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A novel Babesia sp. associated with clinical signs of babesiosis in domestic cats in South Africa

Parasites & Vectors201912:138

https://doi.org/10.1186/s13071-019-3395-x

  • Received: 16 November 2018
  • Accepted: 11 March 2019
  • Published:

Abstract

Background

Feline babesiosis, sporadically reported from various countries, is of major clinical significance in South Africa, particularly in certain coastal areas. Babesia felis, B. leo, B. lengau and B. microti have been reported from domestic cats in South Africa. Blood specimens from domestic cats (n = 18) showing clinical signs consistent with feline babesiosis and confirmed to harbour Babesia spp. piroplasms by microscopy of blood smears and/or reverse line blot (RLB) hybridization were further investigated. Twelve of the RLB-positive specimens had reacted with the Babesia genus-specific probe only, which would suggest the presence of a novel or previously undescribed Babesia species. The aim of this study was to characterise these organisms using 18S rRNA gene sequence analysis.

Results

The parasite 18S rRNA gene was cloned and sequenced from genomic DNA from blood samples. Assembled sequences were used to construct similarity matrices and phylogenetic relationships with known Babesia spp. Fifty-five 18S rRNA gene sequences were obtained. Sequences from 6 cats were most closely related to published B. felis sequences (99–100% sequence identity), while sequences from 5 cats were most closely related to B. leo sequences (99–100% sequence identity). One of these was the first record of B. leo in Mozambique. One sequence had 100% sequence identity with the published B. microti Otsu strain. The most significant finding was that sequences from 7 cats constituted a novel Babesia group with 96% identity to Babesia spp. previously recorded from a maned wolf (Chrysocyon brachyurus), a raccoon (Procyon lotor) from the USA and feral raccoons from Japan, as well as from ticks collected from dogs in Japan.

Conclusions

Babesia leo was unambiguously linked to babesiosis in cats. Our results indicate the presence of a novel potentially pathogenic Babesia sp. in felids in South Africa, which is not closely related to B. felis, B. lengau and B. leo, the species known to be pathogenic to cats in South Africa. Due to the lack of an appropriate type-specimen, we refrain from describing a new species but refer to the novel organism as Babesia sp. cat Western Cape.

Keywords

  • 18S rRNA gene
  • Babesia leo
  • Babesia sp. cat Western Cape
  • Domestic cat
  • Felidae
  • Phylogeny
  • South Africa

Background

Domestication of cats occurred in the Near East, probably by natural selection, the ancestor being the local feline subspecies, Felis silvestris lybica [1]. From here, domestic cats (Felis silvestris catus) have spread world-wide with a current total population of kept or feral cats estimated at nearly one billion [2]. With the exception of Australia, all inhabited continents also harbour indigenous felid species from which pathogens could conceivably be transferred to domestic cats. Feline babesiosis may be a case in point. Although cases of cats showing clinical signs of babesiosis have been reported sporadically from various countries, feline babesiosis seems to be an important disease of domestic cats only in South Africa, especially along the eastern and southern seaboard and with a few foci on the eastern escarpment [3, 4].

Babesia felis was described from a c.3-month-old wild-caught Sudanese wild cat (Felis ocreata, presumably a synonym of a F. silvestris subspecies) that was observed for 12 months but showed no overt clinical signs of disease [5]. Parasitaemia, initially 0.5%, soon peaked at 8% (possibly due to stress while the host was adapting to captivity), but gradually decreased over a 3-month period and subsequently fluctuated around 0.4%. Blood from this cat was inoculated into 22 domestic cats. None of these cats showed any overt clinical signs of disease, but all developed a parasitaemia not exceeding 1% initially and then decreasing to a fluctuating low level which persisted indefinitely [5]. Following the classification suggested by Wenyon [6], Davis [5] assigned the novel parasite to the genus Babesia; he did not designate and deposit a type-specimen, however, which led to subsequent confusion.

During the 1930s domestic cats exhibiting clinical signs similar to those of canine babesiosis, i.e. anaemia, icterus and lethargy, were occasionally presented to veterinarians in South Africa, especially in the Western Cape Province [7, 8]. Felis caffra, presumably the local subspecies of F. silvestris, was suspected as being a reservoir host [8]. In the index case report of feline babesiosis [7], the piroplasms seen on blood smears met the description of B. felis piroplasms by Davis [5]. Due to its pathogenicity in domestic cats, in contrast to B. felis (sensu stricto), Jackson et al. [7] proposed the name Nuttalia felis var. domestica for the South African organism. Choosing Nuttalia rather than Babesia as genus name, they followed Carpano et al. [9] in preferring the classification by Du Toit [10] rather than that of Wenyon [6].

Regrettably, Jackson’s [7] conclusion that the South African organism represented a distinct taxon to B. felis (s.s.), being at least a local variety of the latter, was overlooked in subsequent reports on clinical manifestation and treatment of feline babesiosis: the causative organism was merely referred to as B. felis [1113]. This was also the name used when details of molecular characterisation of the Babesia sp. causing disease in cats were deposited in the GenBank database [14]. The matter will only be resolved if Davis’s [5] original specimens are traced, which seems unlikely. Molecular characterisation has since revealed the presence of B. felis (sensu lato) in cheetahs (Acinonyx jubatus), lions (Panthera leo) and servals (Leptailurus serval) in South Africa, Namibia and Zambia [15, 16].

Domestic cats can also be infected with other Babesia spp. A large, unidentified Babesia was incriminated in causing severe clinical signs in a domestic cat in Harare, Zimbabwe [17]. When examining blood smears of sick cats in South Africa, veterinarians occasionally report finding large organisms (Fig. 1), resembling Babesia rossi of dogs rather than the small B. felis (s.l.) (Figs. 2, 3); attempts at identifying these organisms were unsuccessful (pers. obs.). Babesia canis subsp. presentii was described from two cats in Israel, one a subclinical carrier and the other suffering from co-infection of various other pathogens [18].
Fig. 1
Fig. 1

Blood smear from a cat with clinical signs of babesiosis, showing large, extracellular piroplasms (Courtesy: Dr James Hill, Vetdiagnostix, Pietermaritzburg)

Fig. 2
Fig. 2

Blood smear from a cat with clinical signs of babesiosis, showing small, intra-erythrocytic Babesia felis (sensu lato) piroplasms (Courtesy: Dr Sandy Weltan, Vetdiagnostix, Cape Town)

Fig. 3
Fig. 3

Blood smear from a cat with high parasitaemia of Babesia felis (sensu lato) piroplasms

Babesia pantherae, a large piroplasm isolated from leopards (Panthera pardus) in Kenya and B. herpailuri isolated from a jaguarundi (Herpailurus yaguarondi) originating from Venezuela could be established in domestic cats [1921]. In both cases overt clinical signs developed only in asplenic cats; spleen-intact cats developed a long-lasting parasitaemia but remained asymptomatic [19]. Unfortunately, this was before the advent of molecular characterisation of piroplasms.

