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. 2023 Feb 17;122(3):691–704. doi: 10.1007/s00436-023-07792-9

Transovarial transmission of pathogenic protozoa and rickettsial organisms in ticks

Reghu Ravindran 1,✉, Prabodh Kumar Hembram 1, Gatchanda Shravan Kumar 1, Karapparambu Gopalan Ajith Kumar 1, Chundayil Kalarickal Deepa 1, Anju Varghese 1
PMCID: PMC9936132  PMID: 36797442

Abstract

Transovarial transmission (TOT) is an efficient vertical transmission of pathogens that is observed in many arthropod vectors. This method seems to be an evolutionarily unique development observed only in Babesia sensu stricto (clade VI) and Rickettsia spp., whereas transstadial transmission is the common/default way of transmission. Transovarial transmission does not necessarily contribute to the amplification of tick-borne pathogens but does contribute to the maintenance of disease in the environment. This review aims to provide an updated summary of previous reports on TOT of tick-borne pathogens.

Keywords: Babesia, Anaplasma, Rickettsia, Eggs, Larvae, PCR

Background

Ticks are obligate hematophagous ectoparasites of mammals, birds, reptiles, and amphibians found worldwide, and have great medical and veterinary importance (Perez-Sautu et al. 2021). Ticks and tick-borne diseases (TTBDs) can reach serious levels resulting in human mortality and significant economic losses in livestock (Jongejan and Uilenberg 2004; Schnittger et al. 2012; Florin-Christensen et al. 2014). To date, there have been ~ 970 species of ticks identified in the order Ixodida with ~ 750 hard tick species (Ixodidae), ~ 218 species of soft ticks (Argasidae), and single species under the family Nuttalliellidae (Nuttalliella namaqua) and family Deinocrotonidae (Sonenshine 1991; Dantas-Torres and Otranto 2022).

Tick-borne protozoan pathogens infecting domestic animals are various species under the genera Babesia, Theileria, Cytauxzoon, and Hepatozoon. Similarly, many species of Rickettsia are infecting domestic animals. Species under each genus have different modes of transmission. To understand the unique transmission methods in each species, knowledge on the phylogeny or evolutionary history is important. Molecular phylogeny using 18S rRNA genes of piroplasmids infecting mammals resulted in the formation of six clades, viz., I (B. microti group), II (Western clade), III (Cytauxzoon spp.), IV (T. equi), V (Theileria sensu stricto), and VI (Babesia s.s.) (Schnittger et al. 2012, 2022).

Tick-borne pathogens have the potential to be transmitted through horizontal, transstadial, transovarial, venereal, co-feeding, and localized transmission (Parola and Raoult 2001; Turell 2007; Chauvin et al. 2009). During horizontal transmission, parasites are spread from host to tick and vice versa. Transstadial transmission occurs when there is transmission of parasites throughout the development of tick life stages, from the engorgement through moulting into the next unfed stage of the same individual tick. Such transmission may continue through more than one tick stage within a generation (larva to nymph to adult) in the absence of an oral infection (of the nymph) from a vertebrate host, whereas, in the case of transovarial or vertical transmission, the parasite transmission occurs from the female tick to the larvae of the next generation via the eggs (Randolph et al. 1996). The group of true Babesia or Babesia s.s. (clade VI) is characterized by transstadial and transovarial transmission. In contrast, true Theileria or Theileria s.s. (clade V) and T. equi (clade IV or Equus clade) show transstadial transmission and a schizont parasite stage (Schnittger et al. 2022). However, Babesia sensu lato clade I (B. microti group), II (Western clade), and III (Cytauxzoon) exhibit only transstadial transmission.

During localized transmission, an infected tick transmits the parasite to uninfected ticks feeding at the same skin site (transmission may continue beyond the duration of the blood meal of the infected tick). Whereas during co-feeding, the transmission occurs from infected to uninfected ticks while feeding simultaneously on the same host, but not necessarily at the same skin site, in the absence of a systemic infection in the host (Randolph et al. 1996). Lastly, venereal or sexual transmission occurs during the mating of ticks. For a tick to be considered a competent vector, horizontal transmission and at least one of these other transmission routes must be present (Kahl et al. 2002; Pfaffle et al. 2013; Schnittger et al. 2022).

