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Transstadial transmission of Hepatozoon canis from larvae to nymphs of Rhipicephalus sanguineus
Veterinary Parasitology 196 (2013) 1–5
Contents lists available at SciVerse ScienceDirect
Veterinary Parasitology journal homepage: www.elsevier.com/locate/vetpar
Transstadial transmission of Hepatozoon canis from larvae to nymphs of Rhipicephalus sanguineus Alessio Giannelli a , Rafael Antonio Nascimento Ramos a , Giancarlo Di Paola a , Norbert Mencke b , Filipe Dantas-Torres a,c , Gad Baneth d , Domenico Otranto a,∗ a b c d
Dipartimento di Medicina Veterinaria, Università degli Studi di Bari, Valenzano, Bari, Italy Bayer Animal Health, Global Veterinary Services – CAP, Leverkusen, Germany Departamento de Imunologia, Centro de Pesquisas Aggeu Magalhães (Fiocruz-PE), Recife, Pernambuco, Brazil School of Veterinary Medicine, Hebrew University, Rehovot, Israel
a r t i c l e
i n f o
Article history: Received 13 December 2012 Received in revised form 19 January 2013 Accepted 17 February 2013 Keywords: Hepatozoon canis Biology Transmission Transstadial Dog Rhipicephalus sanguineus Larva Nymph
a b s t r a c t Hepatozoon canis is an apicomplexan parasite of dogs, which is known to become infected by ingesting Rhipicephalus sanguineus adult ticks. To investigate the possibility of H. canis transovarial and transstadial transmission from larvae to nymphs, engorged adult female ticks were collected from a private animal shelter in southern Italy, where H. canis infection is highly prevalent. Female ticks (n = 35) and egg batches were tested by PCR for H. canis. All eggs examined were PCR-negative whereas 88.6% of females from the environment tested positive. Additionally, fed larvae (n = 120) from a dog naturally infected by H. canis were dissected at different time points post collection (i.e. 0, 10, 20 and 30 days). Molted nymphs dissected at 20 days post collection revealed immature oocysts displaying an amorphous central structure in 50% of the specimens, and oocysts containing sporocysts with sporozoites were found in 53.3% of the nymphs dissected at 30 days post collection. This study demonstrates that H. canis is not transmitted transovarially, but it is transmitted transstadially from larvae to nymphs of R. sanguineus and develops sporozoites in oocysts that may infect dogs. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Hepatozoonosis is a cosmopolitan canine vector-borne disease transmitted by ingestion of ixodid ticks harboring the pathogen (Baneth, 2011). Two protozoan species, i.e. Hepatozoon americanum and Hepatozoon canis infect dogs. While H. americanum is restricted to the new world, H. canis presents a cosmopolitan distribution and has been reported from different regions of southern Europe, America, Africa, and Asia (Götsch et al., 2009; Abd Rani et al., 2011; Baneth, 2011; Dantas-Torres et al., 2011; Otranto et al., 2011). Dogs become infected with H. canis through the ingestion of ticks or tick parts containing mature
∗ Corresponding author. Tel.: +39 080 4679839; fax: +39 080 4679839. E-mail address: [email protected] (D. Otranto). 0304-4017/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.vetpar.2013.02.017
oocysts with infective sporozoites. H. canis sporozoites are released in the upper gastrointestinal tract, penetrate the gut wall and reach the blood or lymph circulations (Baneth et al., 2007). The complex life cycle of H. canis includes several developmental stages within the canine host (Baneth et al., 2001, 2007). H. canis meronts develop within the dog tissues and can be found in lymphatic organs (e.g., spleen and lymph nodes), the bone marrow and liver as early as 13 days post infection (Baneth et al., 2001). Micromerozoites penetrate leukocytes, usually neutrophils and monocytes, in which they mature to the gamont stage, which is detected in circulating leukocytes of parasitaemic dogs, from 28 days post infection (Baneth et al., 2007). Gamonts are ingested by ixodid tick vectors during feeding (Baneth et al., 2007). In pregnant bitches, H. canis may cross the placenta and be transmitted to the foetus (Murata et al., 1993). In ticks, gametogenesis is followed by sporogony,
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A. Giannelli et al. / Veterinary Parasitology 196 (2013) 1–5
which occurs within the tick haemocoel (Baneth et al., 2007). The life cycle in the tick and vertebrate host can be completed within about 81 days (Baneth et al., 2007). Although H. canis has been detected in several tick species (e.g., Rhipicephalus (Boophilus) microplus, Amblyomma ovale, Haemaphysalis longicornis and Haemaphysalis flava) (Murata et al., 1995; Rubini et al., 2009; de Miranda et al., 2011), the “brown dog tick” Rhipicephalus sanguineus is regarded as the most important vector worldwide (Dantas-Torres, 2008; Baneth, 2011). Following infection of the protozoa at the nymphal stage, R. sanguineus adults may transmit H. canis to the dog, soon after sporogony (Baneth et al., 2007). The role of R. sanguineus larvae as vector of H. canis has been investigated, but the transstadial transmission from larvae to the nymphal stage remains uncertain (Christophers, 1912). Conversely, transmission of H. americanum in Amblyomma maculatum ticks (from fully engorged larvae to nymphs) has been proven (Ewing et al., 2002). Therefore, the purpose of this study was to investigate two possible routes of H. canis transmission in R. sanguineus ticks: the transovarial route (from eggs laid by female ticks naturally infected by H. canis) and the transstadial route from infected larvae to the nymphal stage. 