A previous South African survey of cats with clinical signs consistent with babesiosis suggested the presence of further potentially pathogenic piroplasms [15]. Subsequent molecular characterisation revealed that the pathogen involved in two fatal cases of feline babesiosis, one being the first record of cerebral babesiosis in a domestic cat, showed a high similarity with B. lengau, previously described from asymptomatic cheetahs [22, 23].

The aim of the present study was to characterise piroplasms from domestic cats in South Africa (Western Cape and KwaZulu-Natal) and Mozambique (Maputo) exhibiting clinical signs of babesiosis, using 18S rRNA gene sequence data and phylogenetic analysis.

Methods

Blood samples from 18 domestic cats, submitted for diagnostic purposes by private veterinary practitioners to the Department of Veterinary Tropical Diseases, Faculty of Veterinary Science, University of Pretoria, were included in the study (Table 1). Inclusion criteria were clinical signs of babesiosis, identification of piroplasms on blood smears and/or positive reverse line blot (RLB) hybridization assay results. Except for one specimen from Maputo, Mozambique, all samples originated from coastal areas in the Western Cape and KwaZulu-Natal provinces of South Africa (Fig. 4).
Table 1

List of domestic cat samples used, with details on the origin, microscopic examination of blood smears, RLB results and phylogenetic classification

Sample ID

Origin

Microscopy

RLB results

No. of clones

Phylogenetic classification

BF221

Cascades, KZN, RSA

Babesia spp.

Babesia genus-specific only

Not applicablea

B. leo

BF238

Durban, KZN, RSA

Babesia spp.

Babesia genus-specific only

Not applicable

B. leo

BF272

Hermanus, WC, RSA

No parasites seen

Babesia genus-specific only

Not applicable

B. felis

BF284

Bellville, WC, RSA

Large Babesia spp.

Babesia genus-specific only

Not applicable

B. felis

BF341

Durban, KZN, RSA

Babesia spp.

Babesia genus-specific only

Not applicable

B. leo

BF342

Bellville, WC, RSA

Large Babesia spp.

Negative/Below detection limit

Not applicable

Novel Babesia sp. variant 1

BF461

Maputo, Mozambique

Babesia spp.

Babesia genus-specific only

Not applicable

B. leo

BF472

Durban, KZN, RSA

Large Babesia spp.

B. felis

Not applicable

Novel Babesia sp. variant 3

BF475

Durban, KZN, RSA

Babesia spp.

B. felis

Not applicable

B. felis

Cat01

Cape Town, WC, RSA

Large Babesia spp.

B. felis

2

Novel Babesia sp. variant 1

    

6

Novel Babesia sp. variant 2

Cat02

Cape Town, WC, RSA

Babesia spp.

B. felis, B microti

1

B. microti

    

5

Novel Babesia sp. variant 1

    

5

Novel Babesia sp. variant 3

Cat03

Cape Town, WC, RSA

Babesia spp.

Not tested

5

Novel Babesia sp. variant 1

    

1

Novel Babesia sp. variant 2

Cat05

Sedgefield, WC, RSA

Babesia spp.

B. felis

8

B. felis

Cat06

Sedgefield, WC, RSA

Babesia spp.

B. felis

9

B. felis

Cat07

Pietermaritzburg, KZN, SA

Large Babesia spp.

Not tested

1

B. leo

Cat08

Paarl, WC, RSA

Large Babesia spp.

Not tested

1

Novel Babesia sp. variant 2

Cat09

Paarl, WC, RSA

Large Babesia spp.

Not tested

1

Novel Babesia sp. variant 1

Cat10

Durban, KZN, RSA

Babesia spp.

Not tested

1

B. felis

aPCR amplicon directly sequenced (not subjected to cloning)

Abbreviations: KZN, KwaZulu-Natal; RSA, Republic of South Africa; WC, Western Cape Province

Fig. 4
Fig. 4

Map of Southern Africa showing the origin of the samples

DNA was extracted according to the manufacturer’s instructions using the QIAamp® DNA Mini Kit (Qiagen, Whitehead Scientific, South Africa). The V4 hypervariable region of the parasite 18S rRNA gene was PCR amplified using Babesia and Theileria genus-specific primers RLB-F2 and biotin-labelled RLB-R2 [24, 25]; PCR reaction conditions were as described by Tembo et al. [26]. DNA extracted from blood from a known T. parva-infected buffalo [27] was used as a positive control, while PCR master mix without DNA was used as a negative control. A touch down thermal cycler programme was used to amplify the DNA [25]. The PCR products were then analysed using the RLB hybridization technique as previously described [24, 25, 28, 29]. Genus- and species-specific probes as described by Tembo et al. [26] were included on the membrane; in addition to this, a B. lengau probe [22] was also included.

The near full-length parasite 18S rRNA gene (~1700 bp) was PCR amplified using primers Nbab_1F [30] and TB Rev [31], as previously described by Bosman et al. [22]. Four separate reactions were prepared per sample. Amplicons of all four reactions per sample were pooled to avoid Taq polymerase-induced errors and purified using a QIAquick PCR purification kit (Qiagen, Southern Cross Biotechnology, South Africa) according to the manufacturer’s instructions. Nine of the samples (labelled BF; Table 1) that had been yielded positive RLB results in a previous study [15], were subjected to direct (bi-directional) sequencing on an ABI 3500XL genetic analyser using the amplification primers. For the other nine specimens, PCR amplicons were cloned prior to sequencing (in case of mixed infections not being detected or masked by the RLB assay) into the pGEM-T Easy vector (Promega, Anatech, South Africa) and transformed into competent Escherichia coli JM109 cells (JM109 high-efficiency competent cells, Promega). Recombinant plasmids were directly (bi-directional) sequenced on the ABI 3500XL genetic analyser at Inqaba Biotechnical Industries using the vector primers SP6 and T7.

