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RNA interference for the study and genetic manipulation of ticks
Review
TRENDS in Parasitology
Vol.23 No.9
Special issue: Tick–host–pathogen interactions in the post-genomic era
RNA interference for the study and genetic manipulation of ticks Jose´ de la Fuente1,2, Katherine M. Kocan1, Consuelo Almaza´n1,3 and Edmour F. Blouin1 1
Department of Veterinary Pathobiology, Center for Veterinary Health Sciences, Oklahoma State University, Stillwater, OK 74078, USA 2 Instituto de Investigacio´n en Recursos Cinege´ticos IREC (CSIC-UCLM-JCCM), Ronda de Toledo s/n, 13071 Ciudad Real, Spain 3 Present address: Facultad de Medicina Veterinaria y Zootecnia, Universidad Auto´noma de Tamaulipas, Km. 5 carretera Victoria-Mante, CP 87000 Victoria, Tamaulipas, Mexico
Ticks are ectoparasites of wild and domestic animals, and humans. A more comprehensive understanding of tick function and the tick–pathogen interface is needed to formulate improved tick-control methods. RNA interference (RNAi) is the most widely used gene-silencing technique in ticks where the use of other methods of genetic manipulations has been limited. In the short time that RNAi has been available, it has proved to be a valuable tool for studying tick gene function, the characterization of the tick–pathogen interface, and the screening and characterization of tick protective antigens. This review considers the applications of RNAi to tick research and the potential of this technique for tick functional studies, and to elucidate the tick–pathogen and tick–host interface. It is probable that the knowledge gained from this experimental approach will contribute to development of vaccines to control tick infestations and the transmission of tickborne pathogens. Introduction Ticks are obligate hematophagous ectoparasites of wild and domestic animals, and humans. They are considered to be second to mosquitoes as vectors of human diseases worldwide [1] and the most important vectors affecting the cattle industry worldwide [2]. Ticks are classified in the subclass Acari, order Parasitiformes, suborder Ixodida and are distributed worldwide from Arctic to tropical regions [3]. Despite efforts to control tick infestations, these ectoparasites remain a serious problem for human and animal health [4,5]. RNA interference (RNAi) [6] is a nucleic acid-based reverse genetic approach that involves disruption of gene expression to determine gene function or its effect on a metabolic pathway. Small interfering RNAs (siRNAs) are the effector molecules of the RNAi pathway, which is initiated by double-stranded RNA (dsRNA) and results in a potent sequence-specific degradation of cytoplasmic mRNAs containing the same sequence as the dsRNA trigger [7–9]. Post-transcriptional gene silencing mechanisms Corresponding author: de la Fuente, J. ([email protected]). Available online 25 July 2007. www.sciencedirect.com
initiated by dsRNA have been discovered in all eukaryotes studied thus far, and RNAi has been rapidly developed in a variety of organisms as a tool for functional genomics studies and other applications [10]. RNAi has become the most widely used gene-silencing technique in ticks and other organisms where alternative approaches for genetic manipulation are not available or are unreliable [5,11]. The genetic characterization of ticks has been limited until the recent application of RNAi [12,13]. In this paper, we review current research using RNAi in ticks and discuss future applications of RNAi to tick research. Mechanism of RNAi in ticks Little is known about the mechanism of RNAi in ticks. Most of our understanding of RNAi is derived from studies on Drosophila melanogaster, Homo