2. Materials and methods 2.1. Transovarial transmission experiment Thirty-five fully engorged female ticks were collected in June 2012 from the environment (i.e., from the ground, rocks, and walls) of a private animal shelter located in the municipality of Putignano (province of Bari, southern Italy), where a high incidence of H. canis infection was recorded in dogs (43.9%) two years before (Otranto et al., 2011). Indeed, the dogs maintained in this shelter are constantly exposed to R. sanguineus ticks, mainly during the summer (Lorusso et al., 2010; Dantas-Torres et al., 2011). The collected engorged females were identified as R. sanguineus according to their morphology (Walker et al., 2000) and placed in individual plastic vials with some holes on the top to allow air entrance. Then, the vials were placed in an incubator under controlled conditions (27 ± 1 ◦ C, RH > 70% and complete darkness). Each female was monitored daily until the end of the oviposition period. Female ticks and their egg batches (∼10 mg each) were separated for subsequent DNA extraction (see below). 2.2. Transstadial transmission experiment At the same time, 400 engorged detached larvae of R. sanguineus were collected from a dog diagnosed positive for H. canis by detection of gamonts in blood smears (parasitaemia of 80% of peripheral blood neutrophils) and by molecular methods (see below). Briefly, blood was stained with the May Grunwald Giemsa Quick Stain (Bio Optica, Milano, Italy) on glass slides, which were examined under light microscopy for the presence of H. canis gamonts for 10 min (approx. 100 fields) under 100 × oil immersion objective. The larvae were maintained in an incubator at
the same controlled conditions described above, until 120 of them were dissected as described below. 2.3. Detection of H. canis in larvae and nymphs In order to detect the presence of H. canis in the tick haemocoel (in larvae or nymphs depending on the time of dissection), 30 specimens were dissected at each follow-up time point (i.e., at the same day of collection from the dog (T1) and 10, 20 and 30 days (T2–T4) after sampling). Briefly, ticks were fixed on slides containing a drop of 0.9% NaCl solution, dissected with a sterile scalpel and then examined immediately under a light microscope (Leica® , DL MB2), at different magnifications. Oocysts were morphologically and morphometrically identified (Baneth et al., 2007) with the aid of an image analyser programme (Leica® , LAS 4.1). Finally, the 30 ticks dissected at each follow-up time point were grouped in 6 pools of 5 individuals for a total of 24 pools, and stored in phosphate buffered saline (PBS) at −20 ◦ C, until PCR analysis. Genomic DNA was extracted from ticks (i.e., engorged females, eggs, larvae and nymphs) using the guanidine isothiocyanate-phenol technique, as described by Sangioni et al. (2005), and from the dog blood sample using a commercial kit (Qiagen, DNeasy Blood & Tissue Kit, Milan, Italy), following the manufacturer’s instructions. Samples were tested by a conventional PCR for the detection of H. canis (Inokuma et al., 2002). A fragment of the 18S rRNA gene (666 base pair in size) was amplified, using primers HepF (5 -ATACATGAGCAAAATCTCAAC-3 ) and HepR (5 -CTTATTATTCCATGCTGCAG-3 ). PCR amplification was carried out in a total volume of 50 l, including ∼100 ng of genomic DNA, 10 mM TrisHCl (pH 8.3) and 50 mM KCl, 2.5 mM MgCl2 , 250 M of each dNTP, 50 pmol of each primer and 1.25 U of AmpliTaq Gold (Applied Biosystems, Foster City, CA, USA). The reactions were run in a thermal cycler (2720, Applied Biosystems, Foster City, CA, USA). Negative (no DNA template, negative reference blood samples) and positive controls (H. canis DNA from a positive tick) were included in all PCR reactions. Amplicons were resolved in ethidium bromide-stained agarose (Gellyphor, EuroClone, Milan, Italy) gels (1.5%) and sized by comparison with Gene RulerTM 100-bp DNA Ladder (MBI Fermentas, Vilnius, Lithuania) as molecular marker, and finally gels were photographed using Gel Doc 2000 (BioRad, Hercules, CA, USA). Amplicons were purified using Ultra-free-DA columns (Amicon, Millipore, Milan, Italy) and sequenced directly (Applied Biosystems, Monza, Milan, Italy) using the TaqDyeDeoxyTerminator Cycle Sequencing Kit (Applied Biosystems, Monza, Milan, Italy). Sequences were determined in both directions (using the same primers individually as for the PCR) and compared with those available in GenBank using the Basic Local Alignment Search Tool (BLAST). 3. Results H. canis DNA was amplified from 31 (88.6%) of the engorged females collected from the dog shelter. However, none of the egg batches from any of the infected female ticks were molecularly positive for H. canis.
A. Giannelli et al. / Veterinary Parasitology 196 (2013) 1–5
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Table 1 Number of ticks (engorged larvae at T1 and T2, and nymphs at T3 and T4) positive for H. canis oocysts at tick dissection. Dissection time (days after collection from a positive dog)
T1 (+0) T2 (+10) T3 (+20) T4 (+30) a
Infested ticks (%)
– – 15/30 (50.0%) 16/30 (53.3%)
H. canis oocysts
Mean
Range
Total
Mean sizes and SD (m)a
– – 5.4 7.2
– – 1–24 1–36
– – 81 115
– – 147.40 ± 46.89 × 139.10 ± 45.07 240.68 ± 66.17 × 217.44 ± 57.76
Values referred to different 15 forms measured.