Sequences were assembled and edited using GAP 4 of the Staden package (Version 1.6.0 for Windows) [32]. A search for homologous sequences was performed using BLASTn [33]. The sequences were aligned with sequences of related genera from GenBank using ClustalX (Version 1.81 for Windows). Alignment files were also analysed with CLC Main Workbench version 4.0 (CLC bio, Aarhus, Denmark) to test consistency of the alignment. The alignment was manually truncated to the size of the smallest sequence (1421 bp). The genetic distances between the sequences were estimated by determining the number of nucleotide differences between sequences using MEGA version 7 [34]. Phylogenetic trees were constructed by the Neighbor-Joining (NJ) and Maximum Likelihood (ML) methods as implemented in MEGA 7. The two-parameter model of Kimura [35] was used to construct similarity matrices by single distance from the aligned sequence data; a NJ phylogenetic tree [36] was constructed in combination with the bootstrap method (1000 replicates/tree) [37]. The Hasegawa-Kishino-Yano (HKY + G + I) substitution model [38], determined as the best-fit model using MEGA 7, was used to infer a ML tree in combination with the bootstrap method (1000 replicates/tree) [37]. The 18S rDNA sequences of Cardiosporidium ciona (EU052685), the closest species for which data are available according to Schnittger et al. [39], was included as the outgroup. All consensus trees were edited using MEGA 7. The GenBank accession numbers of reference sequences used in this study are reported in Table 2. The 18S rRNA gene sequences obtained in this study were submitted to GenBank; the accession numbers are reported in Table 3.
Table 2

Accession numbers for GenBank reference sequences used in the present study

GenBank ID

Species

Origin

Host

References

AY072926

B. canis

Croatia

Dog

Caccio et al. [52]

AY272047

B. canis presentii

Israel

Cat

Baneth et al. [18]

AF158702

B. conradae

USA

Dog

Kjemtrup et al. [53]

U16370

B. divergens

USA

Cattle

Holman [54]

AF158700

B. duncani

USA

Human

Kjemtrup et al. [55]

AF244912

B. felis

South Africa

Domestic cat

Penzhorn et al. [40]

AY278443

B. gibsoni

Spain

Dog

Criado-Fornelio et al. [56]

GQ411417

B. lengau

South Africa

Cheetah

Bosman et al. [22]

KC790443

B. lengau

South Africa

Domestic cat

Bosman et al. [23]

KC833036

B. lengau

South Africa

Domestic cat

Bosman et al. [23]

AF244911

B. leo

South Africa

Lion

Penzhorn et al. [40]

AY452708

B. leo

South Africa

Domestic cat

Wuerth (unpubl.)

AB071177

B. microti (Munich)

Europe

Human

Tsuji et al. (unpubl.)

AB119446

B. microti (Otsu)

Japan

Field rodent

Saito-Ito et al. [57]

AF231348

B. microti (GI)

USA

Human

Zahler et al. [58]

AY693840

B. microti (Gray)

USA

Human

Slemenda et al. (unpubl.)

XR002459986

B. microti (R1)

USA

Human

Cornillot et al. [59]

U16369

B. odocoilei

USA

Cervid

Holman et al. [60]

AY661502

B. odocoilei

USA

Bighorn sheep

Schoelkopf et al. (unpubl.)

M87565

B. rodhaini

Australia

Cell culture

Ellis et al. [61]

DQ111760

B. rossi

Sudan

Dog

Oyamada et al. [62]

AY190123

Babesia sp. Akita610 Dog tick

Japan

Ixodes ovatus

Inokuma et al. [46]

AB251608

Babesia sp. MA#230

Japan

Raccoon

Jinnai et al. [44]

KR017880

Babesia sp. Maned wolf

USA

Maned wolf

Wasserkrug Naor et al. [42]

AB935172

Babesia sp. YA23175

Japan

Raccoon

Komura et al. (unpubl.)

AB935330

Babesia sp. SW-R-090616_T1

Japan

Raccoon

Hirata et al. (unpubl.)

AB935331

Babesia sp. SW-R-092616_T2

Japan

Raccoon

Hirata et al. (unpubl.)

DQ028958

Babesia sp. AJB-2006

USA

Raccoon

Birkenheuer et al. (unpubl.)

KX218429

Babesia sp. 1 1093 cl9

Botswana

Lion

McDermid et al. [63]

KX218430

Babesia sp. 10 1092 cl9

Botswana

Lion

McDermid et al. [63]

KX218431

Babesia sp. 3 1093 cl8

Botswana

Lion

McDermid et al. [63]

KX218432

Babesia sp. 4 1093 cl2

Botswana

Lion

McDermid et al. [63]

KX218433

Babesia sp. 5 1093 c17

Botswana

Lion

McDermid et al. [63]

KX218434

Babesia sp. 6 1092 cl1

Botswana

Lion

McDermid et al. [63]

KX218435

Babesia sp. 7 1092 cl3

Botswana

Lion

McDermid et al. [63]

KX218436

Babesia sp. 8 1092 cl5

Botswana

Lion

McDermid et al. [63]

KX218437

Babesia sp. 9 1093 cl1

Botswana

Lion

McDermid et al. [63]

KX218438

Babesia sp. 10 1092 cl9

Botswana

Lion

McDermid et al. [63]

KX218439

Babesia sp. 11 1095

Botswana

Lion

McDermid et al. [63]

KX218440

Babesia sp. 12 1101

Botswana

Lion

McDermid et al. [63]

AF244913

Babesia sp. Strain A Caracal

South Africa

Caracal

Penzhorn et al. [40]

AF244914

Babesia sp. Strain B Caracal

South Africa

Caracal

Penzhorn et al. [40]

KF724377

B. venatorum

China

Human

Sun et al. [64]

AY072925

B. vogeli

Italy

Dog

Caccio et al. [52]

EU052685

Cardiosporidium cionae

Ciona intestinalis

Ciancio et al. [65]