sapiens, Caenorhabditis elegans, Arabidopsis thaliana and Neurospora crassa cell systems [7–10]. The most closely related organism to ticks (in terms of evolution) in which the mechanism of RNAi has been characterized is D. melanogaster. However, recent results have provided evidence of both similarities and differences between the processes of RNAi in D. melanogaster and in Anopheles gambiae, suggesting that although many of the basic mechanisms of RNAi are evolutionarily conserved, the role of particular protein components of the RNAi machinery differs [14]. The proteins involved in the process of RNAi in ticks have not been identified. However, herein we propose a model for RNAi in ticks based on current information on RNAi in D. melanogaster [8] and mosquitoes [14] (Figure 1). This model is consistent with the results of experiments characterizing viral suppressors of RNAi in tick cells, which suggest that ticks utilize a mechanism of dsRNA-mediated RNAi similar to that described in other eukaryotes [15,16]. Exogenous or viral dsRNA enter the cytoplasm, where it is first processed into double-stranded siRNAs 21–23 nucleotides in length. The key protein for this specific degradation in D. melanogaster is Dicer-1. This RNase III-like dsRNAspecific ribonuclease contains RNase III, helicase, PAZ (Piwi/Argonaute/Zwille) domains that are involved in protein–protein interactions, a dsRNA-binding domain
1471-4922/$ – see front matter ß 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.pt.2007.07.002
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Figure 1. Postulated model of molecular events in tick dsRNA-mediated RNAi. The systemic RNAi pathway might use a protein similar to Sid-1 for dsRNA cell uptake. Exogenous and viral dsRNA enter the cytoplasm, where it is first processed into double-stranded siRNAs 21–23 nucleotides in length. The key protein for this specific degradation, the D. melanogaster RNase III-like dsRNA-specific ribonuclease, Dicer-1, has not been identified in ticks. Dicer-1 then presents the siRNAs to the RNA-induced silencing complex (RISC), which incorporates the siRNAs and targets, and degrades any mRNA with cognate sequences. None of the RISC components have been identified in ticks.
and a DEAD (i.e. the single letter code for Asp–Glu–Ala– Asp)-box helicase domain [8]. Dicer-1 then presents the siRNAs to the RNA-induced silencing complex (RISC), which incorporates the siRNAs and targets, and degrades any mRNA with cognate sequences [7]. Other protein components of RISC, such as the Argonaute (Ago) family, might be required for RNAi [7]. A sequence with identity to Ago-2, found in the Boophilus microplus expressed sequence tag (EST) database (TC984; http://compbio.dfci.harvard.edu/ tgi/cgi-bin/tgi/gimain.pl?gudb=b_microplus), is the only tick sequence associated with the RNAi machinery currently identified. In plants and C. elegans, RNAi is amplified by a process that is dependent on a RNA-dependent RNA polymerase (RdRP), thus resulting in a systemic RNAi effect that can spread from the target tissue to other tissues [7]. However, although RdRP-like activity has been detected in Drosophila embryos [17], a homolog has not been identified in the genomic sequences of higher organisms, such as D. melanogaster and H. sapiens. Furthermore, RdRP-dependent RNAi is not present in D. melanogaster [18] and the mechanism described for RNAi in mosquitoes suggests that RdRP-mediated transitive amplification is absent in this species [14]. Taken together, these observations suggest that RdRP, if present, has a minor role in RNAi in higher eukaryotes. The application of dsRNA via body cavity injection, feeding or soaking leads to global and persistent gene silencing in treated ticks and their progeny [19–22]. This systemic RNAi involves the uptake of dsRNA and might be evolutionarily conserved [23]. At least one protein, systemic RNA interference-deficient-1 (Sid-1), has been identified as necessary and sufficient for systemic RNAi induction in C. elegans and the grasshopper (Schistocerca americana) [23–25]. However, the sid-1 gene has not been identified in ticks [19]. Additional evidence suggests that the basic mechanism of RNAi in ticks is similar to that reported in other www.sciencedirect.com
