to understand the interactions between parasites and the cells of their vertebrate [21,22,23] or invertebrate hosts [24,25,26,27]. However, and certainly because of the difficulties inherent to the studied model—including Babesia spp. culture, tick colony maintenance, and animal models—relatively few studies have been based on the establishment, under experimental conditions, of complete parasite transmission cycles from one vertebrate host to another via the tick bite. Thanks to the experimental models developed for that purpose, these few studies have nevertheless made significant advances in (1) the definition/confirmation of the vector competence of various tick species; (2) the understanding of the modalities of parasite acquisition and transmission by ticks; (3) the discovery of the molecular interactions between the parasite and its invertebrate hosts; (4) the evaluation of some control methods. The aim of this review is to summarize studies that include both tick infection on Babesia-infected animals and Babesia infection of ticks through artificial systems, and to comment on the major results they achieved.

2. General Description of the Babesia Life Cycle

Babesia spp. are transmitted by hard tick (ixodid) vectors. The tick vectors and reservoir hosts differ depending on Babesia sp. and geographical location considered [1]. Several tick species have been mentioned in the literature as vectors of Babesia sp., but, as shown in Table 1, vector competence through experimental transmission has not been validated for all of them. For those mentioned here as "suspected vectors" without realization of the complete transmission cycle under experimental conditions, their involvement is mostly based on epidemiological evidence (e.g., correlation of tick species presence with disease occurrence). Each of the three active stages of hard ticks (larva, nymph, and adult) takes a single blood meal from a vertebrate host in order to mature to the next stage or lay eggs for the female. Most of the tick species listed as confirmed or suspected vectors of some Babesia species are three-hosts ticks—meaning that they take their three blood meals on three different hosts. Some of them, however, have a two-host cycle such as some Hyalomma spp., Rhipicephalus evertsi, and Rhipicephalus bursa, whereas Dermacentor nitens, Rhipicephalus annulatus, Rhipicephalus microplus, and Rhipicephalus decoloratus are one-host ticks that use the same individual host animal for all active tick stages.
The Babesia life cycle includes both asexual multiplication in the erythrocytes of the vertebrate host and sexual reproduction in the tick vector [3,61]. The general life cycle of the Babesia species is summarized in Figure 1 for Babesia sensu stricto (s.s.) species. Indeed, it is necessary to specify here that, quite recently, molecular phylogeny studies using the 18S rRNA gene have led to the division of Babesia species into two large groups: Babesia s.s. and Babesia sensu lato (s.l.). Species belonging to the latter group, such as B. microti, are not capable of transovarial transmission within the tick but only a transstadial mode of transmission [62,63]. Vertebrate hosts are infected by the injection of sporozoites present in tick saliva during the tick bite. Each sporozoite penetrates the cell membrane of an erythrocyte with the aid of a specialized apical complex. Once inside, the parasite produces two merozoites by a process of merogony. Merozoites are then intermittently released following erythrocyte lysis to infect new erythrocytes. The parasite may then persist asymptomatically within its host for several years or lead to acute disease. When they are ingested by the tick during the blood meal, some parasites present in infected erythrocytes (pre-gametocytes) undergo further development in the passage from host blood to the midgut of the tick vector to evolve into gametocytes. The sexual reproduction between gametocytes takes place in the tick gut and leads to a zygote that penetrates the gut epithelium, where further multiplication occurs, with development to motile and haploid kinetes that escape into the tick hemolymph. The kinetes then infect a variety of tick cell types and tissues, including the ovary in the female tick—for Babesia s.s. species—and the tick salivary glands, where successive cycles of asexual multiplication take place. In this last organ, sporozoite development usually only begins when the infected tick attaches to the vertebrate host. The ticks thus transmit the sporozoites to a new host during a new blood meal of the next life-stage for the ticks with several hosts or of the next generation after transovarian transmission for the one-host ticks.