Table 3

Accession numbers for the 18S rRNA gene sequences generated in the present study

GenBank ID

Sample

Phylogenetic classification

Origin

KC790441

BF461a

B. leo

Maputo, Mozambique

KC790442

BF472

Babesia sp. Variant3

Durban, KZN, RSA

KC790444

BF341Aa

B. leo

Durban, KZN, RSA

KR611115

Cat05_8

B. felis

Sedgefield, WC, RSA

KR611116

Cat05_24

B. felis

Sedgefield, WC, RSA

KR611117

Cat05_18

B. felis

Sedgefield, WC, RSA

KR611118

Cat05_14

B. felis

Sedgefield, WC, RSA

KR611119

Cat05_13

B. felis

Sedgefield, WC, RSA

KR611120

Cat05_12

B. felis

Sedgefield, WC, RSA

KR611121

Cat05_6

B. felis

Sedgefield, WC, RSA

KR611122

Cat06_H5

B. felis

Sedgefield, WC, RSA

KR611123

Cat06_G5

B. felis

Sedgefield, WC, RSA

KR611124

Cat06_D5

B. felis

Sedgefield, WC, RSA

KR611125

Cat06_C5

B. felis

Sedgefield, WC, RSA

KR611126

Cat06_B6

B. felis

Sedgefield, WC, RSA

KR611127

Cat06_A6

B. felis

Sedgefield, WC, RSA

KR611128

Cat06_A5

B. felis

Sedgefield, WC, RSA

KR611129

Cat06_B5

B. felis

Sedgefield, WC, RSA

KR611130

Cat06_F5

B. felis

Sedgefield, WC, RSA

KR611131

Cat05_11

B. felis

Sedgefield, WC, RSA

KR611132

Cat07_5E

B. leo

Pietermaritzburg, KZN, RSA

KR611133

Cat03_5

Babesia sp. Variant1

Cape Town, WC, RSA

KR611134

Cat03_10

Babesia sp. Variant1

Cape Town, WC, RSA

KR611135

Cat03_3

Babesia sp. Variant1

Cape Town, WC, RSA

KR611136

Cat03_1

Babesia sp. Variant1

Cape Town, WC, RSA

KR611137

Cat02_3

Babesia sp. Variant1

Cape Town, WC, RSA

KR611138

Cat03_9

Babesia sp. Variant1

Cape Town, WC, RSA

KR611139

Cat02_6

Babesia sp. Variant1

Cape Town, WC, RSA

KR611140

Cat02_10

Babesia sp. Variant1

Cape Town, WC, RSA

KR611141

Cat02_12

Babesia sp. Variant1

Cape Town, WC, RSA

KR611142

Cat03_8

Babesia sp. Variant2

Cape Town, WC, RSA

KR611143

Cat02_4

Babesia sp. Variant1

Cape Town, WC, RSA

KR611144

Cat02_2

Babesia sp. Variant3

Cape Town, WC, RSA

KR611145

Cat02_1

Babesia sp. Variant3

Cape Town, WC, RSA

KR611146

Cat02_9

Babesia sp. Variant3

Cape Town, WC, RSA

KR611148

Cat02_13

Babesia sp. Variant3

Cape Town, WC, RSA

KR611149

Cat01_G

Babesia sp. Variant1

Cape Town, WC, RSA

KR611150

Cat01_K

Babesia sp. Variant2

Cape Town, WC, RSA

KR611151

Cat01_F

Babesia sp. Variant1

Cape Town, WC, RSA

KR611152

Cat01_A

Babesia sp. Variant2

Cape Town, WC, RSA

KR611153

Cat01_J

Babesia sp. Variant2

Cape Town, WC, RSA

KR611154

Cat01_I

Babesia sp. Variant2

Cape Town, WC, RSA

KR611155

Cat01_B

Babesia sp. Variant2

Cape Town, WC, RSA

KR611156

Cat01_E

Babesia sp. Variant2

Cape Town, WC, RSA

KR611158

Cat08_13

Babesia sp. Variant2

Paarl, WC, RSA

KR611159

Cat02_4b

Babesia sp. Variant3

Cape Town, WC, RSA

KR732967

BF475

B. felis

Durban, KZN, RSA

KR732968

BF284

B. felis

Bellville, WC, RSA

KR732969

BF272

B. felis

Hermanus, WC, RSA

KR732970

BF342

Babesia sp. Variant1

Bellville, WC, RSA

KR732971

BF461Aa

B. leo

Maputo, Mozambique

KR732972

BF341a

B. leo

Durban, KZN, RSA

KR732973

BF238

B. leo

Durban, KZN, RSA

KR732974

BF221

B. leo

Cascades, KZN, RSA

KT182985

Cat10_14_11

B. felis

Durban, KZN, RSA

KT182986

Cat9_8

Babesia sp. Variant1

Paarl, WC, RSA

MK095342

46Cat02_10bb

B. microti

Cape Town, WC, RSA

MK095343

46Cat02_10b_newb

B. microt

Cape Town, WC, RSA

aDuplicate samples received per animal (BF341 and 461)

bSequences derived from the same clone (46Cat02_10b and 46Cat02_10b_new)

Abbreviations: KZN, KwaZulu-Natal; RSA, Republic of South Africa; WC, Western Cape Province

Results

Clinical reports indicated that 15 cats showed severe clinical signs of babesiosis, e.g. lethargy, anaemia, icterus and fever. Although no detailed clinical reports were available for three cats (BF341, BF472 and BF455) (Table 1), the attending veterinarians had made tentative diagnoses of babesiosis. With the exception of one cat (BF272), organisms morphologically consistent with piroplasms were seen on microscopic examination of blood smears from 17 of the cats; seven of these had been reported as a “large” Babesia (Table 1).

The RLB hybridization assay results revealed that of the 13 samples tested, six (46.2%) tested positive for the presence of B. felis DNA. One of these samples (Cat02) had a mixed species infection with B. microti (Table 1). This was subsequently confirmed by cloning and sequencing analysis of the 18S rRNA gene. PCR amplicons from a further six samples (46.2%) hybridized with the Babesia genus-specific probe only, suggesting the presence of a potentially novel Babesia species. One sample (BF342) tested negative or below the detection limit of the assay although a large Babesia had been observed by microscopy.

A total of 55 nearly full-length (1484–1525 bp) parasite 18S rRNA gene sequences were obtained from the 18 samples. Of these, nine were directly sequenced and the rest were cloned prior to sequencing, yielding a further 46 sequences from the clones (Table 1). A BLASTn search revealed that sequences from six cats (two of from Durban, KwaZulu-Natal, and four from the Western Cape) were most closely related to a published 18S rRNA gene sequence of B. felis (AF244912) which was previously described from a domestic cat and caused severe clinical babesiosis in naturally and experimentally infected cats in South Africa [11, 13]. One of these 21 sequences (Cat06_A6) had 100% sequence identity to the published B. felis sequence, while the remaining sequences had 99% identity, differing by one nucleotide from the published B. felis 18S rRNA gene sequence over a 1525 bp region.

Sequences from four cats had 100% identity with published B. leo sequences, while one sequence (Cat07_5E) had 99% identity with B. leo (with a 3 nucleotide difference over a 1520 bp region). Babesia leo was previously described from lions in the Kruger National Park, South Africa, and was shown to be a distinct species from B. felis and other felid piroplasms [40]. One specimen was from Maputo, Mozambique, the other four being from KwaZulu-Natal, i.e. all on the north-eastern seaboard of southern Africa.