The results of tick dissection (i.e., larvae and nymphs depending on the time of examination) and data on different H. canis stages found at each follow-up point are reported in Table 1. Briefly, sub-spherical and immature oocysts (Fig. 1) were observed in R. sanguineus nymphs 20 days post detachment (T3). Oocysts displayed an amorphous central structure, condensed in a plasmatic matrix as shown in Fig. 1. Many mature oocysts, with inner oval-shaped sporocysts (34.75 ± 5.6 × 19.23 ± 3.53 m) containing sporozoites (Fig. 2) were detected 30 days post detachment (T4). In two nymphs, developing oocysts as well as free sporocysts were also found, whereas free sporocysts were found in the haemolymph of 13 nymphs (Fig. 3). Twenty-three (95.8%) of the 24 pools of ticks dissected at different time points were positive for H. canis by PCR. In particular, 5/6 (83.3%) ticks pools were PCR-positive at T1 and all tick pools from T2 to T4. The sequences derived from the amplicons matched (100% identity) a GenBank H. canis reference sequence (GenBank accession number JF827277).
Fig. 2. Light micrograph of an H. canis oocyst, containing sporocysts and sporozoites, 30 days post-larval detachment.
4. Discussion This study demonstrated that H. canis is not transmitted transovarially from R. sanguineus eggs to larvae, and demonstrated, for the first time, that larvae can become infected by H. canis when feeding on a positive dog. Indeed, mature forms (oocysts containing sporozoites) develop in newly moulted nymphs, thus indicating that transstadial transmission occurs.
Fig. 1. Light micrograph of an immature H. canis oocyst from R. sanguineus nymph, 20 days post-collection.
Fig. 3. Light micrograph of broken H. canis mature oocyst and free sporocysts in a R. sanguineus nymph (30 days post-detachment).
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A previous study failed to demonstrate the completion of the protozoa sporogonic cycle in R. sanguineus larvae (Christophers, 1912) and, since then, almost all studies on the biology of this protozoa have focused on adult ticks as vectors (Baneth et al., 2001; Rubini et al., 2009). Undoubtedly, the absence of H. canis DNA in eggs laid by positive engorged females indicates that transovarial transmission does not occur in the tick vector, as previously demonstrated by microscopy without the use of more sensitive PCR (Baneth et al., 2001). However, ticks may acquire the infection as larvae during their first blood meal. In addition, the detection of developing and mature H. canis oocysts and sporozoites in ticks at 20 and 30 days postdetachment, respectively, suggests that sporogony occurs in nymphs after about 30 days post-larval detachment from an infected dog. In a previous study, immature oocysts were found in adult ticks at 27 days post-detachment (Baneth et al., 2007), and mature oocysts, featured by distinct sporocysts and sporozoites, were detected initially in unfed adults, as early as 53 days post detachment (Baneth et al., 2007). This data suggests that sporogony in R. sanguineus fed larvae may occur in a shorter time span than in nymphs. This feature well overlaps R. sanguineus moulting developmental times recorded from larvae to nymphs (8–13 days) and from nymphs to adults (11–15 days), under environmental and laboratory-controlled conditions (Dantas-Torres et al., 2011, 2012a). Based on this evidence it may be argued that a strict co-evolution occurred between the sporogony of H. canis in R. sanguineus and its development in larval and nymphal tick stages. A similar synchrony in another parasite transmitted by R. sanguineus has been recently suggested for a filarioid parasite of the genus Cercopithifilaria (Brianti et al., 2012). To further understand and substantiate this hypothesis, the metabolic mechanisms regulating H. canis maturation in tick vectors should be investigated. The finding of 81 (mean, 5.4 per tick) and 115 (7.2 per tick) oocysts in ticks dissected at T3 and T4 may suggest that a correlation exists between number and diameter of oocysts. This is possibly related to the increase in the tick body dimension. The same relationship was observed in oocysts developing in adult ticks at different time points (Baneth, 2011). The data fits well with the report on the development of H. americanum in A. maculatum, where oocysts in unfed nymphs were significantly fewer and smaller than those in adult ticks, as a likely consequence of limited space available for sporogony within the tick’s haemocoel (Ewing et al., 2002). The finding of free sporocysts external to the oocyst membrane in 81.3% of the positive nymphs at 30 days postdetachment, suggests that oocysts were mature and thus susceptible to rupture when external stimuli occurs, for instance, oral ingestion including mechanical disruption of ticks by a dog. From an epidemiological standpoint, the findings of this study suggest that the larval and nymph stages of R. sanguineus are important in maintaining hepatozoonosis in canine populations. This is especially important in certain areas such as southern Europe, where larvae and nymphs present the highest mean abundance on hosts throughout the year, when compared to the adult stage (Lorusso