3. Experimental Models of Babesia Species—Transmission by Ticks

3.1. Tick Infestations on Babesia spp. Infected Animals

The first experiments to transmit Babesia spp. naturally were carried out by applying ticks suspected of being vectors on infected animals. Due to the huge economic importance of bovine babesiosis, these studies on vector competence for Babesia sp. were first conducted on species that infect cattle. Indeed, in 1893, two American researchers, Smith and Kilborne—the first authors to demonstrate the transmission of a disease organism from an arthropod to a mammalian host—showed the vector competence of R. annulatus for B. bigemina by placing ticks infected on animal onto naïve cattle that developed the associated disease [42]. Thereafter, different species of animals were used depending on the species of Babesia studied. Over time, the animal models and methodology used were refined to optimize the infection of both animals and ticks, and to comply with health and safety rules, and animal accommodation and tick containment methods during the tick feeding process have been the subject of several tests and evolutions (see examples in Figure 2A–F). In most cases, for cattle, animals were housed in individual, tick-proof pens surrounded by moats with or without detergent or insecticide [43,45]. Concerning the tick containment methods, in 1961, Callow and Hoyte used a hessian rug to protect the larvae until adult repletion to demonstrate the transmission of B. bigemina to cattle by R. microplus [43]. Ticks were either allowed to spread at will over the animal, or were confined to one site by releasing them under a fabric patch (nylon or organdie), which was glued along its edges to the flank of the bovine. A few years later, for the demonstration of the vector competence of R. decoloratus for B. bigemina in Kenya, ticks were fed on cattle until adult repletion by sprinkling larvae on the backs of the animals [45]. For the first experimental transmission of B. divergens by Ixodes ricinus achieved by Joyner et al in 1963, the ticks were contained in ear bags [36], a method also used later for sheep [64]. Regarding rodents, several laboratory studies involving complete transmission cycles of B. microti to the vertebrate host through the tick bite were performed [28,29,30,31,32,33]. In most instances, rodents were maintained over trays of water from which detached engorged ticks—applied by brush to animals—were harvested, whereas in some cases, ticks were contained in plastic capsules attached with different adhesives. For horses, the first studies aimed to validate the vector competence of D. nitens for B. caballi used larvae applied by brush on the animal [51], whereas in subsequent studies, tick feedings were accomplished by placing the larvae under a cloth patch glued to the back of the host [52].
The identification of a pathogen or pathogen DNA alone—which is even less convincing—in an arthropod cannot be sufficient to prove its ability to transmit this pathogen. Indeed, demonstrations of parasite presence in unfed field ticks, in tick salivary glands, eggs, or unfed larvae, while more convincing than detection in ticks collected from animals, also require confirmation only provided by the validation of vector competence in a controlled experimental model. The best illustration of this corresponds to the following studies performed in Iran in order to identify the vector of B. ovis to sheep. The kinetes of B. ovis were observed in hemolymph and egg smears of Rhipicephalus sanguineus and Hyalomma marginatum field ticks collected from sheep infected with B. ovis [69], so the vector competence of both tick species was further evaluated by placing pairs of adult ticks on sheep inoculated with B. ovis, but no transmission by any of the succeeding tick stages could be demonstrated, thus showing that these tick species are not vectors [64].
As recently reviewed by Gray and co-workers, the establishment of experimental models of pathogens transmission by ticks using live animals has led to significant advances in the understanding of transmission modalities and tick-parasite interactions [70]. For example, it allowed the demonstration that transovarial transmission does not occur in B. microti, leading to no longer classifying this parasite in Babesia spp. s.s. [31,32,33], and that, in I. ricinus, the infection only survives one molt [31]. Laboratory models have also demonstrated that B. microti may promote its transmission in rodents by enhancing the feeding success and survival of its tick vector, Ixodes trianguliceps [29]. Likewise, the establishment of the transmission of B. divergens by I. ricinus in a gerbil experimental model [35] provided proof of sexual development of Babesia through DNA measurements on the developmental stages of B. divergens in the blood of the vertebrate host and in the gut, hemolymph, and salivary glands of the tick vector [38,39]. In addition, several studies have been performed in order to establish which tick life stages are able to acquire and/or to transmit the parasite. Most of them have concluded that only adult stages were able to acquire Babesia sp. s.s. from infected animals, while all succeeding stages (larvae, nymphs, and adults) were able to re-transmit the parasites to susceptible animals [36,37,43,44,48]. However, Schwint and co-workers demonstrated that only the first of three subsequent generations from D. nitens females was able to transmit B. caballi to naïve horses, showing that the parasite is unable to persist in ticks without continuing alimentary infection of adult females [52], whereas other studies have also shown the acquisition of this parasite by nymphs of R. evertsi [53].
Models of transmission of B. microti to rodents via ticks in the laboratory have also made it possible to carry out studies on the phenomena of co-infections. In fact, a lower transmission efficiency of B. microti than Borrelia burgdorferi to Ixodes dammini from both hamsters [71] and white-footed mice [72] has been demonstrated. In the meantime, it was shown that ticks that fed on mice with these concurrent pathogen infections exhibited twice the incidence of B. burgdorferi infection compared with B. microti [72]. Twenty-five years later, however, this laboratory model also showed an increase of the frequency of B. microti-infected Ixodes scapularis (formerly I. dammini) nymphs when they fed as larvae on white-footed mice coinfected with B. burgdorferi, as well as an increase of B. microti parasitemia in co-infected mice [28]. This enhancement of B. microti establishment by B. burgdorferi has been attributed to an immunological conflict in the adaptive immune response of the vertebrate host against the two tick-borne pathogens [73].
The development of experimental models using animals also provided knowledge on the infection acquisition by the vertebrate host following a Babesia-infected tick bite. For example, by developing a laboratory model of B. bovis infection of calves through the bite of R. microplus-infected ticks, Smith and co-workers demonstrated, in 1978, that tick-induced infection was more severe than in calves infected with carrier blood, even when very low numbers of infected larvae were applied [41]. They attributed this difference in virulence to the large number of infective doses injected by each infected tick but Salivary-Assisted Transmission of tick-borne pathogens (see review in [74]) probably also contributed to this observation.
Although, compared to other pathogens, few molecular studies have involved Babesia parasites [70,75], experimental tick-transmission models of Babesia spp. using animals have also made it possible to identify molecules potentially involved in this transmission. Such studies are helping us to better understand the interactions involved and to identify potential targets for blocking parasite transmission. Thus, infection of H. longicornis on dogs infected with B. gibsoni has allowed to implicate a tick protein, longipain, in the transmission of the parasite by its vector [76], and to demonstrate that the vitellogenin receptor on the surface of tick oocytes is essential for its transovarial transmission [77]. Experimental models of B. bovis infection of cattle by R. microplus has also been used to study the function of the tick protein Bm86 during B. bovis infection [78]. In the same way, experimental tick infection models of both B. bigemina and B. bovis-infected cattle were used to perform functional genomics studies on R. annulatus and R. microplus genes that are differentially expressed in response to parasite infection [79,80].
Both the discovery and validation of methods to control the transmission of tick-borne pathogens require experimental designs that include complete transmission cycles. Regarding the Babesia spp. that infect cattle, experimental tick infection models for B. bigemina-infected cattle were used to evaluate the efficiency of some vaccine candidates and drugs against parasite transmission by R. microplus [81,82]. Rhipicephalus microplus experimentally infected with B. bovis were also used to demonstrate the inefficiency of the injectable and pour-on forms of both ivermectin and moxidectin to prevent parasite transmission by ticks [82]. In Argentina, Mangold and co-workers developed a laboratory model of B. bovis transmission to cattle by R. microplus in order to demonstrate the non-transmissibility to ticks of an attenuated vaccine strain of the parasite [83]. Several studies also involved experimental transmission of B. canis to dogs through the bite of infected D. reticulatus ticks in order to evaluate the usefulness of different acaricides to prevent parasite transmission [55,56,57,84], whereas similar experiments were performed regarding B. canis transmission by R. sanguineus [85,86], and B. rossi transmission by Haemaphysalis elliptica [60].

