- Email: [email protected]
Phylogeographical structure of the tick Ixodes persulcatus: A novel view
Ticks and Tick-borne Diseases 3 (2012) 212–218
Contents lists available at SciVerse ScienceDirect
Ticks and Tick-borne Diseases journal homepage: www.elsevier.de/ttbdis
Original article
Phylogeographical structure of the tick Ixodes persulcatus: A novel view Sergey Y. Kovalev ∗ , Tatyana A. Mukhacheva Laboratory of Molecular Genetics, Department of Biology, Ural Federal University, Lenin Avenue 51, Yekaterinburg 620000, Russia
a r t i c l e
i n f o
Article history: Received 18 January 2012 Received in revised form 19 March 2012 Accepted 19 March 2012 Keywords: Ixodes persulcatus Phylogeography Preferential amplification rRNA Population structure
a b s t r a c t The tick Ixodes persulcatus Schulze, 1930, has a wide distribution from the Baltic to the Far East and is a vector of a number of human pathogens. Thus, the study of the genetic structure and evolution of this species is of great epidemiological importance. rRNA genes were used as genetic markers to identify the phylogeographical structure of the ticks. The sequences of gene fragments of 28S (expansion segment D3) and mitochondrial 12S rRNA for 25 and 76 ticks, respectively, that had been collected in various regions of Russia in 2007–2011, were obtained. The sequences of the 28S rRNA D3 segment were identical for all ticks within the studied area. Analysis of the sequences of the mitochondrial 12S rRNA fragment revealed 4 haplotypes with one occurring at a frequency of 0.96. It is shown that the ‘deep’ population structure of I. persulcatus (McLain et al., 2001) was erroneous because of the inclusion of contaminating fungi sequences of 28S rRNA in the phylogenetic analysis. This was, possibly, due to the use of universal PCR primers that amplify the DNA of a wide range of eukaryotes, particularly of fungi which are common in samples of ticks. The influence of PCR conditions on the preferential amplification of the DNA of different organisms is also demonstrated. © 2012 Elsevier GmbH. All rights reserved.
Introduction The tick Ixodes persulcatus Schulze, 1930, commonly known as the taiga tick, is a vector of a number of human pathogens including tick-borne encephalitis virus (Zilber, 1939), Borrelia burgdorferi s.l. (Kriuchechnikov et al., 1988), and Anaplasma phagocytophilum (Shpynov et al., 2004). This species belongs to the Ixodes ricinus complex – a group of ticks distributed in almost all geographic regions of the world (Xu et al., 2003). The tick I. persulcatus inhabits the taiga zone from the Baltic to the Russian Far East as well as Japan, northern China, Mongolia, and Kazakhstan (Fig. 1) (Filippova, 1985) and is the principal vector of tick-borne infections in Russia. Given such a wide range and epidemiological significance, the evolution of I. persulcatus is of great scientific interest. To date, a lot of information concerning the morphology, biology, and ecology of I. persulcatus has been acquired (Alekseev et al., 2000; Filippova, 1985, 2002; Filippova and Musatov, 1996). Moreover, some attempts to study the intraspecific structure of the taiga tick based on a comparison of morphological features were made, and some differences in body size between Asian and European populations were found (Filippova and Musatov, 1996). The study by Filippova (1985) highlights the difficulty of investigating the geographic variation in this species because of the wide geographic range and the inconsistency of data on the
∗ Corresponding author. Tel.: +7 343 2616826; fax: +7 343 3507401. E-mail address: [email protected] (S.Y. Kovalev). 1877-959X/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.ttbdis.2012.03.005
variability of morphological structures at different developmental stages of ticks (Filippova, 1985). Generally, morphological traits may have a complex genetic basis and are often dependent on environmental conditions. Therefore, genetic markers should be used for such population studies. They were shown to be useful in both systematic and population analyses of ticks (Nava et al., 2009). Several studies on ticks belonging to the Ixodes ricinus complex (Casati et al., 2008; Norris et al., 1996) allowed a comparative analysis of their genetic structure to be carried out and the processes governing tick distribution to be understood. However, little is known about the population structure of I. persulcatus. In this regard, special attention should be paid to the work of McLain et al. (2001) who suggested extreme heterogeneity and deep geographical structure of the I. persulcatus population, based on the nucleotide sequence of the expansion segment D3 28S rRNA, as well as a significant difference between the secondary structure of this segment compared to other ticks of the I. ricinus complex (McLain, 2001). Such a structure suggested a long and independent evolution of I. persulcatus populations. As pathogens transmitted by I. persulcatus also show heterogeneous genetic structure (Ecker et al., 1999; Fomenko et al., 2009; Nefedova et al., 2010), they can form an excellent basis for the study of coevolutionary processes in populations of pathogens and their vectors. Since McLain et al. (2001) did not include an analysis of ticks from the central part of the range, it was decided to fill this gap and determine the genetic structure of I. persulcatus primarily in the Middle Urals and Western Siberia and selectively in some regions of Russia. For more accurate analysis, a fragment of the
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
213
Fig. 1. The distribution range of the tick, Ixodes persulcatus (colored in gray). Collection sites are indicated by a black circle (䊉).
mitochondrial 12S rRNA gene, which has been widely used in population genetic studies of ticks (Beati and Keirans, 2001; Casati et al., 2008; Norris et al., 1996), was used as an additional marker. Our results do not support the finding of McLain et al. (2001) leading us to reconsider the existing view on the genetic structure and evolution of I. persulcatus. Materials and methods Ticks, altogether 77 (76 I. persulcatus and 1 I. pavlovskyi), were collected in various regions of Russia in 2007–2011. Sampling was carried out in such a way as to represent a large portion of the species distribution. Collection sites are indicated on the map (Fig. 1). Detailed information on the collection sites as well as GenBank accession numbers are presented in Table 1. Extraction of nucleic acids and reverse transcription Ticks were frozen in liquid nitrogen, crushed, and homogenized in physiological salt solution (0.154 M NaCl) in order to obtain a suspension. Nucleic acid was extracted using a Ribo-Sorb kit for RNA/DNA extraction (Interlabservis, Russia), and reverse transcription was performed using a Reverta kit (Interlabservis), both according to the manufacturer’s instructions. PCR and sequencing The sequences of primers used for amplification are given in previously published papers (12S rRNA, Beati and Keirans, 2001; 28S rRNA, McLain et al., 2001). The PCR reaction mixture (total volume 25 l) contained 3.0 mM MgCl2 , 0.4 mM dNTP, 0.4 M of each primer, 1 U DiaTaq polymerase (Interlabservis), 5.0 l cDNA
template. PCR amplifications were performed using a VeritiTM Thermal Cycler (Applied Biosystems, USA) with the following conditions: 94 ◦ C for 2 min; 42 cycles: melting 94 ◦ C 10 s, annealing of primers 59 ◦ C 10 s; synthesis of DNA 72 ◦ C 15 s, and 72 ◦ C 3 min. Hot start (wax barrier method) was used to improve the specificity of PCR. However, in some cases, this method was shown to be inappropriate for our purposes (see ‘Results’). PCR products were separated by electrophoresis in 2% agarose gel, stained with ethidium bromide and visualized with a UV transilluminator. The DNA sequencing was performed on a Genetic Analyzer ABI PRIZM® 310 (Applied Biosystems, USA) using the reagent kit, BigDye Terminator v.3.1., according to the manufacturer’s instructions. Phylogenetic analysis Analysis, alignment of sequences and reconstruction of phylogenetic trees were performed using MEGA 5.05 (Tamura et al., 2011). Phylogenetic trees were constructed using Neighbor-joining algorithm (all ambiguous positions were removed for each sequence pair), with bootstrapping over 500 pseudo-replicates. Secondary structure prediction The secondary structure of the 28S rRNA D3 segment was predicted for the sequence of I. persulcatus obtained in our study as well as for sequences from GenBank (I. persulcatus AF303997, Cordyceps scarabaeicola AF339524, Mus musculus NR 003279). The secondary structure prediction was carried out using the program, Mfold (http://mobyle.pasteur.fr/cgi-bin/portal.py?form=mfold), based on a minimum free energy algorithm (Zuker et al., 1999). The resulting structures were visualized using RnaViz (De Rijk et al., 2003).