arthropods. Based on the known tolerance for mismatches and gaps in sequence base-pairing with target RNA during RNAi [26], experiments were designed to assay the effect of the injection of cross-species subolesin (4D8) dsRNAs on the reduction of tick weight, oviposition and survival in Ixodes scapularis and Amblyomma americanum (Figure 2). A reduction in tick weight, oviposition and survival was observed with the injection of heterologous subolesin dsRNA, probably owing to the 72% identity between I. scapularis and A. americanum subA sequences [27]. Recently, Nijhof et al. [19] and Kocan et al. [28] demonstrated in B. microplus, A. americanum, Dermacentor variabilis and I. scapularis the effect of systemic RNAi on tick progeny when dsRNA injected in the hemolymph of replete female ticks caused silencing in the next generation larvae. However, silencing of gene expression was not observed in adult ticks [19]. This result can be explained by slow dilution of the gene silencing factors and argues against amplification of RNAi in ticks, a mechanism responsible for long-term gene silencing by RNAi in C. elegans [29]. Although indirect, these results suggest a conserved RNAi mechanism in ticks. However, sequence information and functional studies are required to identify and characterize components of the RNAi machinery in ticks. Methods for RNAi in ticks Four different methods have been used to deliver dsRNA for RNAi in ticks: (i) injection; (ii) soaking; (iii) feeding; and (iv) virus production of dsRNA (Table 1). (i) Injection of dsRNA. The injection of dsRNA into ticks has been done manually with Hamilton (Reno, NV, USA) syringes (‘injection’ in Table 1) or by using microinjectors (‘microinjection’ in Table 1). Aljamali et al. [12] were the first to report the application of RNAi to ticks. Although not described in detail, they manually injected dsRNA into tick hemolymph to induce RNAi. Subsequently, de la Fuente et al.
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Table 1. Summary of RNA interference experiments in ticks Tick species Target gene a Study of tick gene function Histamine-binding Amblyomma protein (HBP) americanum
Delivery of dsRNA b
Phenotype
Refs
Soaking of salivary glands Injection in ticks
Reduction of histamine-binding activity in salivary glands and aberrant tick feeding pattern Inhibition of PGE2-stimulated anticoagulant protein secretion by isolated salivary glands Reduction in anticoagulant protein release by salivary glands and aberrant tick feeding Inhibition of tick feeding and reduced tick weight Reduction in tick survival, weight and oviposition Reduction in anticoagulant activity in salivary glands and reduction in tick weight Reduction in mRNA levels but no effect on tick feeding Inhibition of tick feeding and reduction in tick weight Reduction in tick weight and oviposition Reduction in tick survival, weight and oviposition Suppression of RNAi induced by SFV replicon
[12,13]
A. americanum
Synaptobrevin
Soaking of salivary glands
A. americanum
nSec1
Soaking of salivary glands Injection in ticks
A. americanum
Synaptobrevin cystatin
A. americanum
Subolesin (4D8)