et al., 2010). Infection of dogs with H. canis by nymphs might explain why the highest incidence of canine hepatozoonosis in confined dog populations from the same area was recorded during the summer (Dantas-Torres et al., 2012b), when the immature tick population reaches its peak (Lorusso et al., 2010). The present study’s findings also imply that acaricidal control measures against canine hepatozoonosis should target larvae and nymphs stages (Otranto et al., 2005) in addition to eliminating adults, as these frequently unapparent stages are also involved in H. canis transmission. Acknowledgments This study was supported by a grant from Bayer Animal Health GmbH, Leverkusen, Germany. References Abd Rani, P.A., Irwin, P.J., Coleman, G.T., Gatne, M., Traub, R.J., 2011. A survey of canine tick-borne diseases in India. Parasit. Vector 4, 141. Baneth, G., Samish, M., Shkap, V., 2007. Life cycle of Hepatozoon canis (Apicomplexa: Adeleorina: Hepatozoidae) in the tick Rhipicephalus sanguineus and domestic dog (Canis familiaris). J. Parasitol. 93, 283–299. Baneth, G., Samish, M., Alekseev, E., Aroch, I., Shkap, V., 2001. Transmission of Hepatozoon canis to dogs by naturally-fed or percutaneouslyinjected Rhipicephalus sanguineus ticks. J. Parasitol. 87, 606–611. Baneth, G., 2011. Perspectives on canine and feline hepatozoonosis. Vet. Parasitol. 181, 3–11. Brianti, E., Otranto, D., Dantas-Torres, F., Weigl, S., Latrofa, M.S., Gaglio, G., Napoli, E., Brucato, G., Cauquil, L., Giannetto, S., Bain, O., 2012. Rhipicephalus sanguineus (Ixodida, Ixodidae) as intermediate host of a canine neglected filarial species with dermal microfilariae. Vet. Parasitol. 10, 330–337. Christophers, S.R., 1912. The development of Leucocytozoon canis in the tick with a reference to the development of Piroplasma. Parasitology 5, 37–48. Dantas-Torres, F., 2008. The brown dog tick, Rhipicephalus sanguineus (Latreille, 1806) (Acari: Ixodidae): from taxonomy to control. Vet. Parasitol. 152, 173–185. Dantas-Torres, F., Figueredo, L.A., Otranto, D., 2011. Seasonal variation in the effect of climate on the biology of Rhipicephalus sanguineus in southern Europe. Parasitology 138, 527–536. Dantas-Torres, F., Giannelli, A., Otranto, D., 2012a. Starvation and overwinter do not affect the reproductive fitness of Rhipicephalus sanguineus. Vet. Parasitol. 185, 260–264. Dantas-Torres, F., Latrofa, M.S., Weigl, S., Tarallo, V.D., Lia, R.P., Otranto, D., 2012b. Hepatozoon canis infection in ticks during spring and summer in Italy. Parasitol. Res. 110, 695–698. de Miranda, R.L., de Castro, J.R., Olegário, M.M., Beletti, M.E., Mundim, A.V., O’Dwyer, L.H., Eyal, O., Talmi-Frank, D., Cury, M.C., Baneth, G., 2011. Oocysts of Hepatozoon canis in Rhipicephalus (Boophilus) microplus collected from a naturally infected dog. Vet Parasitol. 177, 392–396. Ewing, S.A., Du Bois, J.G., Mathew, J.S., Panciera, R.J., 2002. Larval Gulf Coast ticks (Amblyomma maculatum) (Acari: Ixodidae) as host for Hepatozoon americanum (Apicomplexa: Adeleorina). Vet. Parasitol. 103, 43–51. Götsch, S., Leschnik, M., Duscher, G., Burgstaller, J.P., Wille-Piazzai, W., Joachim, A., 2009. Ticks and haemoparasites of dogs from Praia, Cape Verde. Vet. Parasitol. 166, 171–174. Inokuma, H., Okuda, M., Ohno, K., Shimoda, K., Onishi, T., 2002. Analysis of the 18S rRNA gene sequence of a Hepatozoon detected in two Japanese dogs. Vet. Parasitol. 106, 265–271. Lorusso, V., Dantas-Torres, F., Lia, R.P., Tarallo, V.D., Mencke, N., Capelli, G., Otranto, D., 2010. Seasonal dynamics of the brown dog tick, Rhipicephalus sanguineus, on a confined dog population in Italy. Med. Vet. Entomol. 24, 309–315. Murata, T., Inoue, M., Tateyama, S., Taura, Y., Nakama, S., 1993. Vertical transmission of Hepatozoon canis in dogs. J. Vet. Med. Sci. 55, 867–868. Murata, T., Inoue, M., Taura, Y., Nakama, S., Abe, H., Fujisaki, K., 1995. Detection of Hepatozoon canis oocyst from ticks collected from the infected dogs. J. Vet. Med. Sci. 57, 111–112.