Among the different transmission mechanisms, transstadial transmission is the key survival strategy for many piroplasms. The lifelong carrier status of the vertebrate host ensures the infection of different lifecycle stages of ticks. In contrast, in the case of transovarial transmission, the parasite will be passed on to the next generation once a tick is infected, without the need for prior feeding on an infected host. Here, the tick functions as a carrier of the pathogen. Hence, a prolonged carrier status of the vertebrate host is not necessary for the transovarial transmission. The transovarial transmission facilitates species diversification by host switching to other vertebrate host species (Schnittger et al. 2022). It is not necessarily contributing to the amplification of tick-borne disease, rather contributes to the maintenance of disease in the environment. Hence, more detailed studies on transovarial transmission are essential. Here, in this review, the updated information on the reports of transovarial transmission of pathogens in ticks is presented.

Babesia spp.

Babesiosis was the first arthropod-borne mammalian disease discovered and has been shown to spread from one generation of a hard tick to the next via a transovarial transmission (Smith and Kilbourne 1893; Demessie and Derso 2015). Ticks become infected with Babesia parasites when they ingest blood cells containing gametocytes, which develop into ray bodies or Strahlenkörper (male and female gametes) in their midgut (Uilenberg 2006), which fuse to form a motile zygote (ookinete), that invades the tick gut cells and undergoes meiotic division, resulting in the production of kinetes. Kinetes disseminate via hemolymph to peripheral tick tissues, including ovarian cells leading to the infection of eggs (Jalovecka et al. 2019).

Babesia sensu stricto (clade VI) shows a unique evolutionary pattern because of their ability of transovarial transmission which facilitates the diversification of Babesia s.s. species to all groups of vertebrates around the world. All ruminant infecting Babesia species (B. bigemina, B. bovis, B. divergens, B. ovata, B. major, B. occultans, B. orientalis, Babesia sp. Mymensingh nk, Babesia sp. Tengchong, B. ovis, B. crassa, B. motasi, B. motasi-like, Babesia sp. Xinjiang, etc.) belong to the Babesia sensu stricto group (clade VI) (Schnittger et al. 2012, 2022), which are characterized by the lack of a schizont stage, asexual reproduction exclusively within red blood cells in vertebrate hosts, and the occurrence of transovarial transmission in the tick vector (Uilenberg 2006; Schnittger et al. 2012; Jalovecka et al. 2019). Ticks belonging to the genera Rhipicephalus and Ixodes are generally implicated in the transmission of bovine babesiosis.

Babesia spp. that cause infections in dogs are divided into the Babesia s.s. clade (clade VI) (B. vogeli, B. canis, B. rossi, B. gibsoni, Rangelia vitalii, and Babesia sp. Coco, Babesia sp. Akita610) and two clearly identifiable Babesia s.l. clades, namely, the Western clade (clade II) (B. negevi, B. conradae) and B. vulpes group (clade Ib) (B. vulpes) (Jalovecka et al. 2019). In horses, only B. caballi is recognised as a true Babesia species (Babesia s.s) (clade VI), while B. equi is reclassified as T. equi (clade IV). Babesia caballi is transmitted transovarially. Among the babesias (B. trautmanni, B. perroncitoi, Babesia sp. Suis) that cause infections in swine, Babesia sp. Suis was recently characterized as Babesia s.s. (clade VI), based on the molecular phylogeny using 18S rRNA genes (Avenant et al. 2021).

Table 1 lists the reports of transovarial transmission of different Babesia spp. In ticks, the transovarial infection rate (the percentage of female ticks that pass microorganisms to their progeny) (Burgdorfer and Varma 1967) with different Babesia spp. ranged from 0.5 to 100% while the filial infection rate (the percentage of infected progeny derived from an infected female tick) (Burgdorfer and Varma 1967) for the same were not available in the published reports.

Table 1.