Soaking of salivary glands Injection in ticks Injection in ticks
Ixodes scapularis
Salp14
Microinjection in ticks
I. scapularis
Salp14
I. scapularis
Actin
I. scapularis I. scapularis
Subolesin (4D8) 50 nucleotidase (4F8)
Microinjection into nymphal ticks Microinjection and injection in ticks Injection in ticks Injection in ticks
I. scapularis
Semliki Forest virus (SFV) replicon
I. scapularis Haemaphysalis longicornis H. longicornis
Subolesin (4D8) Cubilin-related serine proteinase (HISP) Follistatin-related protein (FRP) Longepsin Subolesin (4D8) Rs86 (homologue of Bm86) Subolesin (4D8)
Injection in ticks Injection in ticks
Subolesin (4D8)
Injection in ticks
Subolesin (4D8)
Injection in unfed and replete ticks
B. microplus
Bm86
B. microplus
Bm91
Injection in unfed and replete ticks Injection in unfed and replete ticks
H. longicornis Rhipicephalus sanguineus R. sanguineus Dermacentor variabilis
Dermacentor marginatus Boophilus microplus
Characterization of tick–pathogen interface TROSPA I. scapularis
Infection of ISE6 cells with recombinant SFV expressing viral suppressors Soaking of IDE8 cells Microinjection in ticks
[22]
[34] [27] [32]
[50] [30,32] [27,30] [30] [16]
c
Reduction in mRNA levels Reduction in tick weight
[38]
Microinjection in ticks
Reduction in tick oviposition
[40]
Injection in ticks Injection in ticks
Reduction in mRNA and protein levels Reduction in tick survival, weight and oviposition Reduction in tick weight and oviposition Reduction in tick survival, weight and oviposition. Diminished reproductive performance and absence of viable offspring Reduction in tick survival, weight and oviposition Reduction in tick survival, weight and oviposition. Abnormal egg development. Systemic RNAi on tick progeny Systemic RNAi on tick progeny
[51] [27,39]
[19]
Systemic RNAi on tick progeny
[19]
Reduction in infection and transmission of B. burgdorferi Reduction in the capacity of tick-borne B. burgdorferi to infect mice Reduction of survival of A. phagocytophilum in ticks Reduction in mRNA levels but no effect on acquisition of A. phagocytophilum and B. burgdorferi Reduction in tick weight and infection with B. burgdorferi and affected expression of salivary gland and nonsalivary gland proteins Reduction in tick infection by A. marginale
[45]
Microinjection nymphal ticks Microinjection nymphal ticks Microinjection nymphal ticks Microinjection nymphal ticks
into
I. scapularis
Salp15
I. scapularis
Salp16
I. scapularis
Salp14
I. scapularis
Isac
Nymph capillary feeding
D. variabilis
Subolesin (4D8)
Injection in ticks
into into into
Screening and characterization of tick protective antigens cDNA pools Injection in ticks I. scapularis Subolesin (4D8) and Rs86 Injection in ticks R. sanguineus
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[21]
Reduction in tick weight and oviposition Synergistic effect on tick weight and oviposition
[39] [27,31]
[27] [19]
[46] [47] [50]
[20]
[48] [30] [39]
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Table 1 (Continued ) Tick species B. microplus
Target gene a Subolesin (4D8)
Delivery of dsRNA b Injection in unfed and replete ticks
B. microplus
Bm86
B. microplus
Bm91
Injection in unfed and replete ticks Injection in unfed and replete ticks
Phenotype Reduction in tick survival, weight and oviposition. Abnormal egg development. Systemic RNAi on tick progeny Systemic RNAi on tick progeny
Refs [19]
[19]
Systemic RNAi on tick progeny
[19]
a
Abbreviations: Bm86, B. microplus 86; Bm91, B. microplus 91; Isac, I. scapularis anticomplement; nSec1, neuronal secretory protein 1; Rs86, R. sanguineus 86; Salp14/15/16, salivary gland protein 14/15/16. b dsRNA delivered to unfed adult ticks unless indicated otherwise. c Blouin, E.F. et al., unpublished.
