Seasonal dynamics, spatial distribution and genetic analysis of Anaplasma species infecting small ruminants from Northern Tunisia

Accepted Manuscript Seasonal dynamics, spatial distribution and genetic analysis of Anaplasma species infecting small ruminants from Northern Tunisia

Hanène Belkahia, Mourad Ben Said, Narjesse El Mabrouk, Mariem Saidani, Chayma Cherni, Mariem Ben Hassen, Ali Bouattour, Lilia Messadi PII: DOI: Reference:

S1567-1348(17)30203-4 doi: 10.1016/j.meegid.2017.06.016 MEEGID 3178

To appear in:

Infection, Genetics and Evolution

Received date: Revised date: Accepted date:

28 March 2017 16 May 2017 15 June 2017

Please cite this article as: Hanène Belkahia, Mourad Ben Said, Narjesse El Mabrouk, Mariem Saidani, Chayma Cherni, Mariem Ben Hassen, Ali Bouattour, Lilia Messadi , Seasonal dynamics, spatial distribution and genetic analysis of Anaplasma species infecting small ruminants from Northern Tunisia, Infection, Genetics and Evolution (2017), doi: 10.1016/j.meegid.2017.06.016

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Seasonal dynamics, spatial distribution and genetic analysis of Anaplasma species infecting small ruminants from Northern Tunisia. Authors: Hanène Belkahia 1, Mourad Ben Said 1, Narjesse El Mabrouk 1, Mariem Saidani 1, Chayma Cherni 1, Mariem Ben Hassen 1, Ali Bouattour 2, Lilia Messadi 1,* 1

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Service de Microbiologie et Immunologie, Ecole Nationale de Médecine Vétérinaire, Université de La Manouba, 2020 Sidi Thabet, Tunisie 2 Service d’Entomologie Médicale, Institut Pasteur de Tunis, Université Tunis El Manar, 1002 Tunis, Tunisie *

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Corresponding author: Pr. Lilia Messadi Service de Microbiologie et Immunologie, Ecole Nationale de Médecine Vétérinaire, 2020 Sidi Thabet, Tunisie Tel: +216 71 552 200 Fax: +216 71 552 441 E-mail: [email protected]

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Seasonal dynamics, spatial distribution and genetic analysis of Anaplasma species infecting small ruminants from Northern Tunisia.

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Abstract To date, there have been no reports on seasonal variations of Anaplasma spp. in South Mediterranean small ruminants. In this longitudinal field study, single and mixed Anaplasma spp. infections in small ruminants from five different governorates belonging to three bioclimatic zones from the North of Tunisia were evaluated according to seasons. A total of 1685 blood small ruminant samples were collected in spring (355 sheep and 241 goats), summer (249 sheep and 202 goats), autumn (236 sheep and 186 goats) and winter (132 sheep and 84 goats). Molecular survey of A. ovis and A. bovis showed that average prevalence rates were 35.6% (minimum 30.7% in spring and maximum 43.6% in autumn) and 7.4% (minimum 0.9% in spring and maximum 18.1% in summer), respectively, in sheep, and 46% (minimum 21.7% in summer and maximum 65.5% in winter) and 10.1% (minimum 2.2% in autumn and maximum 23.8% in summer), respectively, in goats. A. phagocytophilum was not detected in all investigated animals. The infection profiles of A. ovis and A. bovis show that anaplasmosis caused by A. ovis is endemic in small ruminants from all investigated bioclimatic areas during the four seasons but conversely, A. bovis infection is highly intensified only in the summer. A. ovis and A. bovis infections were validated by sequencing. The comparison of the 16S rRNA sequences of A. bovis variants showed 100% identity between Tunisian variants isolated from goats, sheep and cattle. The analysis of A. ovis msp4 sequences revealed two different genetic variants previously described in Italy. This is the first survey outlining seasonal dynamics of Anaplasma spp. infections in Tunisian small ruminants. This situation should to be taken into account if anaplasmosis control programs in these domesticated animals are envisaged.

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Keywords: Anaplasma spp.; Co-infection; Seasonal variation; Spatial distribution; Molecular characterization; Tunisian small ruminants

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1. Introduction The classified Anaplasma species that impact small ruminants’ health are A. ovis, A. phagocytophilum and A. bovis. Among these, A. ovis is an inter-erythrocytic pathogen responsible for ovine anaplasmosis in tropical and subtropical regions (Friedhoff, 1997). In small ruminants, infection is usually subclinical; occasionally, it can be severe with haemolytic anaemia, haemoglobinuria and fever (Hornok et al., 2007). A. phagocytophilum infects neutrophil granulocytes of humans, and wild and domestic animals (Dumler et al., 2001). In ruminants, it causes tick-borne fever (TBF) (Stuen, 2007; Woldehiwet, 2010). TBF most common symptoms include high fever, anorexia, dullness and milk yield decrease (Tuomi, 1967; Woldehiwet, 2010). Within the genus Anaplasma, Anaplasma bovis, a monocytotropic bacterial species, has been detected, for the first time, in goats by Liu et al. (2012) and in sheep by Ben Said et al. (2015b). In cattle, A. bovis infection has been reported as usually asymptomatic, but it can cause a variety of clinical symptoms, including reduced body weight, fever, anemia, depression, lymphadenopathy and rarely abortion, and death in some cases (Noaman and Shayan, 2010). In Tunisia, several Anaplasma spp. were identified in small ruminants such as A. ovis in sheep from northern and central Tunisia (Belkahia et al., 2014), and A. ovis, A. bovis, and strains closely related to A. phagocytophilum and A. platys in goats and sheep located in the North of the country (Ben Said et al., 2015a,b and 2017a,b). In these cross-sectional studies, mixed infections by two, three and even four classified and unclassified Anaplasma spp. have been found in these animals and the study of the interactions between these infections is important because it may have an effect on the disease severity that aggravate animal condition, occasionally leading to death (Kocan et al., 2004). In addition, acute illnesses are not only described as being associated with co-infections but also with other reasons such as climate change, nonconforming farming practices, habitat suitability for vectors essentially ticks, animal movement and/or abusive vaccination (Friedhoff, 1997; Kocan et al., 2004; Torina et al., 2008a; Ogden and Lindsay, 2016). The impact of some risk factors in Anaplasma spp. infections was evaluated in Tunisian small ruminants. Indeed, A. ovis and A. bovis prevalence rates in sheep and goats varied according to geographic location, bioclimatic area, age, animal breed and/or tick infestation (Belkahia et al., 2014; Ben Said et al., 2015a, b). However, seasonal dynamics evaluation in infection epidemiology is crucial for estimating endemicity of anaplasmosis and its potential for spread. To date, there is no data concerning the seasonal fluctuations of single and mixed Anaplasma spp. infections in Tunisian small ruminants. Thus, this longitudinal survey aimed to provide spatiotemporal information necessary to evaluate the potentially impact of ovine anaplasmosis in Tunisia. From where, molecular epidemiology of Anaplasma spp. infecting small ruminants was investigated according to seasons by single and nested PCR confirmed by msp4 and 16S rRNA sequence analyses. 2. Materials and methods 2.1. Blood sampling and DNA extraction A longitudinal study was carried out in thirty delegations of Northern Tunisia belonging to five governorates (Tunis, Ariana, Bizerte, Beja and Nabeul) and three bioclimatic areas (lower humid, sub-humid and higher semi-arid) (Supplementary files S1 to S9). According to seasons, a total of 1685 blood small ruminant samples were collected in spring (from March to May 2014; 355 sheep and 241 goats), summer (from June to August 2014; 249 sheep and 202 goats), autumn (from September to November 2014; 236 sheep and 186 goats) and winter (from December 2014 to February 2015; 132 sheep and 84 goats) (Supplementary files S2 to S9). DNA was extracted from 300 μl of each blood sample with Wizard® Genomic DNA purification kit (Promega, Madison, USA) according to manufacturer instructions. 3