214
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Table 1 Origin and sources of ticks used for PCR amplification and GenBank IDs for associated DNA sequences. Locality
Ixodes persulcatus Altai, Southern Siberia Altai, Southern Siberia Irkutsk, Eastern Siberia Khanty-Mansiysk, Western Siberia Kirov, European part Krasnoufimsk, Middle Ural Kurgan, Western Siberia Omsk, Western Siberia Saint-Petersburg, Northwest of European part Sakhalin, Far East Serov, Middle Ural Tavda, Western Siberia Tavda, Western Siberia Tavda, Western Siberia Tyumen, Western Siberia Ufa, Southern Ural Vladivostok, Far East Yekaterinburg, Middle Ural Ixodes pavlovskyi Vladivostok, Far East
28S rRNA
12S rRNA
GenBank ID
Number, sex, dev. stagea
GenBank ID
Number, sex, dev. stagea
Haplotype
JQ085390 JQ085390 HM234635 HM234639 JQ085389 ND HM234636 HM234637 JF357623 HM234638 ND HM234633 ND ND HM234640 HM234632 JQ085391 HM234634
1 m, adult 1 f, adult 2 f, adult 2 f, adult 2 f, adult ND 2 f, adult 1 f, adult 2, adult 1 f, 2 nymphs ND 3 f, adult ND ND 1 f, 1 m, adult 1 f, adult 2 f, adult 1 f, adult
JQ085385 JQ085386 HM234626 HM234630 JQ085384 HM234620 HM234627 HM234628 JF412293 HM234629 HM234621 HM234624 HM234623 HM234622 HM234631 HM234619 JQ085387 HM234625
1 m, adult 1 f, adult 2 f, adult 2 f, adult 2 f, adult 2 f, adult 2 f, adult 2f, adult 7, adult 3 f, adult 2 f, adult 1 m, adult 6 f, 4 m, adult 1 f, adult 2 f,1 m, adult 3 f, adult 2 f, adult 21 f, 8 m, adult, 1 nymph
1 4 1 1 1 1 1 1 1 1 1 3 1 2 1 1 1 1
JQ085392
1 f, adult
JQ085388
1 f, adult
f, female; m, male. a The total number of ticks from one locality and with identical sequences is indicated.
For comparison, the elements of the secondary structure model are numbered according to the model of McLain (2001).
Results Sequence polymorphism of the 28S and 12S fragments The sequences of the gene fragments of 28S (340 bp) and 12S rRNA (351 bp) were obtained for 25 and 76 ticks I. persulcatus, respectively. For the first time, we have determined the sequences of these fragments of I. pavlovskyi Pomerantzev, 1946. GenBank accession numbers are listed in Table 1. It was shown, that the sequence of D3 segment of I. persulcatus is almost identical to those of other Ixodes species, particularly to I. ricinus (FR874103), I. scapularis (FR874102), and I. pavlovskyi (JQ085392), differing from them by not more than 2 nucleotides (Fig. 2A). Moreover, the complete sequence identity of the D3 expansion segment of 28S rRNA in I. persulcatus individuals from all the studied areas was observed. The number of variable nucleotides for the 12S rRNA gene fragment was 1.14% (indels are ignored). The analysis of this marker revealed 4 haplotypes (Fig. 2B), one of which (haplotype 1) occurred at a frequency of 0.96 in the whole studied range from the European part of Russia to the Far East (Table 1). Two haplotypes (2 and 3, HM234624 and HM234622) were found near Tavda (Western Siberia), each in a single case, and differed by only one nucleotide from the haplotype 1. The fourth haplotype (JQ085386), which also had one substitution, was found in the Altai Mountains (Southern Siberia) (Fig. 1). The analysis of 2 available GenBank sequences of the selected region, AB073725 (from Japan) and JF758624 (from China), revealed minor changes (one insertion with one substitution and 2 deletions compared to the haplotype 1, respectively) (Fig. 2B). Using the sequences obtained and related ones for ticks of the I. ricinus complex obtained from GenBank, we constructed phylogenetic trees showing the evolutionary position of I. persulcatus (Fig. 2A and B). No sequence from the McLain study (AF303986–AF303999) was used because of their major differences with our sequences and other ones recently obtained (Anstead et al., 2011). Because of the different number of available sequences
of 28S and 12S rRNA, the comparison of 2 phylogenetic trees would be inappropriate here. The influence of hot start method on PCR results We have shown that the use of hot start (wax barrier method) significantly affected the yield of PCR product (Fig. 4). Since the difference in the length of the D3 fragments between animals and fungi is 50–60 nucleotides, preferential amplification can be easily observed as fragments of different length on agarose gel. Amplified fragments were sequenced and found to belong to different organisms (see “Deep’ genetic structure of I. persulcatus’ for discussion). 28S rRNA D3 structure modeling For comparative analysis, a model of the secondary structure of the D3 segment of I. persulcatus based on our data and on McLain’s sequences was constructed, as well as analogous models for the ascomycete Cordyceps scarabaeicola (whose D3 sequence is almost identical to McLain’s sequences of I. persulcatus AF303992, AF303993 – see “Deep’ genetic structure of I. persulcatus’ for details) and the mouse as a distantly related animal (Fig. 3). The predicted structures correspond to the common model of the D3 segment of eukaryotes (Michot et al., 1984). Remarkably, the secondary structure of our D3 segment sequence of I. persulcatus has an H14 stem and 4 loops (S1–S4), which corresponds to the model of the D3 segment of other ticks, members of the I. ricinus complex (McLain, 2001), and even vertebrate animals (mouse). At the same time, in agreement with McLain’s result, the D3 segment of I. persulcatus has a deletion of 60 nucleotides and lacks the S4 stem, showing that its secondary structure is closer to those of ascomycete fungi, which also have 3 loops (S1–S3; Fig. 3) (Michot et al., 1984). Discussion Intraspecific variability The wide geographic range of I. persulcatus (from the Baltic to the Far East), along with the low mobility of tick individuals,
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
215
Fig. 2. Phylogenetic position of I. persulcatus among species of the I. ricinus complex (Dermacentor albipictus is used as an outgroup). Phylogenetic trees were constructed using Neighbor-joining algorithm (all ambiguous positions were removed for each sequence pair) with bootstrapping over 500 pseudo-replicates (values below 50 are not presented). Alignments of variable nucleotide positions are shown in the right of the trees (the position numbers are given according to the sequences with HM234634 for 28S and HM234619 for 12S rRNA). (A) A tree constructed on the basis of the D3 segment of the 28S rRNA contains all available D3 sequences for Ixodes ticks. (B) A tree constructed on the basis of the fragment of the 12S rRNA. The sequences obtained in this study are marked in bold; the numbers correspond to the haplotypes (details are given in the text).
suggests a marked isolation of populations, which could be observed at the genetic level. However, our studies have shown that some genetic markers, used for studying population diversity in other Ixodes species, did not have sufficient resolution for I. persulcatus ticks. Thus, the levels of intraspecific variability can be compared by using the number of variable nucleotides for each species. European ticks, I. ricinus L., which have an overlapping area with I. persulcatus, are well studied and show a low degree of genetic diversity based on different markers (Casati et al., 2008; Delaye et al., 1997). Nevertheless, the number of variable sites in the fragment of 12S rRNA was 2.5% for I. ricinus that is twofold greater compared to 1.14% for I. persulcatus (Casati et al., 2008). Populations of I. scapularis Say (U.S., East Coast) were shown to be divided into 2 well-isolated groups, northern and southern, with 12.6% of variable sites (Norris et al., 1996). However, our data show an extremely low intraspecific variability as well as the lack of a deep phylogeographical population structure of I. persulcatus in Russia. Moreover, mutations in 12S rRNA which give 4 haplotypes seem to be random and do not reflect
the structure of tick populations. Further research is needed to confirm or disprove this assumption. Our results are indirectly supported by data on the low variability of the internal transcribed spacer 2 (ITS2) within the population of I. persulcatus in Japan (Fukunaga et al., 2000). At the same time, the available data suggest a higher rate of mutation of noncoding ITS compared to 28S rRNA expansion segments that usually show no intraspecific variation (Anstead et al., 2011; Bae et al., 2008). It should be noted that, according to Fukunaga, ticks of the I. ricinus complex (including I. persulcatus) cluster tightly and form a lineage distinct from the other species (Fukunaga et al., 2000). The probable reason for the genetic homogeneity of I. persulcatus in the whole range is not the recent origin of the species, but may reflect the impact of global climate changes in the past, which has been shown for other tick species (Casati et al., 2008; Kain et al., 1999; Qiu et al., 2002). Generally, the consequence of glaciation events is the extinction and the loss of genetic variation in living populations in the affected regions (Hewitt, 2004). In such conditions, many forest species were forced to retreat to the mountains,
216
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Fig. 3. Models of the secondary structure of the D3 segment 28S rRNA of I. persulcatus, the ascomycete Cordyceps scarabaeicola, and the mouse Mus musculus.