[27,30,31] used a similar approach for tick RNAi by injecting dsRNA in the lower right quadrant of the ventral surface of the exoskeleton of unfed female or male ticks using a Hamilton syringe with a one inch, 33-gauge needle (Figure 3). This method has been reliable and reproducible in different tick species [27]. Narasimhan et al. [32] were the first to report the use of microinjection for dsRNA delivery into ticks. The dsRNA was injected into the ventral torso of the idiosoma, away from the anal opening, of adult I. scapularis females using microdispensers (Drummond Scientific, Broomall, PA, USA) and a micromanipulator (Narishige, Tokyo, Japan) connected to a Nanojet microinjector (Drummond Scientific). Both methods deposit dsRNA inside ticks, although microinjection might be a more controlled method owing to its possibility of better controlling the injection volume. However, the survival rate of injected ticks seems to be similar in both methods. Recently, Nijhof et al. [19] developed a method to induce RNAi in B. microplus eggs and larvae by injecting dsRNA into replete female ticks. The dsRNA was injected through the spiracle of the replete tick. This method has been recently reproduced in other tick species [28], thus facilitating gene silencing experiments in ticks and the application of RNAi to a larger number of individuals. However, although successful for tick larvae, the effect of inherited RNAi on nymphs remains to be evaluated. (ii) Soaking or incubation of dsRNA with ticks, tick tissues or cells. Soaking and incubation of dsRNA
with ticks, tick tissues or cells has been used in ex vivo, in vivo and in vitro experiments. Ex vivo soaking of tick-isolated organs in dsRNA solutions have been widely used for the study of A. americanum tick salivary gland physiology [33] (Table 1). In these experiments, salivary glands were incubated for 6 h at 37 8C with dsRNA in cell culture medium to induce RNAi of target genes [13,21,22,34]. This approach for dsRNA delivery might also be useful for RNAi on live ticks. If standardized, soaking of tick larvae into dsRNA could be used as an alternative method to induce RNAi in a large number of ticks and in immature tick stages. Taking advantage of the fact that RNAi is induced in insect cells after incubation with dsRNA [35], in vitro RNAi was obtained in tick IDE8 cells incubated with subolesin dsRNA (E.F. Blouin et al., unpublished) (Table 1). The application of RNAi to cultured tick cells will probably impact on functional genomic studies in tick species from which cell lines have been developed (see Sakyi et al. in this issue). (iii) Feeding of dsRNA to ticks. The feeding of dsRNA to ticks for RNAi was reported by Soares et al. [20], who placed capillary tubes directly over the hypostome of nymphal ticks and the ticks capillary-fed on the dsRNA solution for 2–4 h. They used oral RNAi to demonstrate the role of anticomplement gene (isac) in the nymphal infection by Borrelia burgdorferi [20]. This approach has been used on I. scapularis nymphs only but might prove applicable to other tick species. Capillary feeding is a valuable technique to address a
Figure 2. Sequence requirements for tick RNAi. I. scapularis and A. americanum female ticks were injected with homologous (endogenous; E) or heterologous (H) subolesin (4D8) dsRNA and the percent reduction of tick weight (black bars), oviposition (white bars) and survival (grey bars) was determined in dsRNA-injected ticks when compared to mock saline-injected controls. Two experiments were conducted with similar results. Bars represent mean + SD. All differences were statistically significant with respect to saline-injected controls (P < 0.05; Student’s t-Test). Subolesin dsRNAs were prepared and injected, and ticks were analyzed as previously described [27]. www.sciencedirect.com
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Figure 3. Typical in vivo RNAi experiment. Subolesin (4D8) dsRNA, or injection buffer alone as a control (C), was injected into the lower right quadrant of the ventral surface of the exoskeleton of a female A. americanum tick with a one inch, 33-gauge needle and a Hamilton syringe. After feeding, tick survival, engorgement weight and oviposition were evaluated, and compared between dsRNA-injected and control ticks. Experimental procedures and results of this experiment were published previously [27].