3.2. Tick Infection through Artificial Feeding Systems

The use of natural hosts for direct infection of ticks on infectious animals remains the best method to obtain conditions that are closest to the physiological reality of tick-borne pathogen transmission. However, firstly, ethical considerations lead us to limit the use of animals as much as possible. Secondly, in addition to the constraints associated with licensing the experiments and host specificity, obtaining the animals, keeping them in the laboratory, and handling them can be expensive and difficult or even impossible in the case of most species of wildlife. Finally, the difficulty of controlling parasitaemias of infected animals for the whole feeding period of the ticks is an important consideration. All these reasons have led to the development of artificial methods of tick infection in order to complete the lifecycle of tick-borne pathogens under laboratory conditions. Artificial feeding of ticks, mimicking the natural process, has been used for different purposes including tick rearing, the study of tick physiology and the effects of antibodies or drugs on tick physiology, functional genomic studies, and vaccine candidate discovery, as well as the analysis of tick-borne pathogen transmission (see review by Bonnet and Liu [13]). Nevertheless, relatively few studies have involved Babesia parasites. One of the factors limiting these studies is undoubtedly the need for successful in vitro cultivation of the Babesia species of interest [15].
The use of blood-filled capillary tubes placed over the mouthparts of ticks was first reported in 1938 by Gregson, who used this technique to collect saliva from D. andersoni [87]. Since then, this technique has been used to infect ticks with several tick-borne pathogens, mainly bacteria (see review in [13]). Although this infection process has the advantage of using the natural route of infection, i.e., the digestive tract of the tick, and allows control of the quantity of pathogen ingested, it is quite far from natural conditions because the tick absorbs a large amount of pathogen at once and this, regardless of the "true" full blood meal. The other drawback is that it requires the use of animals before or after infection to feed ticks to repletion. However, its use has led to some significant advances in the study of Babesia spp. transmission. In 1998, Inokuma and Kemp used the capillary feeding technique in order to infect R. microplus with B. bigemina [88]. Adult ticks were pre-fed on cattle and then B. bigemina-infected red blood cells were offered to the ticks using glass capillaries on a warm plate at 35 °C for 18 h in the dark. The authors subsequently demonstrated that ticks were able to acquire the parasite and to transmit it to their progeny. The capillary feeding technique has also been used by Antunes and co-workers in order to evaluate the impact of purified rabbit polyclonal antibodies on some R. microplus proteins associated with both tick physiology and tick infection by B. bigemina [89]. While an effect on tick weight and oviposition was observed, no effect was observed on pathogen DNA levels. However, this work has made it possible to set up an alternative system to the use of animals both to test vaccine candidates and to obtain essential data on tick-pathogen interaction, as this could also be done later with another protein, the calreticulin, identified as being involved in B. bigemina infection in R. annulatus ticks [90].
The membrane feeding technique—consisting of feeding ticks on blood or culture media through a membrane—was first developed by Pierce and Pierce in 1956 in order to feed R. microplus using embryonated hen eggs [91]. Since then, several membranes of animal or artificial origin have been used both to feed and to infect ticks with different pathogens (see review in [13]). In this case, as the pathogen is mixed in blood and absorbed throughout the blood meal via the digestive tract, the method mimics the natural conditions of tick infection more closely than other methods. Its main disadvantages, however, are the need to regularly change the blood used, and the need to use antibiotics and antifungals to avoid contamination. It is also necessary to test pathogen viability under these feeding conditions at regular intervals. In the case of artificial membranes such as silicone ones, feeding systems also need olfactory stimuli for attachment and feeding. Regarding their use to infect ticks with Babesia spp., and due to red blood cell sedimentation, it is important to note that this requires the placing of blood above the membrane to produce a continuous gravitational pressure, ensuring tick absorption of the intraerythrocytic parasites. In 2007, Bonnet and co-workers developed a skin-feeding technique using the skins of both gerbils (for larvae and nymphs) and rabbits (for nymphs and adults) to infect I. ricinus with B. divergens without the need for additional stimuli (Figure 2G,H) [35]. To our knowledge, this is the only membrane-feeding technique that has been used to date to infect ticks with Babesia spp. All tick instars were allowed to acquire the parasite from infected red blood cells maintained at 37 °C in a glass feeder through the animal skin until repletion. Contrary to what was previously observed [36,37], this system showed that in addition to adult females, I. ricinus larvae and nymphs can also acquire B. divergens infections, which persists transtadially in the subsequent nymphal and adult stages (as determined by the detection of DNA in their salivary glands). Babesia divergens DNA was also detected in eggs and larvae produced by females that had fed on parasitized blood, demonstrating the transovarial transmission of the parasite. Later, the use of this artificial tick infection system also allowed the discovery of molecular markers for B. divergens sexual stages [92,93]. Lastly, the same membrane feeding technique allowed the validation of the vector competence of I. ricinus for Babesia (EU1) venatorum [40].
Finally, in order to understand tick-Babesia interactions and to follow parasite development in the vector, Maeda and co-workers used a “semi-artificial” mouse skin membrane feeding technique to infect H. longicornis with B. ovata [94]. In this case, female adult ticks were first allowed to feed on the shaved back of mice, and after 4 to 5 days, a section of the mouse skin with the ticks attached was removed immediately after euthanasia, and set up in artificial feeding units. The ticks were then fed on a mix of media and B. ovata-infected red blood cells through the piece of mouse skin. This technique was then used to demonstrate the transovarial persistence of B. ovata DNA in H. longicornis [50]. Thus, the mixing of animal use and membrane feeding makes it possible to control the parasitemia of the meal offered to ticks, but does not prevent the use of live animals.

3.3. Tick Infection through Injection

Although this method is more distant from physiological reality than the ones previously detailed, some studies have also performed tick infections by injecting the pathogen through the cuticle of the tick. In addition to requiring live animals to feed ticks after the infection, this invasive method also has the disadvantage of a low survival rate of ticks after injection [95]. Nevertheless, its use by some authors has led to important results concerning Babesia spp. In 2018, Antunes and co-workers used a R. bursa-B. ovis-sheep infection model to characterize tick salivary gland genes that were differentially expressed in response to blood feeding and B. ovis infection [96]. In that experiment, female ticks were inoculated with B. ovis in the trochanter-coxae articulation and allowed to feed on rabbits. Vector competence was then confirmed by feeding B. ovis-infected ticks on a naïve lamb. This study allowed both increased understanding of the role of tick salivary gland genes in Babesia infections and identification of potential candidate vaccine antigens for innovative control strategies.

4. Conclusions

It is likely that vector competence for Babesia spp. has yet to be determined in some tick species. Furthermore, our understanding of the life cycles of Babesia spp. is still incomplete, especially regarding the intimate mechanisms of molecular dialogue between the parasite and its vertebrate and invertebrate hosts. Field experiments are less easy to control than those under laboratory conditions, and the gaps in our knowledge can probably only be filled by experimentally reproducing the transmission cycle as closely as possible to reality and by involving all three actors of the vectorial system. Indeed, this review has shown that the use of experimental systems for tick-borne pathogen infections that permit the complete transmission cycles of Babesia parasites has led to major scientific advances in the study of these pathogens. Despite these successes, efforts are still needed to standardize and simplify laboratory protocols to improve our ability to exploit tick artificial infection systems. It is hoped that, in the future, such models of artificial infection will be further developed in order to acquire new knowledge and develop new control strategies while avoiding the use of animals.

Author Contributions

Conceptualization, S.I.B.; formal analysis, S.I.B. and C.N.; writing—original draft preparation, S.I.B. and C.N.; writing—review and editing, S.I.B. and C.N.; supervision, S.I.B. All authors have read and agreed to the published version of the manuscript.

Funding

The PhD of CN is funded by the Institut Français du Cheval et de l’Equitation (IFCE) and by the Fonds Eperon.