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2.2. Anaplasma species detection Primer pair EE1 and EE2 was used in a first PCR for amplifying the 16S rRNA gene of all Anaplasma and Ehrlichia species potentially present in ruminants (Barlough et al., 1996; Ben Said et al., 2017b) (Table 1). PCR reaction was performed in a final volume of 50 µl containing 0,125 U/µl Taq DNA polymerase (Biobasic Inc, Canada), 1x PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs, 50 to 150 ng of genomic DNA, 0.5 µM of primers. Thermal cycling profile was as described by Liu et al. (2012). A. bovis specific primers were used in nested PCR for strains detection and characterization (Table 1). One microliter of the first PCR product was used as target DNA in the nested PCRs, thermally profiled as in Kawahara et al. (2006). Positive 16S rRNA samples by outer primers EE1 and EE2 were tested by hemi-nested PCR, for the specific detection of A. phagocytophilum, using outer primers EphplgroEL-F and EphplgroEL-R, and inner primers EphplgroEL-F and EphgroEL-R amplifying 573 bp sequence of the groel gene (Alberti et al., 2005, Table 1). A. ovis infection was detected by single PCR with the AovisMSP4Fw and AovisMSP4Rev specific primers designed by Torina et al. (2012, Table 1). Distilled water and DNA extracted from A. ovis, A. bovis and A. phagocytophilum, were used as negative and positive controls, respectively. PCR products were electrophoresed in 1% agarose gel and sized with 1 Kb Plus DNA Ladder (Promega, Madison, WI, USA). For msp4 genotyping, selected ovine and caprine samples positive to A. ovis by single PCR based on AovisMSP4Fw and AovisMSP4Rev (Torina et al., 2012) were used in a traditional PCR with MSP45 and MSP43 primers (de la Fuente et al., 2005) (Table 1). Shortly, reactions contained 1x PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs and 0.125 U/µl Taq DNA Polymerase (Biobasic Inc, Canada), 2 µl (50-150 ng) DNA, 0.5 µM of the primers, and milliQ sterile water to a total volume of 50 µl. In each experiment distilled water and DNA extracted from A. ovis (Ben Said et al., 2015a) were used as negative and positive controls, respectively. Thermal cycling reactions were performed as described by de la Fuente et al. (2005). PCR products were electrophoresed in 1.5% agarose gel.

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2.3. DNA sequencing and genetic analysis Selected positive PCR products from primers AB1f/AB1r and MSP45/MSP43 of A. bovis and A. ovis, respectively, were purified with the GF-1 Ambi Clean kit (Vivantis, USA) according to manufacturer instructions. Purified DNA fragments were sequenced in both directions, using the same primers as for the PCR amplifications (Table 1). The reactions were performed using a conventional Big Dye Terminator cycle sequencing ready reaction kit (Perkin Elmer, Applied Biosystems, Foster City, USA) and an ABI3730XL automated DNA sequencer. The chromatograms were evaluated with Chromas Lite v 2.01. The DNAMAN software (Version 5.2.2; Lynnon Biosoft, Que., Canada) was used to perform multiple sequence alignment of 16S rRNA and msp4 sequences and to translate nucleotide to amino acid msp4 sequences. BLAST analysis of GenBank was used to assess the level of similarity with previously reported sequences (http://blast.ncbi.nlm.nih.gov/, Altschul et al., 1997). 2.4. GenBank accession numbers The 16S rRNA partial sequences of A. bovis Ab1 to Ab12 isolates have been deposited under GenBank accession numbers KY655797 to KY655808. The msp4 partial sequences of A. ovis Ao1 to Ao5 isolates have been deposited in the GenBank under accession numbers KY659320 to KY659324 (Table 2). 2.5. Statistical analysis Exact confidence intervals (CI) for prevalence rates at the 95% level were calculated. Comparison of the prevalence of Anaplasma species in sheep and goats according to seasons, bioclimatic zones, 4

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3. Results 3.1. Spatial distribution and seasonal variation of Anaplasma infections 3.1.1. In goats The infection rate of A. ovis in goats begins relatively high (55.2%) during the spring, then decreases significantly during the summer (21.7%), increases during the autumn (41.4%) and reaches its maximum during the winter season (65.5%) (P < 0.001; Figure 1A). Goats located in lower humid and sub-humid areas were statistically more infected by A. ovis, respectively, in spring (74.7%) and winter (100%) than in other seasons (P < 0.001, Figure 2A). In higher semi-arid area, A. ovis infection rates were relatively constant during the four seasons with a slight increase of infection rate in the winter (46.7%; Figure 2A). A. bovis infection and A. ovis/A. bovis co-infection profiles are similar and the rates begin relatively moderate (10.8 %) during the spring, then increase significantly (P < 0.001) and reach their maximum during the summer (21.7 and 21.3%, respectively) and, finally, decrease during the autumn (2.2%) and the winter seasons (3.6%) (Figure 1A). In lower humid, subhumid and higher semi-arid areas, goats analyzed in summer were statistically the most infected by A. bovis than those analyzed in other seasons with prevalence rates estimated at 33.9, 20.9 and 18.6%, respectively (P < 0.001; Figure 3A). As shown in Figure 4A, A. ovis/A. bovis co-infection profiles in the three analyzed bioclimatic zones were very similar to those of A. bovis infection (Figure 3A).