A. Giannelli et al. / Veterinary Parasitology 196 (2013) 1–5 Otranto, D., Lia, R.P., Cantacessi, C., Galli, G., Paradies, P., Mallia, E., Capelli, G., 2005. Efficacy of a combination of imidacloprid 10%/permethrin 50% versus fipronil 10%/(S)-methoprene 12%, against ticks in naturally infected dogs. Vet. Parasitol. 130, 293–304. Otranto, D., Dantas-Torres, F., Weigl, S., Latrofa, M.S., Stanneck, D., Decaprariis, D., Capelli, G., Baneth, G., 2011. Diagnosis of Hepatozoon canis in young dogs by cytology and PCR. Parasit. Vector 4, 55. Rubini, A.S., Paduan, K.S., Martins, T.F., Labruna, M.B., O’Dwyer, L.H., 2009. Acquisition and transmission of Hepatozoon canis
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(Apicomplexa: Hepatozoidae) by the tick Amblyomma ovale (Acari: Ixodidae). Vet. Parasitol. 164, 324–327. Sangioni, L.A., Horta, M.C., Vianna, M.C., Gennari, S.M., Soares, R.M., Galva˜o, M.A., Schumaker, T.T., Ferreira, F., Vidotto, O., Labruna, M.B., 2005. Rickettsial infection in animals and Brazilian spotted fever endemicity. Emerg. Infect. Dis. 11, 265–270. Walker, J.B., Keirans, J.E., Horak, I.G., 2000. The Genus Rhipicephalus (Acari, Ixodidae): A Guide to the Brown Ticks of the World. Cambridge University Press, Cambridge, UK, 655 pp.
Contents lists available at SciVerse ScienceDirect
Veterinary Parasitology journal homepage: www.elsevier.com/locate/vetpar
Transstadial transmission of Hepatozoon canis from larvae to nymphs of Rhipicephalus sanguineus Alessio Giannelli a , Rafael Antonio Nascimento Ramos a , Giancarlo Di Paola a , Norbert Mencke b , Filipe Dantas-Torres a,c , Gad Baneth d , Domenico Otranto a,∗ a b c d
Dipartimento di Medicina Veterinaria, Università degli Studi di Bari, Valenzano, Bari, Italy Bayer Animal Health, Global Veterinary Services – CAP, Leverkusen, Germany Departamento de Imunologia, Centro de Pesquisas Aggeu Magalhães (Fiocruz-PE), Recife, Pernambuco, Brazil School of Veterinary Medicine, Hebrew University, Rehovot, Israel
a r t i c l e
i n f o
Article history: Received 13 December 2012 Received in revised form 19 January 2013 Accepted 17 February 2013 Keywords: Hepatozoon canis Biology Transmission Transstadial Dog Rhipicephalus sanguineus Larva Nymph
a b s t r a c t Hepatozoon canis is an apicomplexan parasite of dogs, which is known to become infected by ingesting Rhipicephalus sanguineus adult ticks. To investigate the possibility of H. canis transovarial and transstadial transmission from larvae to nymphs, engorged adult female ticks were collected from a private animal shelter in southern Italy, where H. canis infection is highly prevalent. Female ticks (n = 35) and egg batches were tested by PCR for H. canis. All eggs examined were PCR-negative whereas 88.6% of females from the environment tested positive. Additionally, fed larvae (n = 120) from a dog naturally infected by H. canis were dissected at different time points post collection (i.e. 0, 10, 20 and 30 days). Molted nymphs dissected at 20 days post collection revealed immature oocysts displaying an amorphous central structure in 50% of the specimens, and oocysts containing sporocysts with sporozoites were found in 53.3% of the nymphs dissected at 30 days post collection. This study demonstrates that H. canis is not transmitted transovarially, but it is transmitted transstadially from larvae to nymphs of R. sanguineus and develops sporozoites in oocysts that may infect dogs. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Hepatozoonosis is a cosmopolitan canine vector-borne disease transmitted by ingestion of ixodid ticks harboring the pathogen (Baneth, 2011). Two protozoan species, i.e. Hepatozoon americanum and Hepatozoon canis infect dogs. While H. americanum is restricted to the new world, H. canis presents a cosmopolitan distribution and has been reported from different regions of southern Europe, America, Africa, and Asia (Götsch et al., 2009; Abd Rani et al., 2011; Baneth, 2011; Dantas-Torres et al., 2011; Otranto et al., 2011). Dogs become infected with H. canis through the ingestion of ticks or tick parts containing mature
∗ Corresponding author. Tel.: +39 080 4679839; fax: +39 080 4679839. E-mail address: [email protected] (D. Otranto). 0304-4017/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.vetpar.2013.02.017
oocysts with infective sporozoites. H. canis sporozoites are released in the upper gastrointestinal tract, penetrate the gut wall and reach the blood or lymph circulations (Baneth et al., 2007). The complex life cycle of H. canis includes several developmental stages within the canine host (Baneth et al., 2001, 2007). H. canis meronts develop within the dog tissues and can be found in lymphatic organs (e.g., spleen and lymph nodes), the bone marrow and liver as early as 13 days post infection (Baneth et al., 2001). Micromerozoites penetrate leukocytes, usually neutrophils and monocytes, in which they mature to the gamont stage, which is detected in circulating leukocytes of parasitaemic dogs, from 28 days post infection (Baneth et al., 2007). Gamonts are ingested by ixodid tick vectors during feeding (Baneth et al., 2007). In pregnant bitches, H. canis may cross the placenta and be transmitted to the foetus (Murata et al., 1993). In ticks, gametogenesis is followed by sporogony,
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A. Giannelli et al. / Veterinary Parasitology 196 (2013) 1–5