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

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]
  2. Lobo, C.A.; Singh, M.; Rodriguez, M. Human babesiosis: Recent advances and future challenges. Curr. Opin. Hematol. 2020, 27, 399–405. [Google Scholar] [CrossRef]
  3. Chauvin, A.; Moreau, E.; Bonnet, S.; Plantard, O.; Malandrin, L. Babesia and its hosts: Adaptation to long-lasting interactions as a way to achieve efficient transmission. Vet. Res. 2009, 40, 37. [Google Scholar] [CrossRef] [Green Version]
  4. Krause, P.J. Human babesiosis. Int. J. Parasitol. 2019, 49, 165–174. [Google Scholar] [CrossRef] [PubMed]
  5. Bock, R.; Jackson, L.; de Vos, A.; Jorgensen, W. Babesiosis of cattle. Parasitology 2004, 129, S247–S269. [Google Scholar] [CrossRef] [PubMed]
  6. Yeruham, I.; Hadani, A.; Galker, F. Some epizootiological and clinical aspects of ovine babesiosis caused by Babesia ovis—A review. Vet. Parasitol. 1998, 74, 153–163. [Google Scholar] [CrossRef]
  7. Tirosh-Levy, S.; Gottlieb, Y.; Fry, L.M.; Knowles, D.P.; Steinman, A. Twenty Years of Equine Piroplasmosis Research: Global Distribution, Molecular Diagnosis, and Phylogeny. Pathogens 2020, 9, 926. [Google Scholar] [CrossRef] [PubMed]
  8. Nadal, C.; Bonnet, S.I.; Marsot, M. Eco-epidemiology of equine piroplasmosis and its associated tick vectors in Europe: A systematic literature review and a meta-analysis of prevalence. Transbound. Emerg. Dis. 2021. [Google Scholar] [CrossRef]
  9. Knowles, D.P.; Kappmeyer, L.S.; Haney, D.; Herndon, D.R.; Fry, L.M.; Munro, J.B.; Sears, K.; Ueti, M.W.; Wise, L.N.; Silva, M.; et al. Discovery of a novel species, Theileria haneyi n. sp., infective to equids, highlights exceptional genomic diversity within the genus Theileria: Implications for apicomplexan parasite surveillance. Int. J. Parasitol. 2018, 48, 679–690. [Google Scholar] [CrossRef]
  10. Bilić, P.; Kuleš, J.; Barić, R.; Mrljak, V. Canine Babesiosis: Where Do We Stand? Acta Vet. 2018, 68, 127–160. [Google Scholar] [CrossRef] [Green Version]
  11. Suarez, C.E.; Noh, S. Emerging perspectives in the research of bovine babesiosis and anaplasmosis. Vet. Parasitol. 2011, 180, 109–125. [Google Scholar] [CrossRef] [PubMed]
  12. Rodriguez-Morales, A.J.; Bonilla-Aldana, D.K.; Escalera-Antezana, J.P.; Alvarado-Arnez, L.E. Research on Babesia: A bibliometric assessment of a neglected tick-borne parasite. F1000Res 2018, 7, 1987. [Google Scholar] [CrossRef]
  13. Bonnet, S.I.; Liu, X.Y. Laboratory artificial infection of hard ticks: A tool for the analysis of tick-borne pathogen transmission. Acarologia 2012, 52, 453–464. [Google Scholar] [CrossRef] [Green Version]
  14. de la Fuente, J.; Antunes, S.; Bonnet, S.; Cabezas-Cruz, A.; Domingos, A.G.; Estrada-Pena, A.; Johnson, N.; Kocan, K.M.; Mansfield, K.L.; Nijhof, A.M.; et al. Tick-Pathogen Interactions and Vector Competence: Identification of Molecular Drivers for Tick-Borne Diseases. Front. Cell Infect. Microbiol. 2017, 7, 114. [Google Scholar] [CrossRef] [Green Version]
  15. Muller, J.; Hemphill, A. In vitro culture systems for the study of apicomplexan parasites in farm animals. Int. J. Parasitol. 2013, 43, 115–124. [Google Scholar] [CrossRef] [PubMed]
  16. Sondgeroth, K.S.; McElwain, T.F.; Ueti, M.W.; Scoles, G.A.; Reif, K.E.; Lau, A.O. Tick passage results in enhanced attenuation of Babesia bovis. Infect. Immun. 2014, 82, 4426–4434. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  17. Stewart, N.P. Differences in the life cycles between a vaccine strain and an unmodified strain of Babesia bovis (Babes, 1889) in the tick Boophilus microplus (Canestrini). J. Protozool. 1978, 25, 497–501. [Google Scholar] [CrossRef]
  18. Zaugg, J.L.; Kuttler, K.L. Experimental infections of Babesia bigemina in American bison. J. Wildl. Dis. 1987, 23, 99–102. [Google Scholar] [CrossRef] [Green Version]
  19. Chauvin, A.; Valentin, A.; Malandrin, L.; L’Hostis, M. Sheep as a new experimental host for Babesia divergens. Vet. Res. 2002, 33, 429–433. [Google Scholar] [CrossRef] [Green Version]
  20. Martinez-Garcia, G.; Santamaria-Espinosa, R.M.; Lira-Amaya, J.J.; Figueroa, J.V. Challenges in Tick-Borne Pathogen Detection: The Case for Babesia spp. Identification in the Tick Vector. Pathogens 2021, 10, 92. [Google Scholar] [CrossRef]
  21. Sun, Y.; Moreau, E.; Chauvin, A.; Malandrin, L. The invasion process of bovine erythrocyte by Babesia divergens: Knowledge from an in vitro assay. Vet. Res. 2011, 42, 62. [Google Scholar] [CrossRef] [Green Version]
  22. Spencer, A.M.; Goethert, H.K.; Telford, S.R., 3rd; Holman, P.J. In vitro host erythrocyte specificity and differential morphology of Babesia divergens and a zoonotic Babesia sp. from eastern cottontail rabbits (Sylvilagus floridanus). J. Parasitol. 2006, 92, 333–340. [Google Scholar] [CrossRef] [PubMed]
  23. O’Connor, R.M.; Long, J.A.; Allred, D.R. Cytoadherence of Babesia bovis-infected erythrocytes to bovine brain capillary endothelial cells provides an in vitro model for sequestration. Infect. Immun. 1999, 67, 3921–3928. [Google Scholar] [CrossRef] [Green Version]
  24. Maeda, H.; Hatta, T.; Alim, M.A.; Tsubokawa, D.; Mikami, F.; Kusakisako, K.; Matsubayashi, M.; Umemiya-Shirafuji, R.; Tsuji, N.; Tanaka, T. Initial development of Babesia ovata in the tick midgut. Vet. Parasitol. 2017, 233, 39–42. [Google Scholar] [CrossRef] [PubMed]
  25. Bhat, U.K.; Mahoney, D.F.; Wright, I.G. The invasion and growth of Babesia bovis in tick tissue culture. Experientia 1979, 35, 752–753. [Google Scholar] [CrossRef] [PubMed]
  26. Ribeiro, M.F.; Bastos, C.V.; Vasconcelos, M.M.; Passos, L.M. Babesia bigemina: In vitro multiplication of sporokinetes in Ixodes scapularis (IDE8) cells. Exp. Parasitol. 2009, 122, 192–195. [Google Scholar] [CrossRef] [PubMed]