where they formed refuges. Apparently, such extinction and refuge formation in the last Pleistocene glaciation, which finished approximately 10,000 years ago, caused the lack of genetic diversity in I. persulcatus at the level of rRNA genes. Consequently, the phylogeographical structure of I. persulcatus could be revealed using more variable genetic markers. ‘Deep’ genetic structure of I. persulcatus Our study of 28S rRNA of I. persulcatus, being actually a reproduction of McLain et al. (2001), led to different results and did not support his conclusions. Thus, McLain found that I. persulcatus from different localities showed high-sequence variation in the D3 segment comparable to those observed between all other Ixodes species. Thus, the population structure would be deep. Furthermore, the analysis of sequences of I. persulcatus revealed
shared deletions of approximately 60 nucleotides. According to McLain, sequence identity among the observed I. ricinus complex species is 60% when I. persulcatus is excluded and only 12% when I. persulcatus is included (McLain et al., 2001). The trees obtained in the present study both for 28S and 12S rRNA show a high degree of similarity within the I. ricinus complex including I. persulcatus (Fig. 2), even if there are some difficulties in interpreting data on the phylogeny of ticks of the I. ricinus complex (Foley et al., 2008). Despite the absence of morphological and genetic evidence of such a great interpopulation variability of I. persulcatus, McLain’s paper suggests an inaccurate concept about the evolution of the species. Moreover, the considerable variability of the D3 segment at the intraspecific level has been cited in some papers reviewing methods of studying genetic structure and phylogeography (Dabert, 2006; Nava et al., 2009).
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Fig. 4. The influence of hot start on the yield of PCR product for 2 selected samples of ticks A and B (see further description in the text). The numbered fragments are D3 segments 28S rRNA of: 1, I. persulcatus (deleterious sequence); 2, fungus; 3, unknown organism; 4, I. persulcatus (target sequence). L-DNA ladder (100–1000 bp).
We have tried to uncover the cause of such great differences between previous results and our data and have concluded that McLain’s results could be explained by the universality of the primers used and an insufficient analysis of the data. Indeed, primers designed on the basis of the complete sequence of 28S rRNA Drosophila melanogaster are highly conserved and may amplify fragments of 28S rRNA of a wide range of eukaryotes (parasites, saprophytes, symbionts of ticks, ticks as well as their hosts). Other authors have also mentioned the problems encountered while working with highly conserved primers in the presence of DNA from more than one species in the sample (Buckler et al., 1997; Navajas and Fenton, 2000; Yli-Mattila et al., 2000). The lack of the GenBank nucleotide sequence for the 28S rRNA D3 segment of ticks and fungi (at the time McLain was carrying out his study) could contribute to an erroneous determination of the source of fragments obtained. With the BLAST program (http://blast.ncbi.nlm.nih.gov/Blast.cgi), we found 2 of McLain’s sequences of I. persulcatus D3 segments (AF303992, AF303993) to be identical with the rRNA of entomopathogenic fungi like Beauveria spp. (e.g. B. caledonica AF339520) and Cordyceps spp. (C. scarabaeicola AF339524) (except for 3 indels). The presence of fungi in the samples is easily explained by tick ecology and the necessity of their obligatory contact with soil and vegetation. The soil and entomopathogenic fungi can colonize the surface of the tick (or their eggs) during their lifetime. Thus, DNA of fungi has been repeatedly isolated from suspensions of ticks (Tuininga et al., 2009). Additionally, 2 Ixodes species (I. kingi and I. sculptus) have been found to harbor fungi, whose 28S rRNA gene fragment was amplified with the primers used by McLain (Anstead et al., 2011). In the case of amplification of rRNA fragments, it is important that these molecules have a rather stable secondary structure. It was shown that the presence of natural contamination of the samples (by fungi, algae) led to possible preferential amplification depending on the PCR conditions (Buckler et al., 1997). We have shown that the use of hot start (wax barrier method) significantly affected the yield of PCR product (Fig. 4). When carrying out PCR with hot start for 2 selected samples, we have not found the target 387-bp fragment (Fig. 4). Thus, the 205-bp fragment (Fig. 4, 1) represents a deleterious sequence of 28S rRNA of I. persulcatus, which probably is a pseudogene or formed during PCR by the phenomenon of jumping PCR (Paabo et al., 1990). 340-bp fragments (Fig. 4, 2) correspond to the rRNA of fungi. Using the BLAST program, we
217
found 100% similarity of one of our sequences with the fungus Cladosporium sphaerospermum (GU017542). In addition, we detected a fragment which was longer than the target by 10 nucleotides (Fig. 4, 3). Unfortunately, we were unable to determine the source of this sequence because of low coverage and identity with database sequences. Without hot start, the balance shifts towards amplification of the rRNA of I. persulcatus ticks (Fig. 4, 4). Hence, under certain PCR conditions, we can observe fragments of a wide range of organisms, but not the required one, which was possibly the case with McLain’s I. persulcatus sequences. Modeling of the secondary structure of the D3 segment, the object of analysis in McLain’s second paper (McLain, 2001), confirms our hypothesis. Many authors indicate the highly conservative secondary structure of rRNA, including the expansion segments (Hancock and Dover, 1990; Michot et al., 1984). At the same time, according to McLain, the I. persulcatus D3 segment differs from those of other species even by the number of stems (3 compared to 4 for other Ixodes species). Therefore, the secondary structure of the I. persulcatus D3 segment, predicted by McLain, is closer to those of fungi rather than to those of other ticks (Fig. 3). All these data prove the hypothesis that McLain’s assumptions about the significant interpopulation variability and deep genetic structure of I. persulcatus ticks are based on the analysis of fungal sequences. In conclusion, the present study indicates that I. persulcatus ticks are not a separate and heterogeneous group. On the contrary, they are an essential part of the I. ricinus complex, as has been demonstrated also by other researchers (Fukunaga et al., 2000; Xu et al., 2003). Among all the ticks of the genus Ixodes studied to date, I. persulcatus shows the lowest genetic variability, which is probably associated with the strong influence of glaciation on its habitat. Acknowledgments We are indebted to O.L. Burundukova, A.V. Ershov, E.G. Filippov, S.I. Ibragimov, N.V. Loginovskikh, M.V. Modorov, T.A. Pimenova, T.E. Snitkovskaya, and N.K. Tokarevich for the collection of material, and to Dr Keith Chamberlain (Rothamsted Research) for his help in preparing the manuscript. The Russian Foundation of Basic Research (No. 10-04-96062) supported this project. References Alekseev, A.N., Jensen, P.M., Dubinina, H.V., Smirnova, L.A., Makrouchina, N.A., Zharkov, S.D., 2000. Peculiarities of behaviour of taiga (Ixodes persulcatus) and sheep (Ixodes ricinus) ticks (Acarina: Ixodidae) determined by different methods. Folia Parasitol. (Praha) 47, 147–153. Anstead, C.A., Krakowetz, C.N., Mann, A.S., Sim, K.A., Chilton, N.B., 2011. An assessment of genetic differences among ixodid ticks in a locus within the nuclear large subunit ribosomal RNA gene. Mol. Cell. Probes 25, 243–248. Bae, C., Szalanski, A., Robbins, R., 2008. Molecular analysis of the lance nematode, Hoplolaimus spp., using the first internal transcribed spacer and the D1–D3 expansion segments of 28S ribosomal DNA1. J. Nematol. 40, 201–209. Beati, L., Keirans, J.E., 2001. Analysis of the systematic relationships among ticks of the genera Rhipicephalus and Boophilus (Acari: Ixodidae) based on mitochondrial 12S ribosomal DNA gene sequences and morphological characters. J. Parasitol. 87, 32–48. Buckler, E.S.T., Ippolito, A., Holtsford, T.P., 1997. The evolution of ribosomal DNA: divergent paralogues and phylogenetic implications. Genetics 145, 821–832. Casati, S., Bernasconi, M.V., Gern, L., Piffaretti, J.C., 2008. Assessment of intraspecific mtDNA variability of European Ixodes ricinus sensu stricto (Acari: Ixodidae). Infect. Genet. Evol. 8, 152–158. Dabert, M., 2006. DNA markers in the phylogenetics of the Acari. Biol. Lett. 43, 97–107. De Rijk, P., Wuyts, J., De Wachter, R., 2003. RnaViz 2: an improved representation of RNA secondary structure. Bioinformatics 19, 299–300. Delaye, C., Beati, L., Aeschlimann, A., Renaud, F., de Meeus, T., 1997. Population genetic structure of Ixodes ricinus in Switzerland from allozymic data: no evidence of divergence between nearby sites. Int. J. Parasitol. 27, 769–773. Ecker, M., Allison, S.L., Meixner, T., Heinz, F.X., 1999. Sequence analysis and genetic classification of tick-borne encephalitis viruses from Europe and Asia. J. Gen. Virol. 80 (Pt 1), 179–185.