variety of biological questions including the study of tick–pathogen [36] and tick–host [37] interfaces. (iv) Virus production of dsRNA. The infection of tick cells with vector-borne RNA viruses, such as Semliki Forest virus (SFV), triggers the RNAi pathway and can be used for the characterization of dsRNAmediated gene silencing in ticks [16]. The RNAi methods discussed above have advantages and disadvantages, which depend on the experimental design and objectives. Injection of dsRNA into ticks might prove to be the most universal method for in vivo RNAi in ticks, particularly with the possibility of generating a high number of treated individuals through inherited RNAi [19,28]. Inherited RNAi and feeding of dsRNA solutions to ticks are relatively easy ways to deliver dsRNA into immature tick stages for the study of tick–pathogen interactions [19,20,28]. However, incubation with dsRNA solutions might be the best approach for ex vivo studies of gene expression in isolated tick organs [33]. Finally, in vitro RNAi experiments with tick cell lines might be the only approach for high throughput screening and functional genomics studies in ticks. RNAi for the study of tick gene function As a reverse genetic approach, RNAi has been used mostly for the analysis of gene function in ticks (Table 1). Functional studies have been conducted in seven tick species of the www.sciencedirect.com
genera Amblyomma, Ixodes, Haemaphysalis, Dermacentor, Rhipicephalus and Boophilus (Table 1). Typically, in vivo RNAi experiments involve the analysis of tick survival, engorgement and oviposition to analyze the effect of gene silencing on tick feeding and reproduction (Figure 3). These experiments have provided valuable information about the function of genes involved in the regulation of tick feeding [12,13,19,21,22,27,30,32,34,38,39] and reproduction [19,27,30,31,39,40]. In the near future, the availability of genome sequence information for several tick species [41,42] would enable the application of RNAi for functional genomewide studies to address fundamental questions on tick physiology, development and gene regulation. RNAi for the characterization of tick–pathogen interface The highest impact of tick infestations on animal and human populations is through the pathogens they transmit [1,2]. Differential gene expression has been characterized in D. variabilis ovaries in response to rickettsial infection [43] and in salivary glands of female Rhipicephalus appendiculatus infected with Theileria parva [44], suggesting that pathogens modify the expression of tick genes that are required for pathogen infection and are involved in tick defense mechanisms to infection. However, the function of these genes in tick–pathogen interactions is largely unknown.
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Recently, tick proteins have been characterized that have a role in the infection and transmission of B. burgdorferi [20,45,46] and Anaplasma phagocytophilum [47] (Table 1). RNAi silencing of genes that encode these proteins results in reduced pathogen infection and/or transmission. One of these proteins, the tick receptor for outer surface protein A (TROSPA) that is required for B. burgdorferi colonization of I. scapularis, exemplifies the association between the tick vector and the pathogen [45]. Pal et al. [45] identified outer surface protein A (OspA) as the bacterial ligand for TROSPA and demonstrated that the blockade of TROSPA by anti-TROSPA antibodies or by the repression of TROSPA expression via RNAi reduced bacterial adherence to the I. scapularis gut in vivo, thereby preventing efficient colonization of the vector, and subsequently reducing pathogen transmission to the mammalian host. Experiments with the tick protective antigen subolesin provided evidence that tick proteins might be targeted to test their role in the vectorial capacity of ticks [48]. Subolesin was recently shown by both RNAi gene silencing and immunization with the recombinant protein to protect against tick infestations, and to cause reduced tick survival and degeneration of gut, salivary gland and reproductive tissues [27,31,37,49]. In addition, targeting subolesin expression by RNAi decreased the ability of ticks to become infected with Anaplasma marginale, the agent of bovine anaplasmosis [48]. Collectively, these results demonstrate that RNAi constitutes an important tool for the study of the tick– pathogen interface, and might contribute to the rapid identification and characterization of potential pathogen transmission-blocking tick vaccine antigens. RNAi for the screening and characterization of tick protective antigens Control of ticks has been attempted primarily by application of acaricides, a method accompanied by serious drawbacks, such as environmental contamination and selection of acaricide-resistant ticks [4,5]. Alternative new approaches have shown promise for tick control, but their use and efficacy have been limited [4]. Vaccines have been shown to be a feasible