  27. de Rezende, J.; Rangel, C.P.; McIntosh, D.; Silveira, J.A.; Cunha, N.C.; Ramos, C.A.; Fonseca, A.H. In vitro cultivation and cryopreservation of Babesia bigemina sporokinetes in hemocytes of Rhipicephalus microplus. Vet. Parasitol. 2015, 212, 400–403. [Google Scholar] [CrossRef]
  28. Dunn, J.M.; Krause, P.J.; Davis, S.; Vannier, E.G.; Fitzpatrick, M.C.; Rollend, L.; Belperron, A.A.; States, S.L.; Stacey, A.; Bockenstedt, L.K.; et al. Borrelia burgdorferi promotes the establishment of Babesia microti in the northeastern United States. PLoS ONE 2014, 9, e115494. [Google Scholar] [CrossRef]
  29. Randolph, S.E. The effect of Babesia microti on feeding and survival in its tick vector, Ixodes trianguliceps. Parasitology 1991, 102 Pt 1, 9–16. [Google Scholar] [CrossRef]
  30. Ruebush, T.K., 2nd; Piesman, J.; Collins, W.E.; Spielman, A.; Warren, M. Tick transmission of Babesia microti to rhesus monkeys (Macaca mulatta). Am. J. Trop. Med. Hyg. 1981, 30, 555–559. [Google Scholar] [CrossRef]
  31. Gray, J.; von Stedingk, L.V.; Gurtelschmid, M.; Granstrom, M. Transmission studies of Babesia microti in Ixodes ricinus ticks and gerbils. J. Clin. Microbiol. 2002, 40, 1259–1263. [Google Scholar] [CrossRef] [Green Version]
  32. Walter, G.; Weber, G. A study on the transmission (transstadial, transovarial) of Babesia microti, strain "Hannover i", in its tick vector, Ixodes ricinus (author’s transl). Tropenmed. Parasitol. 1981, 32, 228–230. [Google Scholar] [PubMed]
  33. Li, L.H.; Zhu, D.; Zhang, C.C.; Zhang, Y.; Zhou, X.N. Experimental transmission of Babesia microti by Rhipicephalus haemaphysaloides. Parasites Vectors 2016, 9, 231. [Google Scholar] [CrossRef] [Green Version]
  34. Wu, J.; Cao, J.; Zhou, Y.; Zhang, H.; Gong, H.; Zhou, J. Evaluation on Infectivity of Babesia microti to Domestic Animals and Ticks Outside the Ixodes Genus. Front. Microbiol. 2017, 8, 1915. [Google Scholar] [CrossRef] [PubMed]
  35. 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]
  36. 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]
  37. 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]
  38. Lewis, D.; Young, E.R. The transmission of a human strain of Babesia divergens by Ixodes ricinus ticks. J. Parasitol. 1980, 66, 359–360. [Google Scholar] [CrossRef]
  39. 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]
  40. Bonnet, S.; Brisseau, N.; Hermouet, A.; Jouglin, M.; Chauvin, A. Experimental in vitro transmission of Babesia sp. (EU1) by Ixodes ricinus. Vet. Res. 2009, 40, 21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  41. Smith, R.D.; Osorno, B.M.; Brener, J.; De La Rosa, R.; Ristic, M. Bovine babesiosis: Severity and reproducibility of Babesia bovis infections induced by Boophilus microplus under laboratory conditions. Res. Vet. Sci. 1978, 24, 287–292. [Google Scholar] [CrossRef]
  42. Smith, T.; Kilborne, F.L. Investigations into the nature, causation and prevention of Southern cattle fever. In Ninth Annual Report of the Bureau of Animal Industry for the Year 1892; Government Printing Office: Washington, DC, USA, 1893; pp. 177–304. [Google Scholar]
  43. Callow, L.; Hoyte, H. Transmission experiments using Babesia bigemina, Theileria mutans, Borrelia sp. and the cattle tick, Boophilus microplus. Aust. Vet. J. 1961, 37, 381–390. [Google Scholar] [CrossRef]
  44. Callow, L.L. Babesia Bigemina in Ticks Grown on Non-Bovine Hosts and Its Transmission to These Hosts. Parasitology 1965, 55, 375–381. [Google Scholar] [CrossRef]
  45. Morzaria, S.P.; Young, A.S.; Hudson, E.B. Babesia bigemina in Kenya: Experimental transmission by Boophilus decoloratus and the production of tick-derived stabilates. Parasitology 1977, 74, 291–298. [Google Scholar] [CrossRef]
  46. Akinboade, O.A.; Dipeolu, O.O.; Adetunji, A. Experimental transmission of Babesia bigemina and Anaplasma marginale to calves with the larvae of Boophilus decoloratus. Zentralbl. Veterinarmed. B 1981, 28, 329–332. [Google Scholar] [CrossRef]
  47. Akinboade, O.A. Experimental transmission of Babesia bigemina in sheep using infective larval tick of Boophilus decoloratus. Rev. Elev. Med. Vet. Pays. Trop. 1981, 34, 271–273. [Google Scholar]
  48. Morzaria, S.P.; Brocklesby, D.W.; Harradine, D.L. Experimental transmission of Babesia major by Haemaphysalis punctata. Res. Vet. Sci. 1977, 23, 261–262. [Google Scholar] [CrossRef]
  49. Yin, H.; Lu, W.; Luo, J.; Zhang, Q.; Lu, W.; Dou, H. Experiments on the transmission of Babesia major and Babesia bigemina by Haemaphysalis punctata. Vet. Parasitol. 1996, 67, 89–98. [Google Scholar] [CrossRef]
  50. Umemiya-Shirafuji, R.; Hatta, T.; Okubo, K.; Sato, M.; Maeda, H.; Kume, A.; Yokoyama, N.; Igarashi, I.; Tsuji, N.; Fujisaki, K.; et al. Transovarial persistence of Babesia ovata DNA in a hard tick, Haemaphysalis longicornis, in a semi-artificial mouse skin membrane feeding system. Acta Parasitol. 2018, 63, 433. [Google Scholar] [CrossRef]
  51. Roby, T.O.; Anthony, D.W.; Thornton, C.W., Jr.; Holbrook, A.A. The Hereditary Transmission of Babesia Caballi in the Tropical Horse Tick, Dermacentor Nitens Neumann. Am. J. Vet. Res. 1964, 25, 494–499. [Google Scholar]
  52. Schwint, O.N.; Knowles, D.P.; Ueti, M.W.; Kappmeyer, L.S.; Scoles, G.A. Transmission of Babesia caballi by Dermacentor nitens (Acari: Ixodidae) is restricted to one generation in the absence of alimentary reinfection on a susceptible equine host. J. Med. Entomol. 2008, 45, 1152–1155. [Google Scholar] [CrossRef] [Green Version]
  53. de Waal, D.T.; Potgieter, F.T. The transstadial transmission of Babesia caballi by Rhipicephalus evertsi evertsi. Onderstepoort. J. Vet. Res. 1987, 54, 655–656. [Google Scholar]