One sequence (Cat02new) had 98–100% sequence identity with published B. microti 18S rRNA gene sequences, including strains from the zoonotic B. microti lineages (USA, Munich, Kobe and Otsu/Hobetsu from Japan). It had 100% sequence identity to the published B. microti Otsu strain (AB119446) and differed by 3–6 nucleotides from the B. microti Gray (AY693840) and B. microti Munich (AB071177) strains, respectively.

The most interesting finding, however, was that sequences obtained from seven cats, six from the Western Cape Province and one from Durban, KwaZulu-Natal, constituted a novel Babesia group with 96% identity to Babesia spp. previously described from captive maned wolves (Chrysocyon brachyurus) [41, 42], raccoons (Procyon lotor) from the USA [43] and Japan [44, 45] and from ticks collected from dogs in Japan [46]. Three genetic variants were identified within this novel Babesia group (designated “Novel Babesia sp. genetic variants 1, 2 and 3”), differing by 1 to 3 nucleotides from each other. Genetic variant 1 was found in five cats, variant 2 in three cats and variant 3 in two cats (Table 2). Three cats were infected with two genetic variants: two with variants 1 and 2, and one with variants 1 and 3.

The observed sequence similarities were subsequently confirmed by phylogenetic analyses. NJ and ML analyses were used to reveal the phylogenetic relationships between the near full-length 18S rRNA gene sequences obtained from this study to related Babesia species previously deposited in GenBank (Table 1). The topologies of both trees were similar. The ML tree is shown in Fig. 5. Three distinct clades, in concordance with Schnittger et al. [39], were obtained representing Clade I (including rodent-infecting B. microti and B. rodhaini, and feline-infecting B. leo and B. felis parasites), Clade II (including B. duncani isolated from humans, canine B. conradae and B. lengau described from cheetah in South Africa) and Clade VI (Babesia (s.s.), including the canine-infecting B. gibsoni, B. canis, B. rossi and B. vogeli, the human-infecting isolate B. venatorum, as well as species infecting ungulates (such as B. divergens and B. odocoilei) and recently described Babesia species infecting other carnivores such as bears, cougars and raccoons, as well as field rodents. The novel Babesia species identified in this study grouped within Clade VI, also referred to as the “carnivore/rodent clade” by Schnittger et al. [39].
Fig. 5
Fig. 5

Maximum likelihood tree showing the evolutionary relationships of the Babesia 18S rDNA sequences obtained, with published sequences. The evolutionary history was inferred by using the Maximum Likelihood method based on the Hasegawa-Kishino-Yano model [38]. A discrete Gamma distribution was used to model evolutionary rate differences among sites [5 categories (+G, parameter = 0.4367)]. The rate variation model allowed for some sites to be evolutionarily invariable [(+I), 57.75% sites]. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. All positions containing gaps and missing data were eliminated. There were a total of 1208 positions in the final dataset. Evolutionary analyses were conducted in MEGA7 [34]

Discussion

The B. felis-positive specimens were from both the Western Cape (n = 4) and KwaZulu-Natal (n = 2). There is a single report of B. leo from a sick cat, but it was a mixed infection with B. felis [15]. The results of the present study unambiguously implicate B. leo in causing clinical babesiosis in domestic cats. The B. leo-positive specimens were all from the north-eastern seaboard of southern Africa: KwaZulu-Natal (n = 4) and Maputo, Mozambique (n = 1), which constituted the first record of B. leo from that country. The Kruger National Park, South Africa, from where B. leo was first described [40], has a 320-km-long border with Mozambique. In a direct line, the south-eastern tip of the Park is only c.70 km from Maputo.

Sequence and phylogenetic analysis of the 18S rRNA gene from seven cats showed that they harboured a novel Babesia sp. which segregated into three separate genetic variants in Babesia clade VI, the carnivore/rodent clade [39]. Babesia felis, B. leo and B. lengau, the three South African felid piroplasms hitherto known to cause clinical signs in domestic cats, were relatively closely related [15, 22, 40]. In contrast, the novel Babesia sp. reported here had only 92% sequence identity with B. felis (AF244912) and 89% sequence identity with B. leo (AF244911) and B. lengau (KC790443 and GQ411417), respectively. The 18S rRNA gene has been widely used to characterize and classify previously unknown Theileria and Babesia parasites [24, 25, 30, 4749]. It has, however, not been established to what extent 18S rRNA gene sequences must differ for the source organisms to be considered different species, rather than merely a genetic variant or genotype within a species [50, 51]. A single gene tree does not necessarily reflect a species tree [39]; therefore, a tree should ideally be constructed using multiple genotypic characters of potentially different evolutionary histories [39].

The novel genetic variants reported here were most closely related (96% identity) to a novel Babesia sp. reported from culled feral raccoons from Japan [44, 45] and from a clinically affected juvenile raccoon from the USA [43]. It is tempting to speculate that feral raccoons may also have been the source of an incidental finding of this Babesia sp. in ticks collected from healthy dogs in Japan [46]. The same Babesia sp. was incriminated in causing severe clinical babesiosis in two South American maned wolves from the same zoological park in Kansas, USA [41, 42].

When examining blood smears, veterinarians described the novel genetic variants reported here as “large” babesias. This may be the elusive large Babesia reported from cats in southern Africa. The arbitrary classification of babesias as either “large” or “small” is not satisfactory, however. For instance, the abovementioned Babesia sp. from raccoons was reported to be closely related to B. odocoilei and B. divergens [44], both generally regarded as “large” species. Nevertheless, the mean length of the round, oval, amoeboid or piriform organisms was 3.13 ± 0.77 µm (range 1.25–4.8 µm) and the mean width was 2.5 ± 0.61 µm [45]. Round, oval and amoeboid forms are trophozoites, which can be expected to increase in size. For comparative purposes, measuring newly formed merozoites should give more consistent and reliable results.

Six of the seven specimens of the novel genetic variants were from a fairly restricted area in the Western Cape Province (Bellville, Cape Town and Paarl). The other case was from Durban, KwaZulu-Natal. No further information was known about the latter case, e.g. whether the cat may originally have come the Cape Town area. It may be possible that the natural hosts and/or vectors of these novel genetic variants are restricted to the Western Cape Province. Due to lacking an appropriate type specimen, we refrain from describing a new species but refer to the novel organism as Babesia sp. cat Western Cape.