which occurs within the tick haemocoel (Baneth et al., 2007). The life cycle in the tick and vertebrate host can be completed within about 81 days (Baneth et al., 2007). Although H. canis has been detected in several tick species (e.g., Rhipicephalus (Boophilus) microplus, Amblyomma ovale, Haemaphysalis longicornis and Haemaphysalis flava) (Murata et al., 1995; Rubini et al., 2009; de Miranda et al., 2011), the “brown dog tick” Rhipicephalus sanguineus is regarded as the most important vector worldwide (Dantas-Torres, 2008; Baneth, 2011). Following infection of the protozoa at the nymphal stage, R. sanguineus adults may transmit H. canis to the dog, soon after sporogony (Baneth et al., 2007). The role of R. sanguineus larvae as vector of H. canis has been investigated, but the transstadial transmission from larvae to the nymphal stage remains uncertain (Christophers, 1912). Conversely, transmission of H. americanum in Amblyomma maculatum ticks (from fully engorged larvae to nymphs) has been proven (Ewing et al., 2002). Therefore, the purpose of this study was to investigate two possible routes of H. canis transmission in R. sanguineus ticks: the transovarial route (from eggs laid by female ticks naturally infected by H. canis) and the transstadial route from infected larvae to the nymphal stage. 2. Materials and methods 2.1. Transovarial transmission experiment Thirty-five fully engorged female ticks were collected in June 2012 from the environment (i.e., from the ground, rocks, and walls) of a private animal shelter located in the municipality of Putignano (province of Bari, southern Italy), where a high incidence of H. canis infection was recorded in dogs (43.9%) two years before (Otranto et al., 2011). Indeed, the dogs maintained in this shelter are constantly exposed to R. sanguineus ticks, mainly during the summer (Lorusso et al., 2010; Dantas-Torres et al., 2011). The collected engorged females were identified as R. sanguineus according to their morphology (Walker et al., 2000) and placed in individual plastic vials with some holes on the top to allow air entrance. Then, the vials were placed in an incubator under controlled conditions (27 ± 1 ◦ C, RH > 70% and complete darkness). Each female was monitored daily until the end of the oviposition period. Female ticks and their egg batches (∼10 mg each) were separated for subsequent DNA extraction (see below). 2.2. Transstadial transmission experiment At the same time, 400 engorged detached larvae of R. sanguineus were collected from a dog diagnosed positive for H. canis by detection of gamonts in blood smears (parasitaemia of 80% of peripheral blood neutrophils) and by molecular methods (see below). Briefly, blood was stained with the May Grunwald Giemsa Quick Stain (Bio Optica, Milano, Italy) on glass slides, which were examined under light microscopy for the presence of H. canis gamonts for 10 min (approx. 100 fields) under 100 × oil immersion objective. The larvae were maintained in an incubator at
the same controlled conditions described above, until 120 of them were dissected as described below. 2.3. Detection of H. canis in larvae and nymphs In order to detect the presence of H. canis in the tick haemocoel (in larvae or nymphs depending on the time of dissection), 30 specimens were dissected at each follow-up time point (i.e., at the same day of collection from the dog (T1) and 10, 20 and 30 days (T2–T4) after sampling). Briefly, ticks were fixed on slides containing a drop of 0.9% NaCl solution, dissected with a sterile scalpel and then examined immediately under a light microscope (Leica® , DL MB2), at different magnifications. Oocysts were morphologically and morphometrically identified (Baneth et al., 2007) with the aid of an image analyser programme (Leica® , LAS 4.1). Finally, the 30 ticks dissected at each follow-up time point were grouped in 6 pools of 5 individuals for a total of 24 pools, and stored in phosphate buffered saline (PBS) at −20 ◦ C, until PCR analysis. Genomic DNA was extracted from ticks (i.e., engorged females, eggs, larvae and nymphs) using the guanidine isothiocyanate-phenol technique, as described by Sangioni et al. (2005), and from the dog blood sample using a commercial kit (Qiagen, DNeasy Blood & Tissue Kit, Milan, Italy), following the manufacturer’s instructions. Samples were tested by a conventional PCR for the detection of H. canis (Inokuma et al., 2002). A fragment of the 18S rRNA gene (666 base pair in size) was amplified, using primers HepF (5 -ATACATGAGCAAAATCTCAAC-3 ) and HepR (5 -CTTATTATTCCATGCTGCAG-3 ). PCR amplification was carried out in a total volume of 50 l, including ∼100 ng of genomic DNA, 10 mM TrisHCl (pH 8.3) and 50 mM KCl, 2.5 mM MgCl2 , 250 M of each dNTP, 50 pmol of each primer and 1.25 U of AmpliTaq Gold (Applied Biosystems, Foster City, CA, USA). The reactions were run in a thermal cycler (2720, Applied Biosystems, Foster City, CA, USA). Negative (no DNA template, negative reference blood samples) and positive controls (H. canis DNA from a positive tick) were included in all PCR reactions. Amplicons were resolved in ethidium bromide-stained agarose (Gellyphor, EuroClone, Milan, Italy) gels (1.5%) and sized by comparison with Gene RulerTM 100-bp DNA Ladder (MBI Fermentas, Vilnius, Lithuania) as molecular marker, and finally gels were photographed using Gel Doc 2000 (BioRad, Hercules, CA, USA). Amplicons were purified using Ultra-free-DA columns (Amicon, Millipore, Milan, Italy) and sequenced directly (Applied Biosystems, Monza, Milan, Italy) using the TaqDyeDeoxyTerminator Cycle Sequencing Kit (Applied Biosystems, Monza, Milan, Italy). Sequences were determined in both directions (using the same primers individually as for the PCR) and compared with those available in GenBank using the Basic Local Alignment Search Tool (BLAST). 3. Results H. canis DNA was amplified from 31 (88.6%) of the engorged females collected from the dog shelter. However, none of the egg batches from any of the infected female ticks were molecularly positive for H. canis.