Transovarial transmission of Babesia spp. in ticks

Place Tick species Study type, test performed Transovarial infection rate Filial infection rate Reference
B. bigemina (bovine)
Australia Rh. microplus Experimental study NA NA Riek (1964)
Brazil Rh. microplus nPCR NA NA Oliveira-Sequeira et al. (2005)
Brazil Rh. microplus Microscopic examination NA NA Oliveira et al. (2005)
Brazil Rh. microplus qPCR 20 to 40% NA Giglioti et al. (2018)
Cuba Rh. microplus qPCR 68% NA Obregon et al. (2020)
India Rh. microplus DNA hybridization with a nonradioactive probe NA NA Ravindran et al. (2006)
India Rh. microplus nPCR 7.41% NA Bhat et al. (2017)
India Rh. annulatus PCR 38% NA Hembram et al. (2022)
Iran Rh. annulatus PCR NA NA Rajabi et al. (2017)
Israel Rh. annulatus Nested PCR NA NA Molad et al. (2015)
Kenya Rh. decoloratus Experimental study NA NA Morzaria et al. (1977)
South Africa Rh. decoloratus Experimental study NA NA Gray and Potgieter 1982
Turkey Rh. annulatus Reverse line blot (RLB) NA NA Ica et al. (2007)
Uruguay Rh. microplus PCR NA NA Gayo et al. (2003)
USA Rh. annulatus Experimental study NA NA Smith and Kilbourne (1893)
B. bovis (bovine)
Australia Rh. microplus Experimental study NA NA Mahoney and Mirre (1979)
Brazil Rh. microplus nPCR NA NA Oliveira-Sequeira et al. (2005)
Brazil Rh. microplus Microscopic examination NA NA Oliveira et al. (2005)
Brazil Rh. microplus qPCR 0.5 to 14.5% NA Giglioti et al. (2018)
Cuba Rh. microplus qPCR 100% NA Obregon et al. (2020)
Uruguay Rh. microplus PCR NA NA Gayo et al. (2003)
USA Rh. microplus Experimental study NA NA Smith et al. (1978)
USA Rh. microplus PCR 12% to 48% NA Howell et al. (2007)
B. ovata (bovine)
Japan Ha. longicornis IFAT NA NA Maeda et al. (2016)
Japan Ha. longicornis nPCR NA NA Shirafuji et al. (2017)
B. occultans (bovine)
South Africa Hy. rufipes Experimental study NA NA Gray and de Vos (1981)
Turkey Hy. marginatum PCR 22.22% NA Aktas et al. (2014)
Turkey Rh. turanicus PCR 50% NA Aktas et al. (2014)
Turkey Hy. marginatum PCR NA NA Orkun (2019)
Turkey Hy. excavatum PCR NA NA Orkun (2019)
B. divergens (bovine)
England I. ricinus Experimental study NA NA Donnelly and Peirce (1975)
France I. ricinus PCR NA NA Bonnet et al. (2007)
B. canis (canine)
England Ha. elliptica Experimental study NA NA Shortt (1973)
England Rh. sanguineus Experimental study NA NA Shortt (1973)
Poland D. reticulatus PCR 100% NA Mierzejewska et al. (2018)
West-central Poland I. ricinus PCR NA NA Liberska et al. (2021)
B. rossi (canine)
Nigeria, West Africa Ha. leachi PCR NA NA Kamani (2021)
B. vogeli (canine)
Taiwan Rh. sanguineus PCR NA NA Jongejan et al. (2018)
B. gibsoni (canine)
Japan Ha. longicornis PCR NA NA Hatta et al. (2012)
Taiwan Ha. hystricis PCR NA NA Jongejan et al. (2018)
B. ovis (ovine)
Iran Rh. bursa (two-host tick) PCR NA NA Esmaeilnejad et al. (2014)
Israel Rh. bursa (two-host tick) Experimental study NA NA Yeruham et al. (2001)
Israel Rh. bursa (two-host tick) PCR NA NA Erster et al. (2016)
Turkey Rh. bursa (two-host tick) PCR NA NA Orkun (2019)
B. motasi (ovine)
Netherlands Ha. punctata Experimental study NA NA Uilenberg et al. (1980)
Great Britain Ha. punctata Experimental study NA NA Alani and Herbert (1988)
B. caballi (equine)
Americas Dermacentor nitens PCR NA NA Schwint et al. (2008)
Brazil Rh. microplus PCR NA NA Battsetseg et al. (2002)
US state, Florida Hy. truncatum Experimental study NA NA de Waal (1990)
B. trautmanni (porcine)
South Africa Rh. simus Experimental study NA NA de Waal et al. (1992)
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NA not available

Theileria spp.