Further characterisation of this novel organism is warranted to understand the pathogenesis and epidemiology, as well as to develop appropriate diagnostic markers. Obtaining appropriate specimens poses a challenge, however. Veterinarians in the feline babesiosis-endemic area usually confirm a diagnosis by finding piroplasms on a blood smear and then treat the cat. Blood specimens are only rarely submitted for confirmation of a diagnosis. Furthermore, our laboratory is in Pretoria, c.600 km from Durban and 1500 km from Cape Town, which hampers routine sampling of clinical cases.

Conclusions

Our results indicate the presence of a novel potentially pathogenic Babesia sp. in felids in South Africa, which is not closely related to Babesia felis, Babesia lengau and Babesia leo, the three species known to be pathogenic to cats. Due to the lack of an appropriate type-specimen, we refrain from describing and a new species but refer to the novel organism as Babesia sp. Cat Western Cape.

Declarations

Acknowledgements

We thank Mr Christiaan Labuschagne and Dr Antoinette van Schalkwyk (Inqaba Biotechnical Industries (Pty) Ltd, Pretoria, South Africa), for their assistance with the sequence analysis, and the veterinarians who submitted specimens for this study, especially Drs Remo Lobetti and Fred Reyers. Professor Melvyn Quan prepared Fig. 4. Publication of this paper has been sponsored by Bayer Animal Health in the framework of the 14th CVBD World Forum Symposium.

Funding

This research was supported financially by the National Research Foundation (NRF), South Africa (grant no 2069496 to BL Penzhorn and grant no. 76529 to MCO).

Availability of data and materials

Data supporting the conclusions of this article are included within the article. The newly generated sequences were submitted to the GenBank database under the accession numbers provided in Table 2.

Authors’ contributions

AMB screened the samples with the reverse line blot, carried out the molecular genetic studies, participated in the sequence alignment and wrote the first draft of the manuscript. BLP coordinated the investigation, conducted literature searches and reviewed and edited all drafts of the manuscript. KAB co-supervised the project, and reviewed and edited the manuscript. TS handled clinical cases and collected most of the specimens. MCO supervised the laboratory work and sequence alignments, constructed the phylogenetic trees, reviewed all drafts of the paper and phylogenetic results and wrote the final version of the manuscript. All authors read and approved the final manuscript.

Ethics approval and consent to participate

This study was approved by the Animal Ethics Committee of the University of Pretoria (ref V116-15) and by the Research Committee of the Faculty of Veterinary Science, University of Pretoria (ref 36-5-613). The South African Department of Agriculture, Forestry & Fisheries granted permission to do research in terms of Section 20 of the Animal Diseases Act, 1984 (Act no. 35 of 1984) (ref 12/11/1/1).

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

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Authors’ Affiliations

(1)
Vectors and Vector-borne Diseases Research Programme, Department of Veterinary Tropical Diseases, Faculty of Veterinary Science, University of Pretoria, Private Bag X04, Onderstepoort, 0110, South Africa
(2)
Centre for Wildlife Studies, Faculty of Veterinary Science, University of Pretoria, Private Bag X04, Onderstepoort, 0110, South Africa
(3)
National Zoological Garden, South African National Biodiversity Institute, PO Box 754, Pretoria, 0001, South Africa
(4)
Department of Veterinary Microbiology and Pathology, Washington State University, Pullman, Washington, USA
(5)
Department of Companion Animal Clinical Studies, Faculty of Veterinary Science, University of Pretoria, Private Bag X04, Onderstepoort, 0110, South Africa
(6)
Cape Animal Medical Centre, 78 Rosmead Avenue, Kenilworth, Cape Town, South Africa