218
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Filippova, N.A., 1985. Taiga Tick Ixodes persulcatus Schulze (Acarina, Ixodidae). Morphology, Systematics, Ecology, Medical Importance. Nauka Publishers, Leningrad, p. 416 (in Russian). Filippova, N.A., 2002. Morphological barrier in mechanisms of reproductive isolation acting in areas of sympatry of closely related species Ixodes persulcatus–I. pavlovskyi and I. persulcatus–I. ricinus (Ixodidae). Parazitologiia 36, 457–468 (in Russian). Filippova, N.A., Musatov, S.A., 1996. Geographic variability in the sexually mature phase of Ixodes persulcatus (Ixodidae). Experience in using databases on morphometry. Parazitologiia 30, 205–215 (in Russian). Foley, J., Nieto, N.C., Foley, P., Teglas, M.B., 2008. Co-phylogenetic analysis of Anaplasma phagocytophilum and its vectors, Ixodes spp. ticks. Exp. Appl. Acarol. 45, 155–170. Fomenko, N.V., Stronin, O.V., Khasnatinov, M.A., Danchinova, G.A., Bataa, J., Gol’tsova, N.A., 2009. The heterogeneity of the ospa gene of Borrelia garinii and Borrelia afzelii in western Siberia and Mongolia. Mol. Gen. Mikrobiol. Virusol., 18–22. Fukunaga, M., Yabuki, M., Hamase, A., Oliver Jr., J.H., Nakao, M., 2000. Molecular phylogenetic analysis of ixodid ticks based on the ribosomal DNA spacer, internal transcribed spacer 2, sequences. J. Parasitol. 86, 38–43. Hancock, J.M., Dover, G.A., 1990. ‘Compensatory slippage’ in the evolution of ribosomal RNA genes. Nucleic Acids Res. 18, 5949–5954. Hewitt, G.M., 2004. The structure of biodiversity – insights from molecular phylogeography. Front. Zool. 1, 4. Kain, D.E., Sperling, F.A., Daly, H.V., Lane, R.S., 1999. Mitochondrial DNA sequence variation in Ixodes pacificus (Acari: Ixodidae). Heredity 83 (Pt 4), 378–386. Kriuchechnikov, V.N., Korenberg, E.I., Shcherbakov, S.V., Kovalevskii, Iu.V., Levin, M.L., 1988. Identification of Borrelia isolated in the USSR from Ixodes persulcatus Schulze ticks. Zh. Mikrobiol. Epidemiol. Immunobiol., 41–44 (in Russian). McLain, D.K., 2001. Evolution of transcript structure and base composition of rDNA expansion segment D3 in ticks. Heredity 87, 544–557. McLain, D.K., Li, J., Oliver Jr., J.H., 2001. Interspecific and geographical variation in the sequence of rDNA expansion segment D3 of Ixodes ticks (Acari: Ixodidae). Heredity 86, 234–242. Michot, B., Hassouna, N., Bachellerie, J.P., 1984. Secondary structure of mouse 28S rRNA and general model for the folding of the large rRNA in eukaryotes. Nucleic Acids Res. 12, 4259–4279. Nava, S., Guglielmone, A.A., Mangold, A.J., 2009. An overview of systematics and evolution of ticks. Front. Biosci. 14, 2857–2877.
Navajas, M., Fenton, B., 2000. The application of molecular markers in the study of diversity in acarology: a review. Exp. Appl. Acarol. 24, 751–774. Nefedova, V.V., Korenberg, E.I., Gorelova, N.B., 2010. Genetic variants of Borrelia garinii, a widely spread Eurasian pathogen of ixodic tick borreliosis. Mol. Gen. Mikrobiol. Virusol., 7–12. Norris, D.E., Klompen, J.S., Keirans, J.E., Black IV, W.C., 1996. Population genetics of Ixodes scapularis (Acari: Ixodidae) based on mitochondrial 16S and 12S genes. J. Med. Entomol. 33, 78–89. Paabo, S., Irwin, D.M., Wilson, A.C., 1990. DNA damage promotes jumping between templates during enzymatic amplification. J. Biol. Chem. 265, 4718–4721. Qiu, W.G., Dykhuizen, D.E., Acosta, M.S., Luft, B.J., 2002. Geographic uniformity of the Lyme disease spirochete (Borrelia burgdorferi) and its shared history with tick vector (Ixodes scapularis) in the Northeastern United States. Genetics 160, 833–849. Shpynov, S.N., Rudakov, N.V., Iastrebov, V.K., Leonova, G.N., Khazova, T.G., Egorova, N.V., Borisova, O.N., Preider, V.P., Bezrukov, G.V., Fedorov, E.G., Fedianin, A.P., Sherstneva, M.B., Turyshev, A.G., Gavrilov, A.P., Tankibaev, M.A., Fournier, P.E., Raoult, D., 2004. New evidence for the detection of Ehrlichia and Anaplasma in Ixodes ticks in Russia and Kazakhstan. Med. Parazitol. (Mosk), 10–14 (in Russian). Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011. MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28, 2731–2739. Tuininga, A.R., Miller, J.L., Morath, S.U., Daniels, T.J., Falco, R.C., Marchese, M., Sahabi, S., Rosa, D., Stafford III, K.C., 2009. Isolation of entomopathogenic fungi from soils and Ixodes scapularis (Acari: Ixodidae) ticks: prevalence and methods. J. Med. Entomol. 46, 557–565. Xu, G., Fang, Q.Q., Keirans, J.E., Durden, L.A., 2003. Molecular phylogenetic analyses indicate that the Ixodes ricinus complex is a paraphyletic group. J. Parasitol. 89, 452–457. Yli-Mattila, T., Paavanen-Huhtala, S., Fenton, B., Tuovinen, T., 2000. Species and strain identification of the predatory mite Euseius finlandicus by RAPD-PCR and ITS sequences. Exp. Appl. Acarol. 24, 863–880. Zilber, L.A., 1939. Spring–summer tick-borne encephalitis. Arkhiv. Biol. Nauk. 56, 255–261. Zuker, M., Mathews, D.H., Turner, D.H., 1999. In: Clark, B.A.B.F.C. (Ed.), Algorithms and Thermodynamics for RNA Secondary Structure Prediction: A Practical Guide in RNA Biochemistry and Biotechnology. Kluwer Academic Publishers.