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Table 1 Number of ticks (engorged larvae at T1 and T2, and nymphs at T3 and T4) positive for H. canis oocysts at tick dissection. Dissection time (days after collection from a positive dog)
T1 (+0) T2 (+10) T3 (+20) T4 (+30) a
Infested ticks (%)
– – 15/30 (50.0%) 16/30 (53.3%)
H. canis oocysts
Mean
Range
Total
Mean sizes and SD (m)a
– – 5.4 7.2
– – 1–24 1–36
– – 81 115
– – 147.40 ± 46.89 × 139.10 ± 45.07 240.68 ± 66.17 × 217.44 ± 57.76
Values referred to different 15 forms measured.
The results of tick dissection (i.e., larvae and nymphs depending on the time of examination) and data on different H. canis stages found at each follow-up point are reported in Table 1. Briefly, sub-spherical and immature oocysts (Fig. 1) were observed in R. sanguineus nymphs 20 days post detachment (T3). Oocysts displayed an amorphous central structure, condensed in a plasmatic matrix as shown in Fig. 1. Many mature oocysts, with inner oval-shaped sporocysts (34.75 ± 5.6 × 19.23 ± 3.53 m) containing sporozoites (Fig. 2) were detected 30 days post detachment (T4). In two nymphs, developing oocysts as well as free sporocysts were also found, whereas free sporocysts were found in the haemolymph of 13 nymphs (Fig. 3). Twenty-three (95.8%) of the 24 pools of ticks dissected at different time points were positive for H. canis by PCR. In particular, 5/6 (83.3%) ticks pools were PCR-positive at T1 and all tick pools from T2 to T4. The sequences derived from the amplicons matched (100% identity) a GenBank H. canis reference sequence (GenBank accession number JF827277).
Fig. 2. Light micrograph of an H. canis oocyst, containing sporocysts and sporozoites, 30 days post-larval detachment.
4. Discussion This study demonstrated that H. canis is not transmitted transovarially from R. sanguineus eggs to larvae, and demonstrated, for the first time, that larvae can become infected by H. canis when feeding on a positive dog. Indeed, mature forms (oocysts containing sporozoites) develop in newly moulted nymphs, thus indicating that transstadial transmission occurs.
Fig. 1. Light micrograph of an immature H. canis oocyst from R. sanguineus nymph, 20 days post-collection.
Fig. 3. Light micrograph of broken H. canis mature oocyst and free sporocysts in a R. sanguineus nymph (30 days post-detachment).
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A previous study failed to demonstrate the completion of the protozoa sporogonic cycle in R. sanguineus larvae (Christophers, 1912) and, since then, almost all studies on the biology of this protozoa have focused on adult ticks as vectors (Baneth et al., 2001; Rubini et al., 2009). Undoubtedly, the absence of H. canis DNA in eggs laid by positive engorged females indicates that transovarial transmission does not occur in the tick vector, as previously demonstrated by microscopy without the use of more sensitive PCR (Baneth et al., 2001). However, ticks may acquire the infection as larvae during their first blood meal. In addition, the detection of developing and mature H. canis oocysts and sporozoites in ticks at 20 and 30 days postdetachment, respectively, suggests that sporogony occurs in nymphs after about 30 days post-larval detachment from an infected dog. In a previous study, immature oocysts were found in adult ticks at 27 days post-detachment (Baneth et al., 2007), and mature oocysts, featured by distinct sporocysts and sporozoites, were detected initially in unfed adults, as early as 53 days post detachment (Baneth et al., 2007). This data suggests that sporogony in R. sanguineus fed larvae may occur in a shorter time span than in nymphs. This feature well overlaps R. sanguineus moulting developmental times recorded from larvae to nymphs (8–13 days) and from nymphs to adults (11–15 days), under environmental and laboratory-controlled conditions (Dantas-Torres et al., 2011, 2012a). Based on this evidence it may be argued that a strict co-evolution occurred between the sporogony of H. canis in R. sanguineus and its development in larval and nymphal tick stages. A similar synchrony in another parasite transmitted by R. sanguineus has been recently suggested for a filarioid parasite of the genus Cercopithifilaria (Brianti et al., 2012). To further understand and substantiate this hypothesis, the metabolic mechanisms regulating H. canis maturation in tick vectors should be investigated. The finding of 81 (mean, 5.4 per tick) and 115 (7.2 per tick) oocysts in ticks dissected at T3 and T4 may suggest that a correlation exists between number and diameter of oocysts. This is possibly related to the increase in the tick body dimension. The same relationship was observed in oocysts developing in adult ticks at different time points (Baneth, 2011). The data fits well with the report on the development of H. americanum in A. maculatum, where oocysts in unfed nymphs were