References

  1. Driscoll CA, Menotti-Raymond M, Roca AL, Hupe K, Johnson WE, Geffen E, et al. The Near Eastern origin of cat domestication. Science. 2007;317:519–23.View ArticleGoogle Scholar
  2. Driscoll CA, Macdonald DW, O’Brien SJ. From wild animals to domestic pets, an evolutionary view of domestication. Proc Natl Acad Sci USA. 2009;106(Suppl. 1):9971–8.View ArticleGoogle Scholar
  3. Jacobson LS, Schoeman T, Lobetti RG. A survey of feline babesiosis in South Africa. J S Afr Vet Assoc. 2000;71:222–8.PubMedGoogle Scholar
  4. Penzhorn BL, Stylianides E, Coetzee MA, Viljoen JM, Lewis BD. A focus of feline babesiosis at Kaapschehoop on the Mpumalanga escarpment. J S Afr Vet Assoc. 1999;70:60.View ArticleGoogle Scholar
  5. Davis LJ. On a piroplasm of the Sudanese wild cat (Felis ocreata). Trans R Soc Trop Med Hyg. 1929;12:523–33.Google Scholar
  6. Wenyon CM. Protozoology. London: Baillière, Tindall & Cox; 1926.Google Scholar
  7. Jackson C, Dunning FJ. Biliary fever (nuttaliosis) of the cat: a case in the Stellenbosch district. J S Afr Vet Med Assoc. 1937;8:83–7.Google Scholar
  8. McNeil J. Piroplasmosis of the domestic cat. J S Afr Vet Med Assoc. 1937;8:88–90.Google Scholar
  9. Du Toit PJ. Zur Systematik der·Piroplasmen. Arch Protist. 1918;39:81–104.Google Scholar
  10. Carpano M. Sulle piroplasmi dei carnivori e su di un nuovo piroplasma de felini (Babesiella felis) nel puma: Felis concolor. Bollettino No. 137. Cairo: Serviz Tecn Scient Min Dell Agric.; 1934.Google Scholar
  11. Futter GJ, Belonje PC. Studies on feline babesiosis. 2. Clinical findings. J S Afr Vet Assoc. 1980;51:143–6.PubMedGoogle Scholar
  12. Potgieter FT. Chemotherapy of Babesia felis infection: efficacy of certain drugs. J S Afr Vet Assoc. 1980;51:289–93.Google Scholar
  13. Penzhorn BL, Lewis BD, López-Rebollar LM, Swan GE. Screening of five drugs for efficacy against Babesia felis in experimentally infected cats. J S Afr Vet Assoc. 2000;71:53–7.PubMedGoogle Scholar
  14. Benson DA, Cavanaugh M, Clark K, Karsch-Mizrachi I, Lipman DJ, Ostell J, et al. GenBank. Nucl Acids Res. 2013;41(Database issue):D36–42.PubMedGoogle Scholar
  15. Bosman AM, Venter EH, Penzhorn BL. Occurrence of Babesia felis and Babesia leo in various wild felid species and domestic cats in southern Africa, based on reverse line blot analysis. Vet Parasitol. 2007;144:33–8.View ArticleGoogle Scholar
  16. Williams BM, Berentsen A, Shock BC, Teixiera M, Dunbar MR, Becker MS, et al. Prevalence and diversity of Babesia, Hepatozoon, Ehrlichia, and Bartonella in wild and domestic carnivores from Zambia, Africa. Parasitol Res. 2014;113:911–8.View ArticleGoogle Scholar
  17. Stewart CG, Hackett KJW, Collett MG. An unidentified Babesia of the domestic cat (Felis domesticus). J S Afr Vet Assoc. 1980;51:219–22.PubMedGoogle Scholar
  18. Baneth G, Kenny MJ, Tasker S, Anug Y, Shkap V, Levy A, et al. Infection with a proposed new subspecies of Babesia canis, Babesia canis subsp. presentii, in domestic cats. J Clin Microbiol. 2004;42:99–105.View ArticleGoogle Scholar
  19. Dennig HK. Babesieninfektionen bei exotischer Katzen and die Bedeutung dieser Blutparasiten für die tierärtzliche Forschung. Acta Zool Pathol Antverpiensia. 1969;48:361–7.Google Scholar
  20. Dennig HK, Brocklesby DW. Babesia pantherae sp. nov., a piroplasm of the leopard (Panthera pardus). Parasitology. 1972;64:525–32.View ArticleGoogle Scholar
  21. Dennig HK. Eine unbekannte Babesienart beim Jaguarundi (Herpailurus yaguarondi). Kleintierpraxis. 1967;12:146–50.Google Scholar
  22. Bosman AM, Oosthuizen EC, Peirce MA, Venter EH, Penzhorn BL. Babesia lengau sp. nov., a novel Babesia species in cheetah (Acinonyx jubatus Schreber, 1775) populations in South Africa. J Clin Microbiol. 2010;48:2703–8.View ArticleGoogle Scholar
  23. Bosman AM, Oosthuizen MC, Venter EH, Steyl JC, Gous TA, Penzhorn BL. Babesia lengau associated with cerebral and haemolytic babesiosis in two domestic cats. Parasit Vectors. 2013;6:128.View ArticleGoogle Scholar
  24. Nijhof AM, Penzhorn BL, Lynen G, Mollel JO, Morkel P, Bekker CPJ, et al. Babesia bicornis sp. nov. and Theileria bicornis sp. nov.: tick-borne parasites associated with mortality in the black rhinoceros. J Clin Microbiol. 2003;41:2249–54.View ArticleGoogle Scholar
  25. Nijhof AM, Pillay V, Steyl J, Prozesky L, Stoltsz WH, Lawrence JA, et al. Molecular characterization of Theileria species associated with mortalities in four species of African antelopes. J Clin Microbiol. 2005;43:5907–11.View ArticleGoogle Scholar
  26. Tembo S, Collins NE, Sibeko-Matjila KP, Troskie M, Vorster I, Byaruhanga C, et al. Occurrence of tick-borne haemoparasites in cattle in the Mungwi District, Northern Province, Zambia. Ticks Tick Borne Dis. 2018;9:707–17.View ArticleGoogle Scholar
  27. Sibeko KP, Oosthuizen MC, Collins NE, Geysen D, Rambritch NE, Latif AA, et al. Development and evaluation of a real-time polymerase chain reaction test for the detection of Theileria parva infections in Cape buffalo (Syncerus caffer) and cattle. Vet Parasitol. 2008;155:37–48.View ArticleGoogle Scholar
  28. Bekker CPJ, de Vos S, Taoufik A, Sparagano OAE, Jongejan F. Simultaneous detection of Anaplasma and Ehrlichia species in ruminants and detection of Ehrlichia ruminantium in Amblyomma variegatum ticks by reverse line blot hybridization. Vet Microbiol. 2000;89:223–38.View ArticleGoogle Scholar
  29. Gubbels J, De Vos A, Van Der Weide M, Viseras J, Schouls LM, De Vries E, et al. Simultaneous detection of bovine Theileria and Babesia species by reverse line blot hybridization. J Clin Microbiol. 1999;37:1782–9.PubMedPubMed CentralGoogle Scholar
  30. Oosthuizen MC, Zweygardt E, Collins NE, Troskie M, Penzhorn BL. Identification of a novel Babesia sp. from sable antelope (Hippotragus niger Harris, 1838). Vet Parasitol. 2008;163:39–46.View ArticleGoogle Scholar
  31. Matjila PT, Leisewitz AL, Oosthuizen MC, Jongejan F, Penzhorn BL. Detection of Theileria species in dogs in South Africa. Vet Parasitol. 2008;157:34–40.View ArticleGoogle Scholar
  32. Staden R, Beal KF, Bonfield JK. The staden package, 1998. Methods Mol Biol. 2000;132:115–30.PubMedGoogle Scholar
  33. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215:403–10.View ArticleGoogle Scholar
  34. Kumar S, Stecher G, Tamura K. MEGA7: Molecular Evolutionary Genetics Analysis version 7.0 for bigger datasets. Mol Biol Evol. 2016;33:1870–4.View ArticleGoogle Scholar
  35. Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16:111–20.View ArticleGoogle Scholar
  36. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–25.PubMedGoogle Scholar