Contents lists available at SciVerse ScienceDirect
Ticks and Tick-borne Diseases journal homepage: www.elsevier.de/ttbdis
Original article
Phylogeographical structure of the tick Ixodes persulcatus: A novel view Sergey Y. Kovalev ∗ , Tatyana A. Mukhacheva Laboratory of Molecular Genetics, Department of Biology, Ural Federal University, Lenin Avenue 51, Yekaterinburg 620000, Russia
a r t i c l e
i n f o
Article history: Received 18 January 2012 Received in revised form 19 March 2012 Accepted 19 March 2012 Keywords: Ixodes persulcatus Phylogeography Preferential amplification rRNA Population structure
a b s t r a c t The tick Ixodes persulcatus Schulze, 1930, has a wide distribution from the Baltic to the Far East and is a vector of a number of human pathogens. Thus, the study of the genetic structure and evolution of this species is of great epidemiological importance. rRNA genes were used as genetic markers to identify the phylogeographical structure of the ticks. The sequences of gene fragments of 28S (expansion segment D3) and mitochondrial 12S rRNA for 25 and 76 ticks, respectively, that had been collected in various regions of Russia in 2007–2011, were obtained. The sequences of the 28S rRNA D3 segment were identical for all ticks within the studied area. Analysis of the sequences of the mitochondrial 12S rRNA fragment revealed 4 haplotypes with one occurring at a frequency of 0.96. It is shown that the ‘deep’ population structure of I. persulcatus (McLain et al., 2001) was erroneous because of the inclusion of contaminating fungi sequences of 28S rRNA in the phylogenetic analysis. This was, possibly, due to the use of universal PCR primers that amplify the DNA of a wide range of eukaryotes, particularly of fungi which are common in samples of ticks. The influence of PCR conditions on the preferential amplification of the DNA of different organisms is also demonstrated. © 2012 Elsevier GmbH. All rights reserved.
Introduction The tick Ixodes persulcatus Schulze, 1930, commonly known as the taiga tick, is a vector of a number of human pathogens including tick-borne encephalitis virus (Zilber, 1939), Borrelia burgdorferi s.l. (Kriuchechnikov et al., 1988), and Anaplasma phagocytophilum (Shpynov et al., 2004). This species belongs to the Ixodes ricinus complex – a group of ticks distributed in almost all geographic regions of the world (Xu et al., 2003). The tick I. persulcatus inhabits the taiga zone from the Baltic to the Russian Far East as well as Japan, northern China, Mongolia, and Kazakhstan (Fig. 1) (Filippova, 1985) and is the principal vector of tick-borne infections in Russia. Given such a wide range and epidemiological significance, the evolution of I. persulcatus is of great scientific interest. To date, a lot of information concerning the morphology, biology, and ecology of I. persulcatus has been acquired (Alekseev et al., 2000; Filippova, 1985, 2002; Filippova and Musatov, 1996). Moreover, some attempts to study the intraspecific structure of the taiga tick based on a comparison of morphological features were made, and some differences in body size between Asian and European populations were found (Filippova and Musatov, 1996). The study by Filippova (1985) highlights the difficulty of investigating the geographic variation in this species because of the wide geographic range and the inconsistency of data on the
∗ Corresponding author. Tel.: +7 343 2616826; fax: +7 343 3507401. E-mail address: [email protected] (S.Y. Kovalev). 1877-959X/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.ttbdis.2012.03.005
variability of morphological structures at different developmental stages of ticks (Filippova, 1985). Generally, morphological traits may have a complex genetic basis and are often dependent on environmental conditions. Therefore, genetic markers should be used for such population studies. They were shown to be useful in both systematic and population analyses of ticks (Nava et al., 2009). Several studies on ticks belonging to the Ixodes ricinus complex (Casati et al., 2008; Norris et al., 1996) allowed a comparative analysis of their genetic structure to be carried out and the processes governing tick distribution to be understood. However, little is known about the population structure of I. persulcatus. In this regard, special attention should be paid to the work of McLain et al. (2001) who suggested extreme heterogeneity and deep geographical structure of the I. persulcatus population, based on the nucleotide sequence of the expansion segment D3 28S rRNA, as well as a significant difference between the secondary structure of this segment compared to other ticks of the I. ricinus complex (McLain, 2001). Such a structure suggested a long and independent evolution of I. persulcatus populations. As pathogens transmitted by I. persulcatus also show heterogeneous genetic structure (Ecker et al., 1999; Fomenko et al., 2009; Nefedova et al., 2010), they can form an excellent basis for the study of coevolutionary processes in populations of pathogens and their vectors. Since McLain et al. (2001) did not include an analysis of ticks from the central part of the range, it was decided to fill this gap and determine the genetic structure of I. persulcatus primarily in the Middle Urals and Western Siberia and selectively in some regions of Russia. For more accurate analysis, a fragment of the
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
213
Fig. 1. The distribution range of the tick, Ixodes persulcatus (colored in gray). Collection sites are indicated by a black circle (䊉).
mitochondrial 12S rRNA gene, which has been widely used in population genetic studies of ticks (Beati and Keirans, 2001; Casati et al., 2008; Norris et al., 1996), was used as an additional marker. Our results do not support the finding of McLain et al. (2001) leading us to reconsider the existing view on the genetic structure and evolution of I. persulcatus. Materials and methods Ticks, altogether 77 (76 I. persulcatus and 1 I. pavlovskyi), were collected in various regions of Russia in 2007–2011. Sampling was carried out in such a way as to represent a large portion of the species distribution. Collection sites are indicated on the map (Fig. 1). Detailed information on the collection sites as well as GenBank accession numbers are presented in Table 1. Extraction of nucleic acids and reverse transcription Ticks were frozen in liquid nitrogen, crushed, and homogenized in physiological salt solution (0.154 M NaCl) in order to obtain a suspension. Nucleic acid was extracted using a Ribo-Sorb kit for RNA/DNA extraction (Interlabservis, Russia), and reverse transcription was performed using a Reverta kit (Interlabservis), both according to the manufacturer’s instructions. PCR and sequencing The sequences of primers used for amplification are given in previously published papers (12S rRNA, Beati and Keirans, 2001; 28S rRNA, McLain et al., 2001). The PCR reaction mixture (total volume 25 l) contained 3.0 mM MgCl2 , 0.4 mM dNTP, 0.4 M of each primer, 1 U DiaTaq polymerase (Interlabservis), 5.0 l cDNA
template. PCR amplifications were performed using a VeritiTM Thermal Cycler (Applied Biosystems, USA) with the following conditions: 94 ◦ C for 2 min; 42 cycles: melting 94 ◦ C 10 s, annealing of primers 59 ◦ C 10 s; synthesis of DNA 72 ◦ C 15 s, and 72 ◦ C 3 min. Hot start (wax barrier method) was used to improve the specificity of PCR. However, in some cases, this method was shown to be inappropriate for our purposes (see ‘Results’). PCR products were separated by electrophoresis in 2% agarose gel, stained with ethidium bromide and visualized with a UV transilluminator. The DNA sequencing was performed on a Genetic Analyzer ABI PRIZM® 310 (Applied Biosystems, USA) using the reagent kit, BigDye Terminator v.3.1., according to the manufacturer’s instructions. Phylogenetic analysis Analysis, alignment of sequences and reconstruction of phylogenetic trees were performed using MEGA 5.05 (Tamura et al., 2011). Phylogenetic trees were constructed using Neighbor-joining algorithm (all ambiguous positions were removed for each sequence pair), with bootstrapping over 500 pseudo-replicates. Secondary structure prediction The secondary structure of the 28S rRNA D3 segment was predicted for the sequence of I. persulcatus obtained in our study as well as for sequences from GenBank (I. persulcatus AF303997, Cordyceps scarabaeicola AF339524, Mus musculus NR 003279). The secondary structure prediction was carried out using the program, Mfold (http://mobyle.pasteur.fr/cgi-bin/portal.py?form=mfold), based on a minimum free energy algorithm (Zuker et al., 1999). The resulting structures were visualized using RnaViz (De Rijk et al., 2003).