  54. Erster, O.; Roth, A.; Wolkomirsky, R.; Leibovich, B.; Savitzky, I.; Shkap, V. Transmission of Babesia ovis by different Rhipicephalus bursa developmental stages and infected blood injection. Ticks Tick Borne Dis. 2016, 7, 13–19. [Google Scholar] [CrossRef]
  55. Jongejan, F.; Fourie, J.J.; Chester, S.T.; Manavella, C.; Mallouk, Y.; Pollmeier, M.G.; Baggott, D. The prevention of transmission of Babesia canis canis by Dermacentor reticulatus ticks to dogs using a novel combination of fipronil, amitraz and (S)-methoprene. Vet. Parasitol. 2011, 179, 343–350. [Google Scholar] [CrossRef] [PubMed]
  56. Beugnet, F.; Halos, L.; Larsen, D.; Labuschagne, M.; Erasmus, H.; Fourie, J. The ability of an oral formulation of afoxolaner to block the transmission of Babesia canis by Dermacentor reticulatus ticks to dogs. Parasites Vectors 2014, 7, 283. [Google Scholar] [CrossRef] [Green Version]
  57. Taenzler, J.; Liebenberg, J.; Roepke, R.K.; Heckeroth, A.R. Prevention of transmission of Babesia canis by Dermacentor reticulatus ticks to dogs after topical administration of fluralaner spot-on solution. Parasites Vectors 2016, 9, 234. [Google Scholar] [CrossRef] [Green Version]
  58. Fourie, J.J.; de Vos, C.; Crafford, D.; Pollmeier, M.; Schunack, B. A study on the long-term efficacy of Seresto(R) collars in preventing Babesia canis (Piana & Galli-Valerio, 1895) transmission to dogs by infected Dermacentor reticulatus (Fabricius, 1794) ticks. Parasites Vectors 2019, 12, 139. [Google Scholar] [CrossRef]
  59. Varloud, M.; Liebenberg, J.; Fourie, J. Early Babesia canis transmission in dogs within 24 h and 8 h of infestation with infected pre-activated male Dermacentor reticulatus ticks. Parasites Vectors 2018, 11, 41. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  60. Beugnet, F.; Lebon, W.; de Vos, C. Prevention of the transmission of Babesia rossi by Haemaphysalis elliptica in dogs treated with Nexgard((R)). Parasite 2019, 26, 49. [Google Scholar] [CrossRef] [Green Version]
  61. Mehlhorn, H.; Shein, E. The piroplasms: Life cycle and sexual stages. Adv. Parasitol. 1984, 23, 37–103. [Google Scholar] [CrossRef]
  62. Gray, J.; Weiss, M.L. Babesia microti. In Emerging Protozoan Pathogens; Naveed Ahmed, K., Ed.; Taylor & Francis: New York, NY, USA, 2007; pp. 303–349. [Google Scholar]
  63. Jalovecka, M.; Sojka, D.; Ascencio, M.; Schnittger, L. Babesia Life Cycle—When Phylogeny Meets Biology. Trends Parasitol. 2019, 35, 356–368. [Google Scholar] [CrossRef]
  64. Razmi, G.; Nouroozi, E. Transovarial Transmission of Babesia ovis by Rhipicephalus sanguineus and Hyalomma marginatum. Iran J. Parasitol. 2010, 5, 35–39. [Google Scholar]
  65. Bonnet, S.; Choumet, V.; Masseglia, S.; Cote, M.; Ferquel, E.; Lilin, T.; Marsot, M.; Chapuis, J.L.; Vourc’h, G. Infection of Siberian chipmunks (Tamias sibiricus barberi) with Borrelia sp. reveals a low reservoir competence under experimental conditions. Ticks Tick Borne Dis. 2015, 6, 393–400. [Google Scholar] [CrossRef]
  66. Almazan, C.; Simo, L.; Fourniol, L.; Rakotobe, S.; Borneres, J.; Cote, M.; Peltier, S.; Maye, J.; Versille, N.; Richardson, J.; et al. Multiple Antigenic Peptide-Based Vaccines Targeting Ixodes ricinus Neuropeptides Induce a Specific Antibody Response but Do Not Impact Tick Infestation. Pathogens 2020, 9, 900. [Google Scholar] [CrossRef] [PubMed]
  67. Almazan, C.; Fourniol, L.; Rouxel, C.; Alberdi, P.; Gandoin, C.; Lagree, A.C.; Boulouis, H.J.; de la Fuente, J.; Bonnet, S.I. Experimental Ixodes ricinus-Sheep Cycle of Anaplasma phagocytophilum NV2Os Propagated in Tick Cell Cultures. Front. Vet. Sci. 2020, 7, 40. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  68. Almazan, C.; Bonnet, S.; Cote, M.; Slovak, M.; Park, Y.; Simo, L. A Versatile Model of Hard Tick Infestation on Laboratory Rabbits. J. Vis. Exp. 2018. [Google Scholar] [CrossRef]
  69. Razmi, G.R.; Naghibi, A.; Aslani, M.R.; Fathivand, M.; Dastjerdi, K. An epidemiological study on ovine babesiosis in the Mashhad suburb area, province of Khorasan, Iran. Vet. Parasitol. 2002, 108, 109–115. [Google Scholar] [CrossRef]
  70. Gray, J.S.; Estrada-Pena, A.; Zintl, A. Vectors of Babesiosis. Annu. Rev. Entomol. 2019, 64, 149–165. [Google Scholar] [CrossRef]
  71. Piesman, J. Intensity and duration of Borrelia burgdorferi and Babesia microti infectivity in rodent hosts. Int. J. Parasitol. 1988, 18, 687–689. [Google Scholar] [CrossRef]
  72. Mather, T.N.; Telford, S.R., 3rd; Moore, S.I.; Spielman, A. Borrelia burgdorferi and Babesia microti: Efficiency of transmission from reservoirs to vector ticks (Ixodes dammini). Exp. Parasitol. 1990, 70, 55–61. [Google Scholar] [CrossRef]
  73. Diuk-Wasser, M.A.; Vannier, E.; Krause, P.J. Coinfection by Ixodes Tick-Borne Pathogens: Ecological, Epidemiological, and Clinical Consequences. Trends Parasitol. 2016, 32, 30–42. [Google Scholar] [CrossRef] [Green Version]
  74. Bonnet, S.; Kazimirova, M.; Richardson, R.; Simo, L. Tick saliva and its role in pathogen transmission. In Skin and Arthropod Vectors; Boulanger, N., Ed.; Elsevier: London, UK, 2018. [Google Scholar]
  75. Liu, X.Y.; Bonnet, S.I. Hard tick factors implicated in pathogen transmission. PLoS Negl. Trop. Dis. 2014, 8, e2566. [Google Scholar] [CrossRef]
  76. Tsuji, N.; Miyoshi, T.; Battsetseg, B.; Matsuo, T.; Xuan, X.; Fujisaki, K. A cysteine protease is critical for Babesia spp. transmission in Haemaphysalis ticks. PLoS Pathog. 2008, 4, e1000062. [Google Scholar] [CrossRef] [Green Version]
  77. Boldbaatar, D.; Battsetseg, B.; Matsuo, T.; Hatta, T.; Umemiya-Shirafuji, R.; Xuan, X.; Fujisaki, K. Tick vitellogenin receptor reveals critical role in oocyte development and transovarial transmission of Babesia parasite. Biochem. Cell Biol. 2008, 86, 331–344. [Google Scholar] [CrossRef]