significantly fewer and smaller than those in adult ticks, as a likely consequence of limited space available for sporogony within the tick’s haemocoel (Ewing et al., 2002). The finding of free sporocysts external to the oocyst membrane in 81.3% of the positive nymphs at 30 days postdetachment, suggests that oocysts were mature and thus susceptible to rupture when external stimuli occurs, for instance, oral ingestion including mechanical disruption of ticks by a dog. From an epidemiological standpoint, the findings of this study suggest that the larval and nymph stages of R. sanguineus are important in maintaining hepatozoonosis in canine populations. This is especially important in certain areas such as southern Europe, where larvae and nymphs present the highest mean abundance on hosts throughout the year, when compared to the adult stage (Lorusso
et al., 2010). Infection of dogs with H. canis by nymphs might explain why the highest incidence of canine hepatozoonosis in confined dog populations from the same area was recorded during the summer (Dantas-Torres et al., 2012b), when the immature tick population reaches its peak (Lorusso et al., 2010). The present study’s findings also imply that acaricidal control measures against canine hepatozoonosis should target larvae and nymphs stages (Otranto et al., 2005) in addition to eliminating adults, as these frequently unapparent stages are also involved in H. canis transmission. Acknowledgments This study was supported by a grant from Bayer Animal Health GmbH, Leverkusen, Germany. References Abd Rani, P.A., Irwin, P.J., Coleman, G.T., Gatne, M., Traub, R.J., 2011. A survey of canine tick-borne diseases in India. Parasit. Vector 4, 141. Baneth, G., Samish, M., Shkap, V., 2007. Life cycle of Hepatozoon canis (Apicomplexa: Adeleorina: Hepatozoidae) in the tick Rhipicephalus sanguineus and domestic dog (Canis familiaris). J. Parasitol. 93, 283–299. Baneth, G., Samish, M., Alekseev, E., Aroch, I., Shkap, V., 2001. Transmission of Hepatozoon canis to dogs by naturally-fed or percutaneouslyinjected Rhipicephalus sanguineus ticks. J. Parasitol. 87, 606–611. Baneth, G., 2011. Perspectives on canine and feline hepatozoonosis. Vet. Parasitol. 181, 3–11. Brianti, E., Otranto, D., Dantas-Torres, F., Weigl, S., Latrofa, M.S., Gaglio, G., Napoli, E., Brucato, G., Cauquil, L., Giannetto, S., Bain, O., 2012. Rhipicephalus sanguineus (Ixodida, Ixodidae) as intermediate host of a canine neglected filarial species with dermal microfilariae. Vet. Parasitol. 10, 330–337. Christophers, S.R., 1912. The development of Leucocytozoon canis in the tick with a reference to the development of Piroplasma. Parasitology 5, 37–48. Dantas-Torres, F., 2008. The brown dog tick, Rhipicephalus sanguineus (Latreille, 1806) (Acari: Ixodidae): from taxonomy to control. Vet. Parasitol. 152, 173–185. Dantas-Torres, F., Figueredo, L.A., Otranto, D., 2011. Seasonal variation in the effect of climate on the biology of Rhipicephalus sanguineus in southern Europe. Parasitology 138, 527–536. Dantas-Torres, F., Giannelli, A., Otranto, D., 2012a. Starvation and overwinter do not affect the reproductive fitness of Rhipicephalus sanguineus. Vet. Parasitol. 185, 260–264. Dantas-Torres, F., Latrofa, M.S., Weigl, S., Tarallo, V.D., Lia, R.P., Otranto, D., 2012b. Hepatozoon canis infection in ticks during spring and summer in Italy. Parasitol. Res. 110, 695–698. de Miranda, R.L., de Castro, J.R., Olegário, M.M., Beletti, M.E., Mundim, A.V., O’Dwyer, L.H., Eyal, O., Talmi-Frank, D., Cury, M.C., Baneth, G., 2011. Oocysts of Hepatozoon canis in Rhipicephalus (Boophilus) microplus collected from a naturally infected dog. Vet Parasitol. 177, 392–396. Ewing, S.A., Du Bois, J.G., Mathew, J.S., Panciera, R.J., 2002. Larval Gulf Coast ticks (Amblyomma maculatum) (Acari: Ixodidae) as host for Hepatozoon americanum (Apicomplexa: Adeleorina). Vet. Parasitol. 103, 43–51. Götsch, S., Leschnik, M., Duscher, G., Burgstaller, J.P., Wille-Piazzai, W., Joachim, A., 2009. Ticks and haemoparasites of dogs from Praia, Cape Verde. Vet. Parasitol. 166, 171–174. Inokuma, H., Okuda, M., Ohno, K., Shimoda, K., Onishi, T., 2002. Analysis of the 18S rRNA gene sequence of a Hepatozoon detected in two Japanese dogs. Vet. Parasitol. 106, 265–271. Lorusso, V., Dantas-Torres, F., Lia, R.P., Tarallo, V.D., Mencke, N., Capelli, G., Otranto, D., 2010. Seasonal dynamics of the brown dog tick, Rhipicephalus sanguineus, on a confined dog population in Italy. Med. Vet. Entomol. 24, 309–315. Murata, T., Inoue, M., Tateyama, S., Taura, Y., Nakama, S., 1993. Vertical transmission of Hepatozoon canis in dogs. J. Vet. Med. Sci. 55, 867–868. Murata, T., Inoue, M., Taura, Y., Nakama, S., Abe, H., Fujisaki, K., 1995. Detection of Hepatozoon canis oocyst from ticks collected from the infected dogs. J. Vet. Med. Sci. 57, 111–112.
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