214
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Table 1 Origin and sources of ticks used for PCR amplification and GenBank IDs for associated DNA sequences. Locality
Ixodes persulcatus Altai, Southern Siberia Altai, Southern Siberia Irkutsk, Eastern Siberia Khanty-Mansiysk, Western Siberia Kirov, European part Krasnoufimsk, Middle Ural Kurgan, Western Siberia Omsk, Western Siberia Saint-Petersburg, Northwest of European part Sakhalin, Far East Serov, Middle Ural Tavda, Western Siberia Tavda, Western Siberia Tavda, Western Siberia Tyumen, Western Siberia Ufa, Southern Ural Vladivostok, Far East Yekaterinburg, Middle Ural Ixodes pavlovskyi Vladivostok, Far East
28S rRNA
12S rRNA
GenBank ID
Number, sex, dev. stagea
GenBank ID
Number, sex, dev. stagea
Haplotype
JQ085390 JQ085390 HM234635 HM234639 JQ085389 ND HM234636 HM234637 JF357623 HM234638 ND HM234633 ND ND HM234640 HM234632 JQ085391 HM234634
1 m, adult 1 f, adult 2 f, adult 2 f, adult 2 f, adult ND 2 f, adult 1 f, adult 2, adult 1 f, 2 nymphs ND 3 f, adult ND ND 1 f, 1 m, adult 1 f, adult 2 f, adult 1 f, adult
JQ085385 JQ085386 HM234626 HM234630 JQ085384 HM234620 HM234627 HM234628 JF412293 HM234629 HM234621 HM234624 HM234623 HM234622 HM234631 HM234619 JQ085387 HM234625
1 m, adult 1 f, adult 2 f, adult 2 f, adult 2 f, adult 2 f, adult 2 f, adult 2f, adult 7, adult 3 f, adult 2 f, adult 1 m, adult 6 f, 4 m, adult 1 f, adult 2 f,1 m, adult 3 f, adult 2 f, adult 21 f, 8 m, adult, 1 nymph
1 4 1 1 1 1 1 1 1 1 1 3 1 2 1 1 1 1
JQ085392
1 f, adult
JQ085388
1 f, adult
f, female; m, male. a The total number of ticks from one locality and with identical sequences is indicated.
For comparison, the elements of the secondary structure model are numbered according to the model of McLain (2001).
Results Sequence polymorphism of the 28S and 12S fragments The sequences of the gene fragments of 28S (340 bp) and 12S rRNA (351 bp) were obtained for 25 and 76 ticks I. persulcatus, respectively. For the first time, we have determined the sequences of these fragments of I. pavlovskyi Pomerantzev, 1946. GenBank accession numbers are listed in Table 1. It was shown, that the sequence of D3 segment of I. persulcatus is almost identical to those of other Ixodes species, particularly to I. ricinus (FR874103), I. scapularis (FR874102), and I. pavlovskyi (JQ085392), differing from them by not more than 2 nucleotides (Fig. 2A). Moreover, the complete sequence identity of the D3 expansion segment of 28S rRNA in I. persulcatus individuals from all the studied areas was observed. The number of variable nucleotides for the 12S rRNA gene fragment was 1.14% (indels are ignored). The analysis of this marker revealed 4 haplotypes (Fig. 2B), one of which (haplotype 1) occurred at a frequency of 0.96 in the whole studied range from the European part of Russia to the Far East (Table 1). Two haplotypes (2 and 3, HM234624 and HM234622) were found near Tavda (Western Siberia), each in a single case, and differed by only one nucleotide from the haplotype 1. The fourth haplotype (JQ085386), which also had one substitution, was found in the Altai Mountains (Southern Siberia) (Fig. 1). The analysis of 2 available GenBank sequences of the selected region, AB073725 (from Japan) and JF758624 (from China), revealed minor changes (one insertion with one substitution and 2 deletions compared to the haplotype 1, respectively) (Fig. 2B). Using the sequences obtained and related ones for ticks of the I. ricinus complex obtained from GenBank, we constructed phylogenetic trees showing the evolutionary position of I. persulcatus (Fig. 2A and B). No sequence from the McLain study (AF303986–AF303999) was used because of their major differences with our sequences and other ones recently obtained (Anstead et al., 2011). Because of the different number of available sequences
of 28S and 12S rRNA, the comparison of 2 phylogenetic trees would be inappropriate here. The influence of hot start method on PCR results We have shown that the use of hot start (wax barrier method) significantly affected the yield of PCR product (Fig. 4). Since the difference in the length of the D3 fragments between animals and fungi is 50–60 nucleotides, preferential amplification can be easily observed as fragments of different length on agarose gel. Amplified fragments were sequenced and found to belong to different organisms (see “Deep’ genetic structure of I. persulcatus’ for discussion). 28S rRNA D3 structure modeling For comparative analysis, a model of the secondary structure of the D3 segment of I. persulcatus based on our data and on McLain’s sequences was constructed, as well as analogous models for the ascomycete Cordyceps scarabaeicola (whose D3 sequence is almost identical to McLain’s sequences of I. persulcatus AF303992, AF303993 – see “Deep’ genetic structure of I. persulcatus’ for details) and the mouse as a distantly related animal (Fig. 3). The predicted structures correspond to the common model of the D3 segment of eukaryotes (Michot et al., 1984). Remarkably, the secondary structure of our D3 segment sequence of I. persulcatus has an H14 stem and 4 loops (S1–S4), which corresponds to the model of the D3 segment of other ticks, members of the I. ricinus complex (McLain, 2001), and even vertebrate animals (mouse). At the same time, in agreement with McLain’s result, the D3 segment of I. persulcatus has a deletion of 60 nucleotides and lacks the S4 stem, showing that its secondary structure is closer to those of ascomycete fungi, which also have 3 loops (S1–S3; Fig. 3) (Michot et al., 1984). Discussion Intraspecific variability The wide geographic range of I. persulcatus (from the Baltic to the Far East), along with the low mobility of tick individuals,
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
215
Fig. 2. Phylogenetic position of I. persulcatus among species of the I. ricinus complex (Dermacentor albipictus is used as an outgroup). Phylogenetic trees were constructed using Neighbor-joining algorithm (all ambiguous positions were removed for each sequence pair) with bootstrapping over 500 pseudo-replicates (values below 50 are not presented). Alignments of variable nucleotide positions are shown in the right of the trees (the position numbers are given according to the sequences with HM234634 for 28S and HM234619 for 12S rRNA). (A) A tree constructed on the basis of the D3 segment of the 28S rRNA contains all available D3 sequences for Ixodes ticks. (B) A tree constructed on the basis of the fragment of the 12S rRNA. The sequences obtained in this study are marked in bold; the numbers correspond to the haplotypes (details are given in the text).
suggests a marked isolation of populations, which could be observed at the genetic level. However, our studies have shown that some genetic markers, used for studying population diversity in other Ixodes species, did not have sufficient resolution for I. persulcatus ticks. Thus, the levels of intraspecific variability can be compared by using the number of variable nucleotides for each species. European ticks, I. ricinus L., which have an overlapping area with I. persulcatus, are well studied and show a low degree of genetic diversity based on different markers (Casati et al., 2008; Delaye et al., 1997). Nevertheless, the number of variable sites in the fragment of 12S rRNA was 2.5% for I. ricinus that is twofold greater compared to 1.14% for I. persulcatus (Casati et al., 2008). Populations of I. scapularis Say (U.S., East Coast) were shown to be divided into 2 well-isolated groups, northern and southern, with 12.6% of variable sites (Norris et al., 1996). However, our data show an extremely low intraspecific variability as well as the lack of a deep phylogeographical population structure of I. persulcatus in Russia. Moreover, mutations in 12S rRNA which give 4 haplotypes seem to be random and do not reflect
the structure of tick populations. Further research is needed to confirm or disprove this assumption. Our results are indirectly supported by data on the low variability of the internal transcribed spacer 2 (ITS2) within the population of I. persulcatus in Japan (Fukunaga et al., 2000). At the same time, the available data suggest a higher rate of mutation of noncoding ITS compared to 28S rRNA expansion segments that usually show no intraspecific variation (Anstead et al., 2011; Bae et al., 2008). It should be noted that, according to Fukunaga, ticks of the I. ricinus complex (including I. persulcatus) cluster tightly and form a lineage distinct from the other species (Fukunaga et al., 2000). The probable reason for the genetic homogeneity of I. persulcatus in the whole range is not the recent origin of the species, but may reflect the impact of global climate changes in the past, which has been shown for other tick species (Casati et al., 2008; Kain et al., 1999; Qiu et al., 2002). Generally, the consequence of glaciation events is the extinction and the loss of genetic variation in living populations in the affected regions (Hewitt, 2004). In such conditions, many forest species were forced to retreat to the mountains,
216
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Fig. 3. Models of the secondary structure of the D3 segment 28S rRNA of I. persulcatus, the ascomycete Cordyceps scarabaeicola, and the mouse Mus musculus.