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3.1.2. In sheep In sheep, A. ovis infection begins relatively low (30.7%) during the spring, then increases during the summer (36.1%), reaches its maximum during the autumn (43.6%) and finally the infection rate decreases during the winter season (31.8%). This fluctuation between seasons in A. ovis infection rate is statistically significant (P = 0.010) (Figure 1B). Sheep located in lower humid, higher semi-arid and sub-humid areas were statistically more infected by A. ovis, respectively, in summer (47.7%), autumn (50%) and winter (52.4%) than in other seasons (Figure 2B). A. bovis infection and A. ovis and A. bovis co-infection profiles are similar and the rates begin relatively low (0.9%) during the spring, then increase and reach their maximum during the summer (18.1 and 17.7%, respectively) and decrease during the autumn (7.6 and 6.8%, respectively) and the winter seasons (3%). These differences between seasons in A. bovis infection and A. ovis/A. bovis co-infection rates are statistically significant (P < 0.001; Figure 1B). In lower humid and higher semi-arid areas, summer was the season when there were more sheep statistically infected by A. bovis (P < 0.001; Figure 3B). In the sub-humid area, sheep tested during the winter (14.3%) were more infected but the difference between seasons remains statistically insignificant in this bioclimatic area (P = 0.090; Figure 3B). As shown in Figure 4B, A. ovis/A. bovis co-infection profiles in the three analyzed bioclimatic zones were very similar to those of A. bovis infection (Figure 3B). 3.2. Molecular characterization of Anaplasma species 3.2.1. Anaplasma ovis msp4 genetic variants A. ovis infection was confirmed by sequencing of 808 bp of the msp4 gene (94.8% of the gene size) from three randomly selected positive goat samples and two positive sheep samples. The msp4 gene sequences obtained in this study shared 99.8 to 100% and 99.9 to 100% nucleotides and amino acids similarity, respectively (Table 2). Alignment of these sequences revealed one genotype (AoOv1) in sheep (GenBank accession numbers KY659320 and KY659321) and one genotype (AoCp1) in goats (GenBank accession numbers KY659322 to KY659324) differing from each other in one nucleotide position (470) conferring one amino acid variation in position 157 (Tables 2 and 3). AoOv1 genotype 5

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revealed 100% homology with GBK1 and AoGOv5 genotypes (GenBank accession numbers KC432641 and KM285222) from Tunisia, and the genotype II represented by "Italy147" A. ovis strain from Sicilian sheep (GenBank accession number AY702924) (Table 3). AoCp1 genotype were 100% identical to GBK2, AoGOv1 and AoGGo1 genotypes (GenBank accession numbers KC432642, KM285218 and KM285217) from Tunisia, and to the genotype III represented by "Italy20" A. ovis strain from Sicilian sheep (GenBank accession number AY702923, Table 3).

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3.2.2. Anaplasma bovis 16S rRNA genetic variants A. bovis infections were validated by sequencing of 511 bp 16S rRNA fragment (33.7% of the gene size) from randomly selected nine goat samples and three sheep samples. Alignment of these sequences revealed one genotype from goats and sheep (AbCpOv1; GenBank accession number KY655797-KY655806), one genotype from goats (AbCp1; GenBank accession number KY655807) and one genotype from sheep (AbOv1; GenBank accession number KY655808). AbCpOv1 was 100% identical to AbGGo1, AbGOv1 and AbGBv3 variants (GenBank accession numbers KM285223, KM285224 and KM401904, respectively) isolated from Tunisian goats, sheep and cattle, respectively and to G55 variant isolated from goat in China (GenBank accession number JN558825) (Table 2). AbOv1 was 100% identical to AbGBv2 genotype isolated from Tunisian cattle and different from AbCpOv1 genotype by one substitution with an identity rate estimated at 99.8% (Tables 2 and 3). AbCp1 was a novel genotype and different from AbCpOv1 and AbOv1 genotypes by one and two substitutions with identity rates estimated at 99.8 and 99.6%, respectively (Tables 2 and 3).

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4. Discussion Previous studies have documented the presence and the phylogeny of Anaplasma species in small ruminants from Tunisia (Belkahia et al., 2014, Ben Said et al., 2015a, b; 2017a, b), but this study is the first outlining seasonal dynamics of Anaplasma spp. infections in sheep and goats from our country. In this longitudinal investigation, results clearly indicate evidence of A. ovis infection in small ruminants sampled in all investigated bioclimatic areas and during the four seasons. In sheep, the average prevalence rate of A. ovis infection was 35.6% (minimum 30.7% in spring and maximum 43.6% in autumn). It was similar to those reported in China (40.5%) (Yang et al., 2015), Sudan (41.7%) (Renneker et al., 2013) and Turkey (31.4%) (Renneker et al., 2013), higher than what observed in Senegal (11.5%) (Djiba et al., 2013) and lower than that estimated in Italy (87-47.3%) (de la Fuente et al., 2005; Torina et al., 2008a, b; 2010; Torina and Caracappa, 2012), Portugal (82.5%) (Renneker et al., 2013) and in other Tunisian regions (70.1 and 93.8%, respectively) (Belkahia et al., 2014; Ben Said et al., 2015a). The average prevalence rate of A. ovis infection in goats was 46% (minimum 21.7% in summer and maximum 65.5% in winter). This rate was higher than that observed in Italy (31.7%) (Torina and Caracappa, 2012) and China (15.3-25.6%) (Liu et al., 2012; Chi et al., 2013) and lower than that estimated in Angola (100%) (Kubelová et al., 2012), Turkey (66.4%) (Altay et al., 2014) and in other Tunisian regions (65.3%) (Ben Said et al., 2015a). Molecular survey of A. bovis in small ruminants showed that average prevalence was 7.4% (minimum 0.9% in spring and maximum 18.1% in summer) in sheep and 10.1% (minimum 2.2% in autumn and maximum 23.8% in summer) in goats. These two close A. bovis prevalence rates were lower than that found in sheep (42.7%) and goats (23.8%) from other Tunisian regions (Ben Said et al., 2015b), in goats from China (49.6%) (Liu et al., 2012) and in deer (23%) (Kawahara et al., 2006) and cattle (15–53.3%) (Ooshiro et al., 2008; Jilintai et al., 2009) from Japan. These same rates were higher than what observed in cattle from Iran (2.7%) (Noaman and Shayan, 2010), India (3.3%) (Nair et al., 2013) and Italy (4.2%) (Ceci et al., 2014).