Theileriosis, (Phylum Apicomplexa; order Piroplasmida; family Theileriidae; genus Theileria) remains a burden for millions of livestock in tropical countries, especially crossbreds and exotic cattle annually (Roy et al. 2021). The dominant Theileria spp. linked to economic loss and mortality worldwide are T. annulata and T. parva (Roy et al. 2021). Mild bovine theileriosis is caused by T. orientalis, T. mutans, T. velifera, and T. taurotragi. Various genotypes of T. orientalis are type 1 (Chitose), type 2 (Ikeda), type 3 (Buffeli), types 4–8, and types N1–N3 (Hammer et al. 2015). Bovine Theileria species, which include T. annulata, T. parva, T. orientalis (syn. T. buffeli/T. sergenti/T. sinensis), T. mutans, T. velifera, and T. taurotragi, belong to a monophyletic group corresponding to clade V (Theileria sensu stricto group) (Schnittger et al. 2012, 2022). Members of this clade exhibit a schizont stage in the lymphoid cells and piroplasms in the red blood cells of the vertebrate host, as well as exclusive transstadial transmission but not transovarial transmission in the tick (Kiara et al. 2018). Theileria annulata (tropical bovine theileriosis) infection is most common in southern Europe, North Africa, the Middle East, and Asia transmitted transstadially by several species of Hyalomma ticks, namely, Hyalomma anatolicum, Hy. dromedarii, Hy. detritum, Hy. scupense, and Hy. lusitanicum (Ali et al. 2013; Jabbar et al. 2015; Gharbi et al. 2020). Hyalomma ticks transmit T. annulata sporozoites into the host and causes a lymphoproliferative disease similar to cancer (Ghosh et al. 2007; Tretina et al. 2015). In addition to a tick bite, transplacental transmission has been detected for T. annulata (Sudan et al. 2015), by PCR analysis. There is no report on the transovarial transmission of T. annulata in ticks (Mehlhorn and Schein 1984; Norval et al. 1992).

Theileria parva, transmitted transstadially, most commonly by Rh. appendiculatus, is present throughout a large part of eastern and southern Africa (Morrison et al. 2020). These parasites also infect the Asian and African species of buffalo (Bubalus bubalis and Syncerus caffer, respectively) (Morrison et al. 2020). Based on the available literature, there are no reports on the transovarial transmission of T. parva in ticks.

Oriental theileriosis was reported in Asia, New Zealand, Australia, and the USA (Oakes et al. 2019). Haemaphysalis longicornis is a known vector tick for T. orientalis in different countries (Fujisaki et al.1994; Hammer et al. 2015, Jabbar et al. 2015). Other potential vectors include Ha. punctata in France (Uilenberg 2000), Rh. microplus in Vietnam and Thailand (Khukhuu et al. 2011; Poolkhetkit et al. 2015), Rh. decoloratus and Rh. evertsi in Ethiopia (Kumsa et al. 2013), and Rh. annulatus in India (Nimisha et al. 2019). Available literature reveals only one report on the detection of the parasite DNA in the eggs of Rh. microplus (Kakati et al. 2015) engorged on a parasite-positive animal. Theileria orientalis can be spread in various ways other than by tick vectors. It has been proven that the infected heifers can transmit the parasite to their foetus or calf (Baek et al. 2003; Lawrence et al. 2016; Swilks et al. 2017; Mekata et al. 2018). Transmission via the transcolostral route is also plausible, but requires more research to confirm this (Emery 2016). In addition, theilerial DNA was detected in mosquitos, lice (Linognathus vituli), and other hematophagous insects (Emery 2016; Hammer et al. 2016). Theileria orientalis can also be transmitted by transfer of piroplasms when contaminated needles (vaccinations), castration knives, and ear notching equipments are reused. In addition, the injuries sustained during yarding and transport of cattle can also assist in the transmission (Hammer et al. 2016).