  78. Bastos, R.G.; Ueti, M.W.; Knowles, D.P.; Scoles, G.A. The Rhipicephalus (Boophilus) microplus Bm86 gene plays a critical role in the fitness of ticks fed on cattle during acute Babesia bovis infection. Parasites Vectors 2010, 3, 111. [Google Scholar] [CrossRef] [Green Version]
  79. Antunes, S.; Galindo, R.C.; Almazan, C.; Rudenko, N.; Golovchenko, M.; Grubhoffer, L.; Shkap, V.; do Rosario, V.; de la Fuente, J.; Domingos, A. Functional genomics studies of Rhipicephalus (Boophilus) annulatus ticks in response to infection with the cattle protozoan parasite, Babesia bigemina. Int. J. Parasitol. 2012, 42, 187–195. [Google Scholar] [CrossRef]
  80. Heekin, A.M.; Guerrero, F.D.; Bendele, K.G.; Saldivar, L.; Scoles, G.A.; Dowd, S.E.; Gondro, C.; Nene, V.; Djikeng, A.; Brayton, K.A. The ovarian transcriptome of the cattle tick, Rhipicephalus (Boophilus) microplus, feeding upon a bovine host infected with Babesia bovis. Parasites Vectors 2013, 6, 276. [Google Scholar] [CrossRef] [Green Version]
  81. Merino, O.; Almazan, C.; Canales, M.; Villar, M.; Moreno-Cid, J.A.; Galindo, R.C.; de la Fuente, J. Targeting the tick protective antigen subolesin reduces vector infestations and pathogen infection by Anaplasma marginale and Babesia bigemina. Vaccine 2011, 29, 8575–8579. [Google Scholar] [CrossRef]
  82. Waldron, S.J.; Jorgensen, W.K. Transmission of Babesia spp. by the cattle tick (Boophilus microplus) to cattle treated with injectable or pour-on formulations of ivermectin and moxidectin. Aust. Vet. J. 1999, 77, 657–659. [Google Scholar] [CrossRef]
  83. Mangold, A.J.; Aguirre, D.H.; Cafrune, M.M.; de Echaide, S.T.; Guglielmone, A.A. Evaluation of the infectivity of a vaccinal and a pathogenic Babesia bovis strain from Argentina to Boophilus microplus. Vet. Parasitol. 1993, 51, 143–148. [Google Scholar] [CrossRef]
  84. Jongejan, F.; de Vos, C.; Fourie, J.J.; Beugnet, F. A novel combination of fipronil and permethrin (Frontline Tri-Act(R)/Frontect(R)) reduces risk of transmission of Babesia canis by Dermacentor reticulatus and of Ehrlichia canis by Rhipicephalus sanguineus ticks to dogs. Parasites Vectors 2015, 8, 602. [Google Scholar] [CrossRef] [Green Version]
  85. Fourie, L.J.; Stanneck, D.; Horak, I.G. The efficacy of collars impregnated with flumethrin and propoxur against experimental infestations of adult Rhipicephalus sanguineus on dogs. J. S. Afr. Vet. Assoc. 2003, 74, 123–126. [Google Scholar] [CrossRef] [Green Version]
  86. Estrada-Pena, A.; Reme, C. Efficacy of a collar impregnated with amitraz and pyriproxyfen for prevention of experimental tick infestations by Rhipicephalus sanguineus, Ixodes ricinus, and Ixodes scapularis in dogs. J. Am. Vet. Med. Assoc. 2005, 226, 221–224. [Google Scholar] [CrossRef]
  87. Gregson, J. Notes on some phenomenal feeding of ticks. Proc. Ent. Soc. Br. Columb. 1938, 34, 8. [Google Scholar]
  88. Inokuma, H.; Kemp, D.H. Establishment of Boophilus microplus infected with Babesia bigemina by using in vitro tube feeding technique. J. Vet. Med. Sci. 1998, 60, 509–512. [Google Scholar] [CrossRef] [Green Version]
  89. Antunes, S.; Merino, O.; Mosqueda, J.; Moreno-Cid, J.A.; Bell-Sakyi, L.; Fragkoudis, R.; Weisheit, S.; Perez de la Lastra, J.M.; Alberdi, P.; Domingos, A.; et al. Tick capillary feeding for the study of proteins involved in tick-pathogen interactions as potential antigens for the control of tick infestation and pathogen infection. Parasites Vectors 2014, 7, 42. [Google Scholar] [CrossRef] [Green Version]
  90. Antunes, S.; Merino, O.; Lerias, J.; Domingues, N.; Mosqueda, J.; de la Fuente, J.; Domingos, A. Artificial feeding of Rhipicephalus microplus female ticks with anti calreticulin serum do not influence tick and Babesia bigemina acquisition. Ticks Tick Borne Dis. 2015, 6, 47–55. [Google Scholar] [CrossRef]
  91. Pierce, A.; Pierce, M. A note on the cultivation of Boophilus microplus (Canestrini, 1887) (Ixodidae: Acarina) on the embryonated hen egg. Aust. Vet. J. 1956, 32, 144–146. [Google Scholar] [CrossRef]
  92. Becker, C.A.; Malandrin, L.; Depoix, D.; Larcher, T.; David, P.H.; Chauvin, A.; Bischoff, E.; Bonnet, S. Identification of three CCp genes in Babesia divergens: Novel markers for sexual stages parasites. Mol. Biochem. Parasitol. 2010, 174, 36–43. [Google Scholar] [CrossRef]
  93. Becker, C.A.; Malandrin, L.; Larcher, T.; Chauvin, A.; Bischoff, E.; Bonnet, S.I. Validation of BdCCp2 as a marker for Babesia divergens sexual stages in ticks. Exp. Parasitol. 2013, 133, 51–56. [Google Scholar] [CrossRef]
  94. Maeda, H.; Hatta, T.; Alim, M.A.; Tsubokawa, D.; Mikami, F.; Matsubayashi, M.; Miyoshi, T.; Umemiya-Shirafuji, R.; Kawazu, S.I.; Igarashi, I.; et al. Establishment of a novel tick-Babesia experimental infection model. Sci. Rep. 2016, 6, 37039. [Google Scholar] [CrossRef] [Green Version]
  95. Rechav, Y.; Zyzak, M.; Fielden, L.J.; Childs, J.E. Comparison of methods for introducing and producing artificial infection of ixodid ticks (Acari: Ixodidae) with Ehrlichia chaffeensis. J. Med. Entomol. 1999, 36, 414–419. [Google Scholar] [CrossRef]
  96. Antunes, S.; Couto, J.; Ferrolho, J.; Rodrigues, F.; Nobre, J.; Santos, A.S.; Santos-Silva, M.M.; de la Fuente, J.; Domingos, A. Rhipicephalus bursa Sialotranscriptomic Response to Blood Feeding and Babesia ovis Infection: Identification of Candidate Protective Antigens. Front. Cell Infect. Microbiol. 2018, 8, 116. [Google Scholar] [CrossRef]