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These differences observed in the prevalence rates of Anaplasma species among different countries and areas of the same country may be due to several factors including the various methods of sample analysis, the presence and abundance of incriminated ticks and/or other vectors and its control programmes, the farm management and husbandry practices, the sampling seasons and/or other bioclimatic or ecological factors, the susceptibility of each host species and breeds, and the presence of wildlife reservoirs (Torina et al., 2008a; Belkahia et al., 2014; Ben Said et al., 2015a and b). However, A. phagocytophilum was not detected in all investigated small ruminants analyzed during the four seasons. Same results were reported in these animal species from Italy, Morocco and other regions from Tunisia (Zobba et al., 2014; Ait Lbacha et al., 2015; Ben Said et al., 2015a). It can be suggested that small ruminants located in these studied sites are not main reservoirs for this zoonotic species, and probably other domestic animals like cattle, dogs and horses could be involved as reservoir hosts in these areas (M’ghirbi et al., 2009, 2012, 2016). However, two types of strains (A. phagocytophilum-like 1 and 2) genetically related to A. phagocytophilum have been found in small ruminants located in these regions, but until now these infected animals have not shown any clinical signs relating to tick-borne fever (Ben Said et al., 2015a, 2017). Except for sheep located in the sub-humid zone, almost all A. bovis infection profiles in overall and according to each bioclimatic area show that small ruminants analyzed during the summer season are statistically the most infected by this Anaplasma species (Figures 1 and 3). This finding suggests that the main vectors of A. bovis in small ruminants have essentially a summer activity such as Rhipicephalus and Hyalomma ticks (Bouattour et al., 2002). This is reinforced by a previous parasitological study carried out on small ruminant populations located in northern Tunisia, analyzed in spring and summer, and found to be infected by A. bovis (Ben Said et al., 2015b). In this study, we have demonstrated the infestation of these animals by several tick species belonging to Rhipicephalus and Hyalomma genera like R. turanicus, R. sanguineus, R. bursa, R. annulatus and H. excavatum. These tick species are known to have a summer infestation activity. In addition, Hyalomma species have been proposed previously as vectors of A. bovis in Africa (Donatien and Lestoquard, 1936). More recently, Harrus et al. (2011) have isolated A. bovis from R. turanicus and R. sanguineus. Until now, the vectors of A. bovis are still unidentified in the country; from where, investigations on ticks are needed to know the main vectors of this bacterium in Tunisian ruminants. In our study, high A. ovis prevalence rates were observed in tested small ruminants in overall (minimum 23.8% and maximum 65.5%) and from each bioclimatic area (minimum 11.6% and maximum 100%) during the four seasons (Figures 1 and 2). It points out that there are probably two groups of vectors transmitting this bacteria in these investigated regions (i) one that has a peak of activity in autumn in Tunisia like Ixodes ricinus, Rhipicephalus (Boophilus) annulatus, Dermacentor marginatus, Haemaphysalis punctata and H. sulcata (Morel, 1965; Van den Ende, 1970; YousfiMonod and Aeschlimann, 1986; Bouattour et al., 1999, 2002) and (ii) another group of vectors that probably has an optimal infestation activity during the spring and/or the summer (from April to August) in this country such as R. turanicus, R. Bursa, R. sanguineus, Hyalomma marginatum marginatum and H. excavatum (Morel, 1965; Van den Ende, 1970; Yousfi-Monod and Aeschlimann, 1986; Bouattour et al., 1999, 2002). However, most of these species have been previously proposed as vectors of A. ovis worldwide including Rhipicephalus spp., Hyalomma spp. and Dermacentor spp. (Friedhoff, 1997; Yin and Luo, 2007). In fact, DNA of A. ovis has been detected in R. sanguineus, R. turanicus and/or R. bursa in the Mediterranean countries such as Italy (Torina et al., 2008a), Turkey (Aktas et al., 2009), France (Dahmani et al., 2017) and Algeria (Aouadi et al., 2017). The tick D. andersoni is considered the main vector of A. ovis infection (Friedhoff, 1997), and Hornok et al. (2007) reported that D. marginatus may be involved in the transmission of this Anaplasma species. In addition, Uilenberg (1997) suggested that several species of arthropods can be involved in the transmission of A. ovis. In this respect, Hashemi-Fesharki (1997) reported that biting flies are 7