Among six Theileria species infecting goats, T. lestoquardi, T. luwenshuni, and T. uilenbergi are extremely pathogenic, causing high mortality, and the remaining three, T. separata, T. ovis and T. recondite, are less pathogenic in small ruminants (Islam et al. 2021). It is believed that these infections are transmitted transstadially through multihost ticks and there were no previous reports on transovarial transmission for these parasites.

Babesia equi in horses is reclassified as T. equi owing to its extraerythrocytic schizogony, erythrocytic invasion, and transstadial transmission in ticks (Mehlhorn and Schein 1998; Ueti and Knowles 2018). However, there are reports on the occurrence of transovarial transmission of T. equi in Rh. microplus (Battsetseg et al. 2002) and Ha. longicornis tick (Ikadai et al. 2007). Phylogenetic studies proved that T. equi does not belong to Theileria s.s., but rather represents a unique separate monophyletic clade (clade IV or Equus group) (Schnittger et al. 2012; Jalovecka et al. 2019; Bhoora et al. 2020).

Cytauxzoon (clade III) is characterized by the presence of a schizont stage that infects host cells of the mononuclear reticulohistiocytic system (Schnittger et al. 2022). Dermacentor variabilis was initially accepted to be the natural tick vector of C. felis (Blouin et al. 1984) while transstadial transmission was experimentally proved recently in Amblyomma americanum (Reichard et al. 2009).

Rickettsia spp.

The organisms assigned to the order Rickettsialses were reclassified based on 16S rRNA genes, groESL, and surface protein genes into two families viz., Anaplasmataceae and Rickettsiaceae (Dumler et al. 2001). All the members of the family Rickettsiaceae are slow-growing gram-negative bacteria that are pleomorphic, obligatory intracellular, have a life cycle that involves both an arthropod vector and a vertebrate host (Portillo et al. 2017; Blanda et al. 2020), and grow freely in the cytoplasm of eukaryotic cells. These bacteria can be transmitted to animals and humans by blood-sucking arthropods, causing specific zoonotic diseases termed rickettsioses (Merhej et al. 2014; de Mera et al. 2018). Rickettsia and Orientia are the two genera causing rickettsioses (Jiang et al. 2021) in animals and man.

Based on the disease presentation, antigenicity, and vectors, rickettsial diseases (and their causative agents) have been traditionally separated into three major groups viz., spotted fever group (SFG), typhus group, and scrub typhus group (Luce-Fedrow et al. 2015; Parola et al. 2013; Abdad et al. 2018; Richards and Jiang 2020). More than 30 species are included in the SFG, with more species being added in each year (https://www.bacterio.net).

Many rickettsial endosymbionts of invertebrates are thought to be vertically transmitted, implying that arthropod vectors serve as reservoirs or amplifiers in nature (Parola et al. 2013). The Rickettsia of spotted fever category encompasses a number of human infections, the majority of which are spread by ticks. The Rickettsia spp. transmitted transovarially in the ticks are shown in Table 2. In ticks, the transovarial infection rate with different Rickettsia spp. ranged from 8 to 100%, while its filial infection rate for the same ranged from 22.7 to 100%.

Table 2.