where they formed refuges. Apparently, such extinction and refuge formation in the last Pleistocene glaciation, which finished approximately 10,000 years ago, caused the lack of genetic diversity in I. persulcatus at the level of rRNA genes. Consequently, the phylogeographical structure of I. persulcatus could be revealed using more variable genetic markers. ‘Deep’ genetic structure of I. persulcatus Our study of 28S rRNA of I. persulcatus, being actually a reproduction of McLain et al. (2001), led to different results and did not support his conclusions. Thus, McLain found that I. persulcatus from different localities showed high-sequence variation in the D3 segment comparable to those observed between all other Ixodes species. Thus, the population structure would be deep. Furthermore, the analysis of sequences of I. persulcatus revealed
shared deletions of approximately 60 nucleotides. According to McLain, sequence identity among the observed I. ricinus complex species is 60% when I. persulcatus is excluded and only 12% when I. persulcatus is included (McLain et al., 2001). The trees obtained in the present study both for 28S and 12S rRNA show a high degree of similarity within the I. ricinus complex including I. persulcatus (Fig. 2), even if there are some difficulties in interpreting data on the phylogeny of ticks of the I. ricinus complex (Foley et al., 2008). Despite the absence of morphological and genetic evidence of such a great interpopulation variability of I. persulcatus, McLain’s paper suggests an inaccurate concept about the evolution of the species. Moreover, the considerable variability of the D3 segment at the intraspecific level has been cited in some papers reviewing methods of studying genetic structure and phylogeography (Dabert, 2006; Nava et al., 2009).
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Fig. 4. The influence of hot start on the yield of PCR product for 2 selected samples of ticks A and B (see further description in the text). The numbered fragments are D3 segments 28S rRNA of: 1, I. persulcatus (deleterious sequence); 2, fungus; 3, unknown organism; 4, I. persulcatus (target sequence). L-DNA ladder (100–1000 bp).
We have tried to uncover the cause of such great differences between previous results and our data and have concluded that McLain’s results could be explained by the universality of the primers used and an insufficient analysis of the data. Indeed, primers designed on the basis of the complete sequence of 28S rRNA Drosophila melanogaster are highly conserved and may amplify fragments of 28S rRNA of a wide range of eukaryotes (parasites, saprophytes, symbionts of ticks, ticks as well as their hosts). Other authors have also mentioned the problems encountered while working with highly conserved primers in the presence of DNA from more than one species in the sample (Buckler et al., 1997; Navajas and Fenton, 2000; Yli-Mattila et al., 2000). The lack of the GenBank nucleotide sequence for the 28S rRNA D3 segment of ticks and fungi (at the time McLain was carrying out his study) could contribute to an erroneous determination of the source of fragments obtained. With the BLAST program (http://blast.ncbi.nlm.nih.gov/Blast.cgi), we found 2 of McLain’s sequences of I. persulcatus D3 segments (AF303992, AF303993) to be identical with the rRNA of entomopathogenic fungi like Beauveria spp. (e.g. B. caledonica AF339520) and Cordyceps spp. (C. scarabaeicola AF339524) (except for 3 indels). The presence of fungi in the samples is easily explained by tick ecology and the necessity of their obligatory contact with soil and vegetation. The soil and entomopathogenic fungi can colonize the surface of the tick (or their eggs) during their lifetime. Thus, DNA of fungi has been repeatedly isolated from suspensions of ticks (Tuininga et al., 2009). Additionally, 2 Ixodes species (I. kingi and I. sculptus) have been found to harbor fungi, whose 28S rRNA gene fragment was amplified with the primers used by McLain (Anstead et al., 2011). In the case of amplification of rRNA fragments, it is important that these molecules have a rather stable secondary structure. It was shown that the presence of natural contamination of the samples (by fungi, algae) led to possible preferential amplification depending on the PCR conditions (Buckler et al., 1997). We have shown that the use of hot start (wax barrier method) significantly affected the yield of PCR product (Fig. 4). When carrying out PCR with hot start for 2 selected samples, we have not found the target 387-bp fragment (Fig. 4). Thus, the 205-bp fragment (Fig. 4, 1) represents a deleterious sequence of 28S rRNA of I. persulcatus, which probably is a pseudogene or formed during PCR by the phenomenon of jumping PCR (Paabo et al., 1990). 340-bp fragments (Fig. 4, 2) correspond to the rRNA of fungi. Using the BLAST program, we
217
found 100% similarity of one of our sequences with the fungus Cladosporium sphaerospermum (GU017542). In addition, we detected a fragment which was longer than the target by 10 nucleotides (Fig. 4, 3). Unfortunately, we were unable to determine the source of this sequence because of low coverage and identity with database sequences. Without hot start, the balance shifts towards amplification of the rRNA of I. persulcatus ticks (Fig. 4, 4). Hence, under certain PCR conditions, we can observe fragments of a wide range of organisms, but not the required one, which was possibly the case with McLain’s I. persulcatus sequences. Modeling of the secondary structure of the D3 segment, the object of analysis in McLain’s second paper (McLain, 2001), confirms our hypothesis. Many authors indicate the highly conservative secondary structure of rRNA, including the expansion segments (Hancock and Dover, 1990; Michot et al., 1984). At the same time, according to McLain, the I. persulcatus D3 segment differs from those of other species even by the number of stems (3 compared to 4 for other Ixodes species). Therefore, the secondary structure of the I. persulcatus D3 segment, predicted by McLain, is closer to those of fungi rather than to those of other ticks (Fig. 3). All these data prove the hypothesis that McLain’s assumptions about the significant interpopulation variability and deep genetic structure of I. persulcatus ticks are based on the analysis of fungal sequences. In conclusion, the present study indicates that I. persulcatus ticks are not a separate and heterogeneous group. On the contrary, they are an essential part of the I. ricinus complex, as has been demonstrated also by other researchers (Fukunaga et al., 2000; Xu et al., 2003). Among all the ticks of the genus Ixodes studied to date, I. persulcatus shows the lowest genetic variability, which is probably associated with the strong influence of glaciation on its habitat. Acknowledgments We are indebted to O.L. Burundukova, A.V. Ershov, E.G. Filippov, S.I. Ibragimov, N.V. Loginovskikh, M.V. Modorov, T.A. Pimenova, T.E. Snitkovskaya, and N.K. Tokarevich for the collection of material, and to Dr Keith Chamberlain (Rothamsted Research) for his help in preparing the manuscript. The Russian Foundation of Basic Research (No. 10-04-96062) supported this project. References Alekseev, A.N., Jensen, P.M., Dubinina, H.V., Smirnova, L.A., Makrouchina, N.A., Zharkov, S.D., 2000. Peculiarities of behaviour of taiga (Ixodes persulcatus) and sheep (Ixodes ricinus) ticks (Acarina: Ixodidae) determined by different methods. Folia Parasitol. (Praha) 47, 147–153. Anstead, C.A., Krakowetz, C.N., Mann, A.S., Sim, K.A., Chilton, N.B., 2011. An assessment of genetic differences among ixodid ticks in a locus within the nuclear large subunit ribosomal RNA gene. Mol. Cell. Probes 25, 243–248. Bae, C., Szalanski, A., Robbins, R., 2008. Molecular analysis of the lance nematode, Hoplolaimus spp., using the first internal transcribed spacer and the D1–D3 expansion segments of 28S ribosomal DNA1. J. Nematol. 40, 201–209. Beati, L., Keirans, J.E., 2001. Analysis of the systematic relationships among ticks of the genera Rhipicephalus and Boophilus (Acari: Ixodidae) based on mitochondrial 12S ribosomal DNA gene sequences and morphological characters. J. Parasitol. 87, 32–48. Buckler, E.S.T., Ippolito, A., Holtsford, T.P., 1997. The evolution of ribosomal DNA: divergent paralogues and phylogenetic implications. Genetics 145, 821–832. Casati, S., Bernasconi, M.V., Gern, L., Piffaretti, J.C., 2008. Assessment of intraspecific mtDNA variability of European Ixodes ricinus sensu stricto (Acari: Ixodidae). Infect. Genet. Evol. 8, 152–158. Dabert, M., 2006. DNA markers in the phylogenetics of the Acari. Biol. Lett. 43, 97–107. De Rijk, P., Wuyts, J., De Wachter, R., 2003. RnaViz 2: an improved representation of RNA secondary structure. Bioinformatics 19, 299–300. Delaye, C., Beati, L., Aeschlimann, A., Renaud, F., de Meeus, T., 1997. Population genetic structure of Ixodes ricinus in Switzerland from allozymic data: no evidence of divergence between nearby sites. Int. J. Parasitol. 27, 769–773. Ecker, M., Allison, S.L., Meixner, T., Heinz, F.X., 1999. Sequence analysis and genetic classification of tick-borne encephalitis viruses from Europe and Asia. J. Gen. Virol. 80 (Pt 1), 179–185.