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involved in the transmission of A. ovis, and Hornok et al. (2011) detected A. ovis in sheep keds (Melophagus ovinus), indicating that these hippoboscid flies which infest sheep and occasionally goats, especially during the spring and winter seasons, may have a transmission role. However, in Tunisia, the detection and the prevalence in vectors in general and especially in ticks and its genetic variability, which could be found between hosts and vectors, have not been studied yet. Furthermore, seasonal A. ovis/A. bovis co-infection and A. bovis infection profiles confirm that almost all small ruminants infected by A. bovis are also infected by A. ovis (Figure 1). The same trend has been observed in a cross-sectional study performed between May and September on Chinese goats (Liu et al., 2012). This finding suggests that, in summer, these two bacterial species have probably one or several common vectors having certainly an intense infestation activity in this season. However, A. ovis infection rate in goats decreased significantly during the summer while A. bovis prevalence rate reached its maximum in the same animals (Figure 1A). Although it has been shown that some strains of A. marginale, A. ovis, and A. phagocytophilum can prevent the multiplication of other strains (de la Fuente et al., 2002, 2007; Stuen et al., 2005), the relation between the present data and the competition among Anaplasma species in the analyzed goats and/or common vectors in favor of A. bovis remains to be confirmed by other investigations. The analysis of A. bovis 16S rRNA sequences revealed a perfect homology between Tunisian variants isolated from goats, sheep and cattle (Table 3) suggesting the occurrence of an unique transmission cycle of A. bovis in analyzed regions involving, at least, these three ruminant species and probably common tick species or/and other vectors. Additionally, the low diversity of the 16S rRNA sequences from A. bovis strains found in several Tunisian regions and the apparent absence of clinical signs associated with this infection suggest a small geographical segregation of these Tunisian strains which seem to have a limited pathogenicity in infected small ruminants. Similar findings were reported in domesticated ruminants from other Tunisian regions (Belkahia et al., 2015; Ben Said et al., 2015b) and in cattle from Japan and India (Jilintai et al., 2009; Nair et al., 2013). The alignment and the comparison of the msp4 sequences of A. ovis variants revealed a low genetic variability and, in five sequenced positive samples, only two genetic variants that differ in C/T substitution at position 470 were detected (Table 3). Both variants were previously described in Italy (AOG2 and AOG3) (de la Fuente et al., 2005). In contrast to our results, msp4 sequence diversity of A. ovis strains from Hungary (Hornok et al., 2007) and Italy (Torina et al., 2010) showed higher polymorphism represented by five and seventeen variants in Hungary and Italy respectively. Additionally, AoOv1 variant was detected only in sheep and AoCp1 variant was detected only in goats. Concordantly, Liu et al. (2012) suggest that A. ovis msp4 genotypes may be different among sheep and goats. In summary, we conclude that anaplasmosis caused by A. ovis is endemic in small ruminants from all investigated bioclimatic areas during the four seasons but conversely, A. bovis infection is highly intensified only in the summer and A. phagocytophilum infection is absent or very rare in small ruminants from these Tunisian regions. This situation with those of further longitudinal surveys, necessary to be performed in other ruminants, must be taken into account if anaplasmosis control programs in these domesticated animals are envisaged. Acknowledgements This work was supported by the research project “2 PS1.3.023 – RESTUS” funded by the European Neighbourhood and Partnership Instrument (ENPI) - Transboundary Cooperation (TC) - ItalyTunisia 2007-2013, the “Laboratoire d’épidémiologie d’infections enzootiques des herbivores en Tunisie” (LR02AGR03), funded by the Ministry of Higher Education and Scientific Research of Tunisia, and the research project “Epidémiologie de maladies bactériennes à transmission vectorielle des herbivores” (06-680-0029) funded by the Ministry of Agriculture of Tunisia. The authors would 8

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like to thank Dr. Leïla Sayeh, Dr. Saber Hdhiri, Dr. Aymen Sahbani, Dr. Tarak Blaïech, Dr. Oussama Mathlouthi, Dr. Said Jaajaa and Dr. Taoufik Ben Hamida and their technicians for their help and facilitating the access to the farmers.

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Competing interests No competing financial interests exist.

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Ceci, L., Iarussi, F., Greco, B., Lacinio, R., Fornelli, S., Carelli, G., 2014. Retrospective study of hemoparasites in cattle in southern Italy by reverse line blot hybridization. J. Vet. Med. Sci. 76, 869–875. Chi, Q., Liu, Z., Li, Y., Yang, J., Chen, Z., Yue, C., Luo, J., Yin, H., 2013. Development of a realtime PCR assay for detection and quantification of Anaplasma ovis infection. Transbound. Emerg. Dis. 60, 119–124. Dahmani, M., Davoust, B., Rousseau, F., Raoult, D., Fenollar, F., Mediannikov, O., 2017. Natural Anaplasmataceae infection in Rhipicephalus bursa ticks collected from sheep in the French Basque Country. Ticks Tick Borne Dis. 8, 18–24. de la Fuente, J., Garcia-Garcia, J.C., Blouin, E.F., Saliki, J.T., Kocan, K.M., 2002. Infection of tick cells and bovine erythrocytes with one genotype of the intracellular ehrlichia Anaplasma marginale excludes infection with other genotypes. Clin. Diagn. Lab. Immunol. 9, 658–668. de la Fuente, J., Torina, A., Caracappa, S., Tumino, G., Furlá, R., Almazán, C., Kocan, K.M., 2005. Serologic and molecular characterization of Anaplasma species infection in farm animals and ticks from Sicily. Vet. Parasitol. 133, 357–362. de la Fuente, J., Atkinson, M.W., Naranjo, V., Fernandez de Mera, I.G., Mangold, A.J., Keating, K.A., Kocan, K.M., 2007. Sequence analysis of the msp4 gene of Anaplasma ovis strains. Vet. Microbiol. 119, 375–381. Djiba, M.L., Mediannikov, O., Mbengue, M., Thiongane, Y., Molez, J.F., Seck, M.T., Fenollar, F., Raoult, D., Ndiaye, M., 2013. Survey of Anaplasmataceae bacteria in sheep from Senegal. Trop. Anim. Health Prod. 45, 1557–1561. Donatien, A., Lestoquard, F., 1936. Rickettsia bovis, nouvelle espèce pathogène pour le boeuf. Bull. Soc. Pathol. Exot. 29, 1057–1061. Dumler, J.S., Barbet, A.F., Bekker, C.P., Dasch, G.A., Palmer, G.H., Ray, S.C., Rikihisa, Y., Rurangirwa, F.R., 2001. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and ‘HGE agent’ as subjective synonyms of Ehrlichia phagocytophilum. Int. J. Syst. Evol. Microbiol. 51, 2145–2165. Friedhoff, K.T., 1997. Tick-borne diseases of sheep and goats caused by Babesia, Theileria or Anaplasma spp. Parassitologia 39, 99–109. Harrus, S., Perlman-Avrahami, A., Mumcuoglu, K.Y., Morick, D., Eyal, O., Baneth, G., 2011. Molecular detection of Ehrlichia canis, Anaplasma bovis, Anaplasma platys, Candidatus Midichloria mitochondrii and Babesia canis vogeli in ticks from Israel. Clin. Microbiol. Infect. 17, 459–463. Hashemi-Fesharki, R., 1997. Tick-borne diseases of sheep and goats and their related vectors in Iran. Parassitologia 39, 115–117. Hornok, S., Elek, V., de la Fuente, J., Naranjo, V., Farkas, R., Majoros, G., Földvári, G., 2007. First serological and molecular evidence on the endemicity of Anaplasma ovis and A. marginale in Hungary. Vet. Microbiol. 122, 316–322. Hornok, S., de la Fuente, J., Biró, N., Fernández de Mera, I.G., Meli, M.L., Elek, V., Gönczi, E., Meili, T., Tánczos, B., Farkas, R., Lutz, H., Hofmann-Lehmann, R., 2011. First molecular evidence of Anaplasma ovis and Rickettsia spp. in keds (Diptera: Hippoboscidae) of sheep and wild ruminants. Vector Borne Zoonotic Dis. 11, 1319–1321. Jilintai, Seino, N., Hayakawa, D., Suzuki, M., Hata, H., Kondo, S., Matsumoto, K.,Yokoyama, N., Inokuma, H., 2009. Molecular survey for Anaplasma bovis and Anaplasma phagocytophilum infection in cattle in a pastureland where sika deer appear in Hokkaido, Japan. Jpn. J. Infect. Dis. 62, 73–75.