Transovarial transmission of Rickettsia spp. in ticks

Place Tick species Study type, test performed Transovarial infection rate Filial infection rate Reference
R. rickettsii
Brazil A. aureolatum PCR 100% 100% Labruna et al. (2011)
Brazil Rh. sanguineus PCR NA  < 50% Piranda et al. (2011)
Brazil Rh. sanguineus PCR NA 100% Pacheco et al. (2011)
Brazil A. cajennense qPCR  < 50%  < 50% Soares et al. (2012)
Brazil A. aureolatum qPCR 25% NA Binder et al. (2021)
USA D. andersoni Experimental study 100% 100% Burgdorfer (1963)
R. conorii conorii
Algeria Rh. sanguineus PCR 100% Up to 99% Socolovschi et al. (2009a, b)
Algeria Rh. sanguineus PCR 100% Up to 99% Socolovschi et al. (2012)
Thailand Rh. sanguineus PCR NA NA Matsumoto et al. (2005a, b)
R. raoultii
India Rh. annulatus PCR 8% NA Hembram et al. (2022)
India Ha. bispinosa PCR 15% NA Hembram et al. (2022)
Netherlands D. reticulatus PCR NA NA Alberdi et al. (2012)
Northern Mongolia D. nuttalli nPCR NA NA Moore et al. (2018)
Turkey D. marginatus PCR NA NA Orkun (2019)
R. slovaca
Turkey D. marginatus PCR NA NA Orkun (2019)
USA D. variabilis PCR  ≥ 99%  ≥ 99% Zemtsova et al. (2016)
R. massiliae
France Rh. turanicus PCR 100% 98.5% Matsumoto et al. (2005a, b)
R. africae
Ivory Coast, Africa A. variegatum PCR 100% 93.4% Socolovschi et al. (2009a, b)
R. bellii
Brazil I. loricatus PCR NA NA Horta et al. (2006)
R. aeschlimannii
Turkey Hy. marginatum PCR 25% NA Orkun (2019)
R. amblyommii
Brazil A. auricularium PCR 100% 100% Saraiva et al. (2013)
R. montana
USA D. variabilis PCR NA NA Macaluso et al. (2001)
R. rhipicephali
USA D. variabilis PCR NA NA Macaluso et al. (2001)
Rickettsia spp.
Northern Germany I. ricinus qPCR NA 22.7% Hauck et al. (2020)
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NA not available

Anaplasma spp.

The organisms placed under family Anaplasmataceae are obligate intracellular parasites found exclusively within the membrane-bound vacuoles in the host cell cytoplasm. The family Anaplasmataceae include four genera viz., Anaplasma, Ehrlichia, Neorickettsia, and Wolbachia. The genus Anaplasma include A. marginale, A. marginale subsp. centrale, A. phagocytophilum, A. bovis, and A. platys (Kocan et al. 2010).

A. marginale

Anaplasmosis causes considerable economic loss to beef and dairy industries globally, including those in the America, Europe, Australia, Asia, and Africa (Aubry and Geale 2011; Atif 2015; Kocan et al. 2015). Anaplasma spp. can be transmitted biologically by ticks, mechanically by blood-sucking arthropods (blood-contaminated mouthparts of biting flies) or blood-contaminated fomites, i.e., castration and dehorning equipment, needles, and ear tag applicators (Kocan et al. 2015; Battilani et al. 2017).

Many species of ticks are reported to serve as the vectors of A. marginale viz., Argas persicus, Dermacentor andersoni, D. albipictus, D. calcaratus, D. variabilis, D. occidentalis, D. hunteri, Hy. excavatum, Hy. rufipes, I. ricinus, I. scapularis, Ornithodoros lahorensis, Rh. microplus, Rh. annulatus, Rh. decoloratus, Rh. evertsi, and Rh. simus, but the most common vectors throughout tropical and subtropical areas of the world are Dermacentor spp. (D. andersoni, D. variabilis, and D. albipictus) and Rhipicephalus (Boophilus) spp. (Rh. microplus and Rh. annulatus) (Rar and Golovljova 2011; Kocan et al. 2015). Tick transmission can occur from stage to stage (interstadial or transstadial) or within a stage (intrastadial) (Stich et al. 1989). Interstadial transmission of A. marginale has been demonstrated by the 3-host ticks, D. andersoni, and D. variabilis in the USA (Kocan 1986; Kocan et al. 1981, 1985; Stiller et al. 1989), Rh. sanguineus in Israel (Shkap et al. 2009) and by Rh. simus in South Africa (Potgieter and Van Rensburg 1980, 1982; Potgieter et al. 1983). Intrastadial transmission of A. marginale is caused by male ticks which serve as the reservoir hosts of the organisms, persistently infecting the cattle (Ge et al. 1996; Kocan et al. 1992, 2000; Palmer et al. 2001). The co-feeding of ticks does not appear to influence the dynamics of A. marginale transmission (Kocan and de la Fuente 2003). Transplacental transmission of A. marginale occurs in cattle, resulting in healthy but persistently infected calves (Grau et al. 2013).