  37. Felsenstein J. Confidence limits on phylogenies: an approach using the bootstrap. Evolution. 1985;39:783–91.View ArticleGoogle Scholar
  38. Hasegawa M, Kishino H, Yano T. Dating the human-ape split by a molecular clock of mitochondrial DNA. J Mol Evol. 1985;22:160–74.View ArticleGoogle Scholar
  39. Schnittger L, Rodriguez AE, Florin-Christensen M, Morrison DA. Babesia: a world emerging. Inf Gen Evol. 2012;12:1788–809.View ArticleGoogle Scholar
  40. Penzhorn BL, Kjemtrup AM, López-Rebollar LM, Conrad PA. Babesia leo n. sp. from lions in the Kruger National Park, South Africa, and its relation to other piroplasms. J Parasitol. 2001;87:881–5.View ArticleGoogle Scholar
  41. Phair K, Carpenter JW, Smee N, Myers CB, Pohlman LM. Severe anemia caused by babesiosis in a maned wolf (Chrysocyon brachyurus). J Zoo Wildl Med. 2012;43:162–7.View ArticleGoogle Scholar
  42. Wasserkrug Naor A, Lindemann DM, Schreeg ME, Marr HS, Birkenheuer AJ, Carpenter JW, et al. Clinical, morphological, and molecular characterization of an undetermined Babesia species in a maned wolf (Chrysocyon brachyurus). Ticks Tick Borne Dis. 2019;10:124–6.View ArticleGoogle Scholar
  43. Birkenheuer AJ, Whittington J, Neel J, Large E, Barger A, Levy MG, et al. Molecular characterization of a Babesia species identified in a North American raccoon. J Wildl Dis. 2006;42:375–80.View ArticleGoogle Scholar
  44. Jinnai M, Kawabuchi-Kurata T, Tsuji M, Nakajima R, Fujisawa K, Nagata S, et al. Molecular evidence for the presence of new Babesia species in feral raccoons (Procyon lotor) in Hokkaido, Japan. Vet Parasitol. 2009;162:241–7.View ArticleGoogle Scholar
  45. Kawabuchi T, Tsuki M, Sado A, Matoba Y, Asakawa M, Ishihara C. Babesia microti-like parasites detected in feral raccoons (Procyon lotor) captured in Hokkaido, Japan. Jpn J Vet Med Sci. 2005;67:825–7.View ArticleGoogle Scholar
  46. Inokuma H, Yoshizaki Y, Shimada Y, Sakata Y, Okuda M, Onishi T. Epidemiological survey of Babesia species in Japan performed with specimens from ticks collected from dogs and detection of new Babesia DNA closely related to Babesia odocoilei and Babesia divergens DNA. J Clin Microbiol. 2003;41:3494–8.View ArticleGoogle Scholar
  47. Gubbels M, Hong Y, van der Weide M, Qi B, Nijman IS, Guangyuan L, et al. Molecular characterisation of the Theileria buffeli/orientalis group. Int J Parasitol. 2000;30:943–52.View ArticleGoogle Scholar
  48. Birkenheuer AJ, Levy MG, Breitschwerdt EB. Development and evaluation of a seminested PCR for detection and differentiation of Babesia gibsoni (Asian genotype) and B. canis DNA in canine blood samples. J Clin Microbiol. 2003;41:4172–7.View ArticleGoogle Scholar
  49. Schnittger L, Yin H, Gubbels MJ, Beyer D, Niemann S, Jongejan F, et al. Phylogeny of sheep and goat Theileria and Babesia parasites. Parasitol Res. 2003;91:398–406.View ArticleGoogle Scholar
  50. Chae JS, Allsopp BA, Waghela SD, Park JH, Kakuda T, Sugimoto C, et al. A study of the systematics of Theileria spp. based upon small-subunit ribosomal RNA gene sequences. Parasitol Res. 1999;85:877–83.View ArticleGoogle Scholar
  51. Allsopp MT, Allsopp BA. Molecular sequence evidence for the reclassification of some Babesia species. Ann NY Acad Sci. 2006;1081:509–17.View ArticleGoogle Scholar
  52. Caccio SM, Antunovic B, Moretti A, Mangili V, Marinculic A, Slemenda SB, et al. Molecular characterisation of Babesia canis canis and Babesia canis vogeli from naturally infected European dogs. Vet Parasitol. 2002;106:285–92.View ArticleGoogle Scholar
  53. Kjemtrup AM, Wainwright K, Miller M, Penzhorn BL, Carreno RA. Babesia conradae, sp. nov., a small canine Babesia identified in California. Vet Parasitol. 2006;138:103–11.View ArticleGoogle Scholar
  54. Holman PJ. Comparative study of cervid Babesia isolates. Thesis, Veterinary Pathobiology, Texas A&M University, College Station, TX, USA; 1994.Google Scholar
  55. Kjemtrup AM, Thomford J, Robinson T, Conrad PA. Phylogenetic relationships of human and wildlife piroplasm isolates in the western United States inferred from the 18S nuclear small subunit RNA gene. Parasitology. 2000;120:487–93.View ArticleGoogle Scholar
  56. Criado-Fornelio A, Gonzalez-del-Rio MA, Buling-Sarana A, Barba-Carretero JC. Molecular characterization of a Babesia gibsoni isolate from a Spanish dog. Vet Parasitol. 2003;117:123–9.View ArticleGoogle Scholar
  57. Saito-Ito A, Dantrakool A, Kawai A, Shiota T. Survey of rodents and ticks in human babesiosis emergence area in Japan: first detection of Babesia microti-like parasites in Ixodes ovatus. J Clin Microbiol. 2004;42:2268–70.View ArticleGoogle Scholar
  58. Zahler M, Rinder H, Gothe R. Genotypic status of Babesia microti within the piroplasms. Parasitol Res. 2000;86:642–6.View ArticleGoogle Scholar
  59. Cornillot E, Hadj-Kaddour K, Dassouli A, Noel B, Ranwez V, Vacherie B, et al. Sequencing of the smallest Apicomplexan genome from the human pathogen Babesia microti. Nucl Acids Res. 2012;40:9102–14.View ArticleGoogle Scholar
  60. Holman PJ, Madeley J, Craig TM, Allsopp BA, Allsopp MT, Petrini KR, et al. Antigenic, phenotypic and molecular characterization confirms Babesia odocoilei isolated from three cervids. J Wildl Dis. 2000;36:518–30.View ArticleGoogle Scholar
  61. Ellis J, Hefford C, Baverstock PR, Dalrymple BP, Johnson AM. Ribosomal DNA sequence comparison of Babesia and Theileria. Mol Biochem Parasitol. 1992;54:87–95.View ArticleGoogle Scholar
  62. Oyamada M, Davoust B, Boni M, Dereure J, Bucheton B, Hammad A, et al. Detection of Babesia canis rossi, B. canis vogeli, and Hepatozoon canis in dogs in a village of eastern Sudan by using a screening PCR and sequencing methodologies. Clin Diagn Lab Immunol. 2005;12:1343–6.PubMedPubMed CentralGoogle Scholar
  63. McDermid KR, Snyman A, Verreynne FJ, Carroll JP, Penzhorn BL, Yabsley MJ. Surveillance for viral and parasitic pathogens in a vulnerable African lion (Panthera leo) population in the northern Tuli Game Reserve, Botswana. J Wildl Dis. 2017;53:54–61.View ArticleGoogle Scholar
  64. Sun Y, Li SG, Jiang JF, Wang X, Zhang Y, Wang H, et al. Babesia venatorum infection in child, China. Emerg Infect Dis. 2014;20:896–7.View ArticleGoogle Scholar
  65. Ciancio A, Scippa S, Finetti-Sialer M, De Candia A, Avallone B, De Vincentiis M. Redescription of Cardiosporidium cionae (Van Gaver and Stephan, 1907) (Apicomplexa: Piroplasmida), a plasmodial parasite of ascidian haemocytes. Eur J Protistol. 2008;44:181–96.View ArticleGoogle Scholar

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