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Kawahara, M., Rikihisa, Y., Lin, Q., Isogai, E., Tahara, K., Itagaki, A., Hiramitsu, Y., Tajima, T., 2006. Novel genetic variants of Anaplasma phagocytophilum, Anaplasma bovis, Anaplasma centrale, and a novel Ehrlichia sp. in wild deer and ticks on two major islands in Japan. Appl. Environ. Microbiol. 72, 1102–1109. Kocan, K.M., de la Fuente, J., Blouin, E.F., Garcia-Garcia, J.C., 2004. Anaplasma marginale (Rickettsiales: Anaplasmataceae): recent advances in defining host-pathogen adaptations of a tick-borne rickettsia. Parasitol. 125, S285–300. Kubelová, M., Mazancová, J., Siroký, P., 2012. Theileria, Babesia, and Anaplasma detected by PCR in ruminant herds at Bié Province, Angola. Parasite 19, 417–422. Liu, Z., Ma, M., Wang, Z., Wang, J., Peng, Y., Li, Y., Guan, G., Luo, J., Yin, H., 2012. Molecular survey and genetic identification of Anaplasma species in goats from central and southern China. Appl. Environ. Microbiol. 78, 464–470. M’ghirbi, Y., Ghorbel, A., Amouri, M., Nebaoui, A., Haddad, S., Bouattour, A., 2009. Clinical, serological, and molecular evidence of ehrlichiosis and anaplasmosis in dogs in Tunisia. Parasitol. Res., 104, 767–774. M’ghirbi, Y., Yaïch, H., Ghorbel, A., Bouattour, A., 2012. Anaplasma phagocytophilum in horses and ticks in Tunisia. Parasit. Vectors 30, 180. M'ghirbi, Y., Bèji, M., Oporto, B., Khrouf, F., Hurtado, A., Bouattour, A., 2016. Anaplasma marginale and A. phagocytophilum in cattle in Tunisia. Parasit. Vectors 9, 556. Morel, P.C., 1965. Les tiques de l’Afrique et du Bassin Méditerranéen (Ixodoidae). Document multigrad. IEMVPT Maisons-Alfort, 332p. Nair, A.S., Ravindran, R., Lakshmanan, B., Sreekumar, C., Kumar, S.S., Raju, R., Tresamol, P.V., Vimalkumar, M.B., Saseendranath, M.R., 2013. Bovine carriers of Anaplasma marginale and Anaplasma bovis in South India. Trop. Biomed. 30, 105–112. Noaman, V., Shayan, P., 2010. Molecular detection of Anaplasma bovis in cattle from central part of Iran. Vet. Res. Forum 1, 117–122. Ogden, N.H., Lindsay, L.R., 2016. Effects of climate and climate change on vectors and vector-borne diseases: Ticks are different. Trends Parasitol. 32, 646-656. Ooshiro, M., Zakimi, S., Matsukawa, Y., Katagiri, Y., Inokuma, H., 2008. Detection of Anaplasma bovis and Anaplasma phagocytophilum from cattle on Yonaguni island, Okinawa, Japan. Vet. Parasitol. 154, 360–364. Renneker, S., Abdo, J., Salih, D.E.A., Karagenç, T., Bilgiç, H., Torina, A., Oliva, A.G., Campos, J., Kullmann, B., Ahmed, J., Seitzer, U., 2013. Can Anaplasma ovis in small ruminants be neglected any longer? Transbound. Emerg. Dis. 60, 105–112. Stuen, S., Dahl, H., Bergström, K., Moum, T., 2005. Unidirectional suppression of Anaplasma phagocytophilum genotypes in infected lambs. Clin. Diagn. Lab. Immunol. 12, 1448–1450. Stuen, S., 2007. Anaplasma phagocytophilum - the most widespread tick-borne infection in animals in Europe. Vet. Res. Commun. 31, 79–84. Torina, A., Alongi, A., Naranjo, V., Estrada-Peña, A., Vicente, J., Scimeca, S., Marino, A.M., Salina, F., Caracappa, S., de la Fuente, J., 2008a. Prevalence and genotypes of Anaplasma species and habitat suitability for ticks in a Mediterranean ecosystem. Appl. Environ. Microbiol. 74, 7578– 7584. Torina, A., Alongi, A., Naranjo, V., Scimeca, S., Nicosia, S., Di Marco, V., Caracappa, S., Kocan, K.M., de la Fuente, J., 2008b. Characterization of Anaplasma infections in Sicily, Italy. Ann. N. Y. Acad. Sci. 1149, 90–93. Torina, A., Galindo, R.C., Vicente, J., Di Marco, V., Russo, M., Aronica, V., Fiasconaro, M., Scimeca, S., Alongi, A., Caracappa, S., Kocan, K.M., Gortazar, C., de la Fuente, J., 2010. Characterization of Anaplasma phagocytophilum and A. ovis infection in a naturally infected sheep flock with poor health condition. Trop. Anim. Health Prod. 42, 1327–1331. 12