There are very few reports on the occurrence of transovarial transmission of A. marginale. Shimada et al. (2004) detected A. marginale major surface protein 5 (msp5) gene in larvae of Rh. microplus by PCR amplification. Amaro Estrada et al. (2020) confirmed the transovarial transmission of A. marginale by detecting it in the unfed larvae hatched from the fully engorged Rh. microplus by the PCR targeting both msp5 and major surface protein1α (msp1α) genes. Kumar et al. (2019) detected this organism in the Rh. microplus ticks and their egg masses. Hembram et al. (2022) detected this organism in the Rh. annulatus ticks, their egg masses, and unfed larvae.

A. bovis

Anaplasma bovis is a bacterium infecting the circulating monocytes (Sreekumar et al. 1996; Liu et al. 2012) and tissue macrophages of domestic and wild ruminants (Worthington and Bigalke 2001). The infection in cattle is normally asymptomatic, although it can induce a number of clinical symptoms, including decreased body weight, fever, anemia, depression, lymphadenopathy, and in rare cases, abortion, as well as death. Anaplasma bovis DNA was detected in the nymphs and larvae of Ha. megaspinosa in Japan (Yoshimoto et al. 2010), Rh. turanicus in Israel (Harrus et al. 2011), engorged female Rh. annulatus in India (Nimisha et al. 2019), and an undescribed tick species in South Africa (Harrison et al. 2011). There are no reports on the transovarial transmission of A. bovis in ticks.

A. phagocytophilum

Anaplasma phagocytophilum, an obligate intracellular gram-negative bacterium is the etiological agent of tick-borne fever (TBF) in ruminants (Atif 2015) and of equine, canine, and human granulocytic anaplasmosis (EGA, CGA, and HGA, respectively) (Dumler et al. 2001; Woldehiwet 2010). Anaplasma phagocytophilum (Rickettsiales: Anaplasmataceae) has become an important tick-borne pathogen in the USA, Europe, and Asia, with increasing numbers of infected people and animals every year (Bakken and Dumler 2015; Tang et al. 2015). Anaplasma phagocytophilum multiplies within a parasitophorous vacuole to form a morula in the cytoplasm of tick and vertebrate host cells (Dumler et al. 2001; Tang et al. 2015; Munderloh et al. 1999). Fatal cases have been reported so far in sheep, cattle, horses, reindeer, roe deer, moose, dogs, and humans (Jenkins et al. 2001; Stuen 2003; Franzen et al. 2007).

The most typical way to spread the A. phagocytophilum is through the bite of an infected tick (Jaarsma et al. 2019). Transstadial transmission is essential in maintaining A. phagocytophilum within its endemic cycles (Medlock et al. 2013; Jahfari et al. 2014; Krucken et al. 2013). Ixodes ricinus (Strle 2004; Parola et al. 2005), I. scapularis, and I. persulcatus (Alekseev et al. 1998; Woldehiwet 2010) were identified as vectors. Anaplasma phagocytophilum has been found in questing ticks belonging to other members of the genus Ixodes like I. trianguliceps (Ogden et al. 1998), I. ventalloi (Santos et al. 2004), I. hexagonus (Pfaffle et al. 2011), and I. nipponensis (Lee et al. 2020). Anaplasma phagocytophilum DNA has also been found in D. reticulatus (Karbowiak et al. 2014), Haemaphysalis punctata, Ha. concinna, and Rh. bursa (Barandika et al. 2007).

Although the transovarial transmission has not been shown in Ixodes species, it has been demonstrated in moose tick D. albipictus, a parasite with a single host life cycle (Baldridge et al. 2009). The presence of A. phagocytophilum was confirmed by PCR in unfed larvae of D. albipictus (Baldridge et al. 2009) and I. ricinus (Hauck et al. 2020). Hembram et al. (2022) detected this organism in the fully repleted Rh. annulatus and Ha. bispinosa ticks as well as their progenies.