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Figure 1. Life cycle of Babesia spp. sensu stricto. Vertebrate hosts are infected following the bite of an infected hard tick, through the invasion of the host erythrocytes by sporozoites excreted in the tick saliva. Inside the erythrocyte, sporozoites develop into trophozoites that undergo an asexual multiplication called merogony, ending in the formation of either merozoites that can infect other erythrocytes, or of gametocytes, which eventually develop into gametes. Infected erythrocytes are taken up by the tick during its blood meal, but only the gametocytes survive, and they then undergo further development, changing into gametes in the tick midgut. Then, sexual multiplication—gamogony—takes place with the fusion of two gametes to form a motile zygote that enters the midgut epithelial cells to develop into motile kinetes through meiotic division. Kinetes disseminate to tick tissues, including ovarian and salivary gland cells. The invasion of tick ovaries results in transovarial transmission while those of salivary glands leads to transmission to the vertebrate host through injection of sporozoites with the saliva. Babesia microti-like species, which belong to Babesia spp. s.l. species, only invade the salivary glands, not the ovary. The schematic representation was made using the software biorender.com.
Figure 1. Life cycle of Babesia spp. sensu stricto. Vertebrate hosts are infected following the bite of an infected hard tick, through the invasion of the host erythrocytes by sporozoites excreted in the tick saliva. Inside the erythrocyte, sporozoites develop into trophozoites that undergo an asexual multiplication called merogony, ending in the formation of either merozoites that can infect other erythrocytes, or of gametocytes, which eventually develop into gametes. Infected erythrocytes are taken up by the tick during its blood meal, but only the gametocytes survive, and they then undergo further development, changing into gametes in the tick midgut. Then, sexual multiplication—gamogony—takes place with the fusion of two gametes to form a motile zygote that enters the midgut epithelial cells to develop into motile kinetes through meiotic division. Kinetes disseminate to tick tissues, including ovarian and salivary gland cells. The invasion of tick ovaries results in transovarial transmission while those of salivary glands leads to transmission to the vertebrate host through injection of sporozoites with the saliva. Babesia microti-like species, which belong to Babesia spp. s.l. species, only invade the salivary glands, not the ovary. The schematic representation was made using the software biorender.com.
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Figure 2. Experimental feeding of Ixodes ricinus ticks on (A) Siberian chipmunks (Tamias sibiricus barberi) [65], (B) mouse [66], (C,D) sheep [67], (E,F) rabbit [68], and (G,H) a membrane artificial feeding system [35].
Figure 2. Experimental feeding of Ixodes ricinus ticks on (A) Siberian chipmunks (Tamias sibiricus barberi) [65], (B) mouse [66], (C,D) sheep [67], (E,F) rabbit [68], and (G,H) a membrane artificial feeding system [35].
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Table
Table 1. Major Babesia species infecting humans, dogs, cattle, sheep, goat, and equids; their suspected or confirmed main vector and vertebrate hosts in the field; and the realization of the whole transmission cycle under experimental conditions.
Table 1. Major Babesia species infecting humans, dogs, cattle, sheep, goat, and equids; their suspected or confirmed main vector and vertebrate hosts in the field; and the realization of the whole transmission cycle under experimental conditions.
Babesia spp.Suspected or Confirmed
Main Vectors		Main Vertebrate HostsRealization of the Complete Transmission Cycle Under Experimental Conditions
Babesia microtiIxodes persulcatus
Ixodes ovatus
Ixodes scapularis
Ixodes trianguliceps
Ixodes dammini
Ixodes ricinus
Rhipicephalus haemaphysaloides
Haemaphysalis longicornis		Human, rodentND
ND
[28]
[29]
[30]
[31,32]
[33]
[34]
Babesia divergensIxodes ricinusHuman, cattle[35,36,37,38,39]
Babesia venatorumIxodes ricinusHuman, roe deer[40]
Babesia duncaniDermacentor albipictusHuman, mule deerND
Babesia bovisRhipicephalus microplus
Rhipicephalus annulatus
Rhipicephalus geigyi		Cattle, buffalo[41]
ND
ND
Babesia bigeminaRhipicephalusannulatus
Rhipicephalus microplus
Rhipicephalus decoloratus
Rhipicephalus geigyi
Rhipicephalus evertsi		Cattle, buffalo[42]
[43,44]
[45,46,47]
ND
ND
Babesia majorHaemaphysalis punctataCattle[48,49]
Babesia ovataHaemaphysalis longicornisCattle[50]
Babesia orientalisRhipicephalus haemaphysaloidesWater buffaloND
Babesia caballiDermacentor nitens
Dermacentor sp.
Hyalomma sp.
Rhipicephalus evertsi		Horse, Donkey, Mule[51,52]
ND
ND
[53]
Babesia ovisRhipicephalus bursaSheep and Goat[54]
Babesia motasiRhipicephalus bursa
Haemaphysalis punctata		Sheep and GoatND
ND
Babesia canisDermacentor reticulatus
Haemaphysalis spp.
Hyalomma spp.		Dog[55,56,57,58,59]
ND
ND
Babesia gibsoniHaemaphysalis sp.
Rhipicephalus sanguineus		DogND
ND
Babesia vogeliRhipicephalus sanguineusDogND
Babesia rossiHaemaphysaliselliptica
Rhipicephalus sanguineus		Dog[60]
ND
ND: no identified data.
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Bonnet, S.I.; Nadal, C. Experimental Infection of Ticks: An Essential Tool for the Analysis of Babesia Species Biology and Transmission. Pathogens 2021, 10, 1403. https://doi.org/10.3390/pathogens10111403

AMA Style

Bonnet SI, Nadal C. Experimental Infection of Ticks: An Essential Tool for the Analysis of Babesia Species Biology and Transmission. Pathogens. 2021; 10(11):1403. https://doi.org/10.3390/pathogens10111403

Chicago/Turabian Style

Bonnet, Sarah I., and Clémence Nadal. 2021. "Experimental Infection of Ticks: An Essential Tool for the Analysis of Babesia Species Biology and Transmission" Pathogens 10, no. 11: 1403. https://doi.org/10.3390/pathogens10111403

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