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Torina, A., Agnone, A., Blanda, V., Alongi, A., D’Agostino, R., Caracappa, S., Marino, A.M.F., Di Marco, V., de la Fuente, J., 2012. Development and validation of two PCR tests for the detection of and differentiation between Anaplasma ovis and Anaplasma marginale. Ticks Tick-borne Dis. 3, 283–287. Torina, A., Caracappa, S., 2012. Tick-borne diseases in sheep and goats: clinical and diagnostic aspects. Small Rum. Res. 106, S6–S11. Tuomi, J., 1967. Experimental studies on bovine tick-borne fever .1. Clinical and haematological data, some properties of the causative agent, and homologous immunity. Acta Pathol. Microbiol. Scand. 70, 429–445. Uilenberg, G., 1997. General review of tick-borne diseases of sheep and goats worldwide. Parassitologia 39, 161–165. Van den Ende, M., 1970. [The ticks (Ixodidae) of domestic animals in Tunisia and their biology]. Arch. Inst. Pasteur de Tunis 47, 253–264. Woldehiwet, Z., 2010. The natural history of Anaplasma phagocytophilum. Vet. Parasitol. 167, 108– 122. Yang, J, Li, Y., Liu, Z., Liu, J., Niu, Q., Ren, Q., Chen, Z., Guan, G., Luo, J., Yin, H., 2015. Molecular detection and characterization of Anaplasma spp. in sheep and cattle from Xinjiang, northwest China. Parasit. Vector 8, 108. Yin, H., Luo, J., 2007. Ticks of small ruminants in China. Parasitol. Res. 101, 187–189. Yousfi-Monod, R., Aeschlimann, A., 1986. [Research on ticks (Acarina, Ixodidae) infesting cattle in north western Algeria. I. Systematical survey and seasonal activity]. Ann. Parasit. Hum. Comp. 61, 341–358. Zobba, R., Anfossi, A.G., Pinna Parpaglia, M.L., Dore, G.M., Chessa, B., Spezzigu, A., Rocca, S., Visco, S., Pittau, M., Alberti, A., 2014. Molecular investigation and phylogeny of Anaplasma spp. in Mediterranean ruminants reveal the presence of neutrophil-tropic strains closely related to A. platys. Appl. Environ. Microbiol. 80, 271–280.

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Figure legends Figure 1 Seasonal profiles of single and mixed infections by Anaplasma ovis and A. bovis in analyzed goats (A) and sheep (B). Abbreviation: P-values are in brackets; *: Statistically significant test.

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Figure 2 Seasonal profiles of Anaplasma ovis infection in tested goats (A) and sheep (B) from each bioclimatic area. Abbreviation: P-values are in brackets; *: Statistically significant test.

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Figure 3 Seasonal profiles of Anaplasma bovis infection in tested goats (A) and sheep (B) from each bioclimatic area. Abbreviation: P-values are in brackets; *: Statistically significant test.

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Figure 4 Seasonal profiles of A. ovis/A. bovis co-infection in investigated goats (A) and sheep (B) from each bioclimatic area. Abbreviation: P-values are in brackets; *: Statistically significant test.

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Supplementary legends Supplementary file S1 Geographic location of the Tunisian studied regions. (A): Map of Tunisia showing the governorates of Tunis, Ariana, Bizerte, Beja and Nabeul, (B): Map of the North of Tunisia showing the 30 delegations according to bioclimatic areas.

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Table 1 Primers used for detection and/or characterization of Anaplasma species in small ruminants in the present study. Targ et gene

Amplic on size (bp)

Refere nce

EE-1

TCCTGGCTCAGAACGAACGCTGGCG GC

16S AR Nr

1433

Barlou gh et al. (1996)

EE-2

AGTCACTGACCCAACCTTAAATGGC TG

AB1f

CTCGTAGCTTGCTATGAGAAC

16S AR Nr

551

Kawah ara et al. (2006)

AB1r

TCTCCCGGACTCCAGTCTG

AovisMSP4 Fw

TGAAGGGAGCGGGGTCATGGG

msp 4

344

Torina et al. (2012)

AovisMSP4 Rev

GAGTAATTGCAGCCAGGGACTCT

MSP45

GGGAGCTCCTATGAATTACAGAGAA TTGTTTAC

msp 4

852

de la Fuente et al. (2005)

groE 624 L

Alberti et al. (2005)

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PCR 22 A. bovis

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PCR 11 Anaplasma spp.

EphplgroEL -F

ATGGTATGCAGTTTGATCGC

EphplgroEL -R EphplgroEL -F EphgroELR

TCTACTCTGTCTTTGCGTTC

AC

Nested PCR5 A. phagocytoph ilum

CCGGATCCTTAGCTGAACAGGAATC TTGC

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MSP43

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Single PCR4 A. ovis

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Single PCR A. ovis

Sequence 5’ to 3’

Primer

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Assay

ATGGTATGCAGTTTGATCGC

573

TTGAGTACAGCAACACCACCGGAA

1

: First PCR allowing the detection of all Anaplasma species. : Second PCR allowing the species detection and characterization of A. bovis. 3 : Single PCR allowing the detection of A. ovis. 4 : Single PCR allowing the characterization of A. ovis after sequencing. 5 : Nested PCR allowing the detection of A. phagocytophilum. 2

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Table 2 Designation and information about sequencing of Anaplasma ovis and A. bovis genetic variants identified in this study. Gene

Sequence type

Isolate

Host ( Reference number)

Geographi cal location

GenBank accession no.