218
S.Y. Kovalev, T.A. Mukhacheva / Ticks and Tick-borne Diseases 3 (2012) 212–218
Filippova, N.A., 1985. Taiga Tick Ixodes persulcatus Schulze (Acarina, Ixodidae). Morphology, Systematics, Ecology, Medical Importance. Nauka Publishers, Leningrad, p. 416 (in Russian). Filippova, N.A., 2002. Morphological barrier in mechanisms of reproductive isolation acting in areas of sympatry of closely related species Ixodes persulcatus–I. pavlovskyi and I. persulcatus–I. ricinus (Ixodidae). Parazitologiia 36, 457–468 (in Russian). Filippova, N.A., Musatov, S.A., 1996. Geographic variability in the sexually mature phase of Ixodes persulcatus (Ixodidae). Experience in using databases on morphometry. Parazitologiia 30, 205–215 (in Russian). Foley, J., Nieto, N.C., Foley, P., Teglas, M.B., 2008. Co-phylogenetic analysis of Anaplasma phagocytophilum and its vectors, Ixodes spp. ticks. Exp. Appl. Acarol. 45, 155–170. Fomenko, N.V., Stronin, O.V., Khasnatinov, M.A., Danchinova, G.A., Bataa, J., Gol’tsova, N.A., 2009. The heterogeneity of the ospa gene of Borrelia garinii and Borrelia afzelii in western Siberia and Mongolia. Mol. Gen. Mikrobiol. Virusol., 18–22. Fukunaga, M., Yabuki, M., Hamase, A., Oliver Jr., J.H., Nakao, M., 2000. Molecular phylogenetic analysis of ixodid ticks based on the ribosomal DNA spacer, internal transcribed spacer 2, sequences. J. Parasitol. 86, 38–43. Hancock, J.M., Dover, G.A., 1990. ‘Compensatory slippage’ in the evolution of ribosomal RNA genes. Nucleic Acids Res. 18, 5949–5954. Hewitt, G.M., 2004. The structure of biodiversity – insights from molecular phylogeography. Front. Zool. 1, 4. Kain, D.E., Sperling, F.A., Daly, H.V., Lane, R.S., 1999. Mitochondrial DNA sequence variation in Ixodes pacificus (Acari: Ixodidae). Heredity 83 (Pt 4), 378–386. Kriuchechnikov, V.N., Korenberg, E.I., Shcherbakov, S.V., Kovalevskii, Iu.V., Levin, M.L., 1988. Identification of Borrelia isolated in the USSR from Ixodes persulcatus Schulze ticks. Zh. Mikrobiol. Epidemiol. Immunobiol., 41–44 (in Russian). McLain, D.K., 2001. Evolution of transcript structure and base composition of rDNA expansion segment D3 in ticks. Heredity 87, 544–557. McLain, D.K., Li, J., Oliver Jr., J.H., 2001. Interspecific and geographical variation in the sequence of rDNA expansion segment D3 of Ixodes ticks (Acari: Ixodidae). Heredity 86, 234–242. Michot, B., Hassouna, N., Bachellerie, J.P., 1984. Secondary structure of mouse 28S rRNA and general model for the folding of the large rRNA in eukaryotes. Nucleic Acids Res. 12, 4259–4279. Nava, S., Guglielmone, A.A., Mangold, A.J., 2009. An overview of systematics and evolution of ticks. Front. Biosci. 14, 2857–2877.
Navajas, M., Fenton, B., 2000. The application of molecular markers in the study of diversity in acarology: a review. Exp. Appl. Acarol. 24, 751–774. Nefedova, V.V., Korenberg, E.I., Gorelova, N.B., 2010. Genetic variants of Borrelia garinii, a widely spread Eurasian pathogen of ixodic tick borreliosis. Mol. Gen. Mikrobiol. Virusol., 7–12. Norris, D.E., Klompen, J.S., Keirans, J.E., Black IV, W.C., 1996. Population genetics of Ixodes scapularis (Acari: Ixodidae) based on mitochondrial 16S and 12S genes. J. Med. Entomol. 33, 78–89. Paabo, S., Irwin, D.M., Wilson, A.C., 1990. DNA damage promotes jumping between templates during enzymatic amplification. J. Biol. Chem. 265, 4718–4721. Qiu, W.G., Dykhuizen, D.E., Acosta, M.S., Luft, B.J., 2002. Geographic uniformity of the Lyme disease spirochete (Borrelia burgdorferi) and its shared history with tick vector (Ixodes scapularis) in the Northeastern United States. Genetics 160, 833–849. Shpynov, S.N., Rudakov, N.V., Iastrebov, V.K., Leonova, G.N., Khazova, T.G., Egorova, N.V., Borisova, O.N., Preider, V.P., Bezrukov, G.V., Fedorov, E.G., Fedianin, A.P., Sherstneva, M.B., Turyshev, A.G., Gavrilov, A.P., Tankibaev, M.A., Fournier, P.E., Raoult, D., 2004. New evidence for the detection of Ehrlichia and Anaplasma in Ixodes ticks in Russia and Kazakhstan. Med. Parazitol. (Mosk), 10–14 (in Russian). Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011. MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28, 2731–2739. Tuininga, A.R., Miller, J.L., Morath, S.U., Daniels, T.J., Falco, R.C., Marchese, M., Sahabi, S., Rosa, D., Stafford III, K.C., 2009. Isolation of entomopathogenic fungi from soils and Ixodes scapularis (Acari: Ixodidae) ticks: prevalence and methods. J. Med. Entomol. 46, 557–565. Xu, G., Fang, Q.Q., Keirans, J.E., Durden, L.A., 2003. Molecular phylogenetic analyses indicate that the Ixodes ricinus complex is a paraphyletic group. J. Parasitol. 89, 452–457. Yli-Mattila, T., Paavanen-Huhtala, S., Fenton, B., Tuovinen, T., 2000. Species and strain identification of the predatory mite Euseius finlandicus by RAPD-PCR and ITS sequences. Exp. Appl. Acarol. 24, 863–880. Zilber, L.A., 1939. Spring–summer tick-borne encephalitis. Arkhiv. Biol. Nauk. 56, 255–261. Zuker, M., Mathews, D.H., Turner, D.H., 1999. In: Clark, B.A.B.F.C. (Ed.), Algorithms and Thermodynamics for RNA Secondary Structure Prediction: A Practical Guide in RNA Biochemistry and Biotechnology. Kluwer Academic Publishers.