Concluding remarks

Numerous factors have contributed to an increase in the incidence and diversity of tick-borne diseases in both humans and animals in recent years. Global climate change favored the spread of vector populations restricted previously to narrow geo-climatic conditions to new and wider areas, thereby spreading the infections carried by them. Urbanization and habitat encroachment caused increased contact of humans/animals with wildlife and new vectors. Human activities including deforestation, reforestation, and plantation lead to a situation with increased interaction of the host with the widely dispersed blood-feeding ectoparasites, previously restricted only to the forest environments. This resulted in changes in the vector ecology. In addition, the availability of better diagnostic tools and increased awareness among the scientific community, veterinarians, physicians, and public health authorities contributed to significant improvement in the knowledge of TTBDs. Presently, TTBDs are considered to be a major problem for both human and animal populations. There is still a great dearth of knowledge regarding the vector potential of many tick species found throughout the world. In order to elucidate the disease biology of tick-borne diseases, it is essential to understand their transmission mechanisms. Among the different transmission mechanisms, transstadial transmission is the key survival strategy for many piroplasms. The lifelong carrier status of the vertebrate host ensures infection of such organisms infecting different lifecycle stages of ticks. In contrast, in the case of transovarial transmission, the parasite will be passed on into the next generation once a tick is infected, without the need for prior feeding on an infected host. Here, the tick functions as a carrier of the pathogen. Hence, a prolonged carrier status of the vertebrate host is not necessary for transmission. The transovarial transmission facilitates species diversification by host switching to other vertebrate host species. Thus, transovarial transmission plays a role in establishing the endemicity of the infection. Hence, it can affect the control efforts against the pathogen in a particular region. The transovarial transmission results in the formation of infected larvae (more when the transovarial and filial infection rates are high) with greater potential for spreading disease compared to nymphal and adult stages since such larvae are minute and difficult to be detected with the naked eye.

Acknowledgements

The second author wishes to acknowledge the Indian Council of Agricultural Research (ICAR) for providing the National Talent Scholarship (NTS) and Kerala Veterinary and Animal Sciences University (KVASU) for the facilities.

Abbreviations

TOT

Transovarial transmission

SFG

Spotted fever group

TTBDs

Ticks and tick-borne diseases

PCR

Polymerase chain reaction

nPCR

Nested polymerase chain reaction

RLB

Reverse line blot

qPCR

Quantitative polymerase chain reaction

IFAT

Indirect immunofluorescent antibody test

msp5

Major surface protein 5

msp1α

Major surface protein1α

TBF

Tick-borne fever

B

Babesia

A

Anaplasma

Th

Theileria

Rh

Rhipicephalus

Ha

Haemaphysalis

Hy

Hyalomma

I

Ixodes

D

Dermacentor

R

Rickettsia

NA

Not available

s.s.

sensu stricto

s.l.

sensu lato

Author contribution

R.R. and P.K.H. wrote the manuscript text and prepared tables. G.S.K. and C.K.D. edited the tables. K.G.A. and A.V. edited the manuscript. All authors read and approved the final manuscript.

Funding

The study was financially supported by the RKVY-RAFTAAR 2019–20 project (KE/RKVY-ANHB/2019/1422), Kerala state plan project (2021–22) (RSP/21–22/VI-7), Indian Council of Agricultural Research (ICAR)-sponsored research projects (NAIP/C2066, NFBSFARA/BSA-4004/2013–14, NASF/ABA-6015/2016–17), Kerala State Council for Science, Technology, and Environment-sponsored research projects (022/YIPB/KBC/2013/CSTE, 010–14/SARD/13/CSTE), and Kerala Veterinary and Animal Sciences University (KVASU/2019/078/MVP/VPR). The funders had no role in the study design, data collection, analysis, decision to publish, or preparation/content of the manuscript.

Data availability

Not applicable.

Code availability

Not applicable.

All the required ethical standards were complied with.

Declarations

Competing interests

The authors declare no competing interests.

Consent of publication

All authors consent to the publication of this manuscript.

Conflict of interest

The authors declare no competing interests.

Footnotes

Publisher's note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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