A. ovis

msp4

AoOv1

Ao1

Ovis aries (T5)

Tunis

KY659320 100% A. ovis (AY70292 4, HQ456348 )

Ao2

Ovis aries (T11)

RI SC

KY65932 1

100% A. ovis (AY70292 4, HQ45634 8)

Capra hircus (T16)

Tunis

KY65932 2

100% A. ovis (AY70292 3, KJ782397 , HQ45634 7)

Ao4

Capra hircus (Bz4)

Bizerte

KY65932 3

100% A. ovis (AY70292 3, KJ782397 , HQ45634 7)

Ao5

Capra hircus (Bz12)

Bizerte

KY65932 4

100% A. ovis (AY70292 3,

Ao3

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KJ782397 , HQ45634 7) Ab1

Ovis aries (Bj14)

Ab2

Capra hircus (Bz8)

Beja

KY65579 7

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16S rRNA AbCpOv1

100% A. bovis (KJ78239 7, HQ45634 7, AY70292 3)

KY65579 8

100% A. bovis (KM2852 23, KM28522 4)

Capra hircus (Bz9)

Bizerte

KY65579 9

100% A. bovis (KM2852 23, KM28522 4)

Ab4

Capra hircus (Bj7)

Beja

KY65580 0

100% A. bovis (KM2852 23, KM28522 4)

Ab5

Capra hircus (Bj8)

Beja

KY65580 1

100% A. bovis (KM2852 23, KM28522 4)

Ab6

Capra hircus

Beja

KY65580 2

100% A. bovis (KM2852

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Ab3

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A. bovis

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(Bj14)

23, KM28522 4)

Capra hircus (Bj15)

Beja

Ab8

Capra hircus (Bj17)

Beja

KY65580 3

100% A. bovis (KM2852 23, KM28522 4)

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Beja

KY65580 5

100% A. bovis (KM2852 23, KM28522 4)

Ab10

Ovis aries (Bz14)

Bizerte

KY65580 6

100% A. bovis (KM2852 23, KM28522 4)

AbCp1

Ab11

Capra hircus (Bz5)

Bizerte

KY65580 7

99% A. bovis (KM2852 23, KM28522 4)

AbOv1

Ab12

Ovis aries (Bz13)

Bizerte

KY65580 8

100% A. bovis (KM4019 03, KM28522

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Capra hircus (Bj18)

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Ab9

100% A. bovis (KM2852 23, KM28522 4)

KY65580 4

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Ab7

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AY702923 and AY702924 are the GenBank accession numbers of Italy 20 and Italy 147 strains isolated from Italian sheep (de la Fuente et al., 2005). KJ782397 is the GenBank accession number of ATS20 isolate found on Chinese sheep (Yang et al., 2015). HQ456347 and HQ456348 are the GenBank accession numbers of Yongjing and Yuzhong isolates found on Chinese sheep (Ma et al., 2011). KM285223 is the GenBank accession number of AbGGo1 variant isolated from Tunisian goat (Ben Said et al., 2015b). KM285224 and KM285225 are the GenBank accession numbers of AbGOv1 and AbGOv2 variants isolated from Tunisian sheep (Ben Said et al., 2015b). KM401903 is the GenBank accession number of AbGBv2 variant isolated from Tunisian cattle (Belkahia et al., 2015).

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Table 3 Diversity among nucleotide and amino acid msp4 sequences from Anaplasma ovis (808 bp) and among nucleotide 16S rRNA sequences from Anaplasma bovis (511 bp) isolated from Tunisian strains.

KC432641

GBK2

KC432642

GB3

KC432643

GK1

KC432644

AoGOv1

KM285218

AoGOv2

KM285219

AoGOv3

KM285220

AoGOv4

KM285221

AoGOv5 AoGGo1

KM285222 KM285217

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NU

Sheep GBK1

PT E CE

Goat

A. bovis

AC

Sheep AoOv1

Nucleotide positions (amino acid Reference positions)3 230 244 470 476 532 (178) (77) (83) (157) (159) G A C C C (L) Belkahia (R) (S) (A) et al. (2014) * * T * * (V) T * T * * (I) (V) T * T * A (I) (I) (V) * * T * * Ben (V) Said et al. (2015a) * G T * * (G) (V) T * T A * (I) (V) * * T A * (V) * * * * * * * T * * (V) * * * * * This study * * T * * (V) 82 166 296 412

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GenBank2

RI

Variant1

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Host

D

Anaplasma Gene sp. A. ovis msp4

KY659320

Goat

AoCp1

KY659322

Goat

AbGGo1

KM285223

A

A

G

G

Sheep AbGOv1 AbGOv2 Cattle AbGBv1

KM285224 KM285225 KM401902

* G G

* T T

* * *

* * *

16S rRNA

Ben Said et al. (2015b)

Belkahia et al. 24

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(2015) AbGBv2 KM401903 AbGBv3 KM401904 AbCpOv1 KY655797

* * *

T * *

* * *

* * *

KY655807 KY655808

* *

* T

A *

C *

Goat and sheep Goat AbCp1 Sheep AbOv1

This study

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: AoOv1 variant was also represented by GenBank accession number KY659321, AoCp1 variant was also represented by GenBank accession numbers KY659323 and KY659324, and AbCpOv1 variant was also represented by GenBank accession numbers KY655798 to KY655806. 2 : GenBank accession number of the genetic variant. 3 : Numbers represent the nucleotide position with respect to A. ovis Italy 147 strain from Italy (GenBank accession number AY702924) and to A. bovis G55 isolate (clone 55) from China (GenBank accession number JN558825). Conserved nucleotide positions relative to the first sequence are indicated with asterisks. Amino acid changes are indicated between parentheses with single letter code. Amino acids: R, Arginine; I, Isoleucine; S, Serine; G, Glycine; V, Valine; A, Alanine; L, Leucine; Nucleotides: T, Thymine; C, Cytosine; G, Guanine; A, Adenine.

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Highlights First longitudinal field study in Anaplasma species in small ruminants from Tunisia.

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Average prevalence rates of A. ovis and A. bovis were 46 and 10.1% in goats, and 35.6 and

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7.4% in sheep, respectively.

A. phagocytophilum was not detected in all investigated animals.

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Seasonal and bioclimatic variations of Anaplasma spp. infection and co-infection rates were

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Genotyping of msp4 and 16S rRNA genes revealed two and three different variants of A. ovis

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and A. bovis, respectively.

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