tick-control method that offers a cost-effective, environmentally friendly alternative to chemical control [4,5]. However, identification of tick protective antigens remains the limiting step in vaccine development. Discovery of protective antigens, along with proper vaccine formulations, field-trial evaluation and commercialization, are all major components required for development of tick vaccines. Recently, de la Fuente et al. [30] demonstrated that RNAi could be used for the screening of tick protective antigens. Through RNAi, it is possible to screen a large number of genes for potential vaccine candidates in a relatively short time and with minimal use of laboratory animals. Selected antigens could then be characterized and evaluated as recombinant proteins in controlled vaccine trials. RNAi might be also used for the preliminary characterization of tick protective antigens and antigen combinations before conducting more expensive vaccine trials for the development of improved vaccine compositions [19,39]. www.sciencedirect.com
Prospects for RNAi applications in ticks The application of RNAi to tick research is still in its infancy. However, RNAi has been shown to be a valuable tool for the study of tick gene function, the characterization of the tick–pathogen interface, and the screening and characterization of tick protective antigens. Tick genomics and proteomics are likely to evolve into projects addressing the sequencing, annotation and functional analysis of tick genomes, providing invaluable information to address basic biological questions and for the development of tick vaccines. For functional analyses, RNAi might become the most valuable tool, used in conjunction with other technologies such as immunomapping and expression library immunization. Methodologically, RNAi will probably evolve into more efficient methods that might enable modification of gene expression in a large number of individuals that would make feasible the application of autocidal control for reduction of tick populations [31]. The mechanism of dsRNA-induced RNAi in ticks should be refined to contribute to a better understanding and utilization of this genetic approach in this species. Finally, RNAi will probably provide comprehensive contributions to the study of the tick– pathogen interface, and might have an impact on the development of vaccines to control tick infestations and the transmission of tick-borne pathogens. Acknowledgements We thank members of our laboratories for fruitful discussions and technical assistance. Part of this work was supported by project no. 1669 of the Oklahoma Agricultural Experiment Station and the Endowed Chair for Food Animal Research (K.M. Kocan), the Instituto de Ciencias de la Salud, Spain (ICS-JCCM) (project no. 06036–00), and the Wellcome Trust under the Animal Health in the Developing World initiative through project 0757990 entitled ‘Adapting recombinant anti-tick vaccines to livestock in Africa’. The writing of this paper has been facilitated through The Integrated Consortium on Ticks and Tick-borne Diseases (ICTTD-3), financed by the International Cooperation Programme of the European Union through Coordination Action Project no. 510561.
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33 Bowman, A.S. and Sauer, J.R. (2004) Tick salivary glands: function, physiology and future. Parasitology 129 (Suppl.), S67–S81 34 Karim, S. et al. (2005) RNAi-mediated gene silencing to assess the role of synaptobrevin and cystatin in tick blood feeding. Biochem. Biophys. Res. Commun. 334, 1336–1342 35 Eggert, U.S. et al. (2004) Parallel chemical genetic and genome-wide RNAi screens identify cytokinesis inhibitors and targets. PLoS Biol. 2, e379 36 Kocan, K.M. et al. (2005) Capillary tube feeding system for studying tick-pathogen interactions of Dermacentor variabilis (Acari: Ixodidae) and Anaplasma marginale (Rickettsiales: Anaplasmataceae). J. Med. Entomol. 42, 864–874 37 Almaza´n, C. et al. (2005) Characterization of three Ixodes scapularis cDNAs protective against tick infestations. Vaccine 23, 4403–4416 38 Miyoshi, T. et al. (2004) Gene silencing of a cubilin-related serine proteinase from the hard tick Haemaphysalis longicornis by RNA interference. J. Vet. Med. Sci. 66, 1471–1473 39 de la Fuente, J. et al. (2006) Synergistic effect of silencing the expression of tick protective antigens 4D8 and Rs86 in Rhipicephalus sanguineus by RNA interference. Parasitol. Res. 99, 108–113 40 Zhou, J. et al. (2006) Identification of a follistatin-related protein from the tick Haemaphysalis longicornis and its effect on tick oviposition. Gene 372, 191–198 41 Hill, C.A. and Wikel, S.K. (2005) The Ixodes scapularis genome project: an opportunity for advancing tick research. Trends Parasitol. 21, 151– 153 42 Guerrero, F.D. et al. (2006) Sequencing a new target genome: the Boophilus microplus (Acari: Ixodidae) genome project. J. Med. Entomol. 43, 9–16 43 Mulenga, A. et al. (2003) Dynamics of Rickettsia-tick interactions: identification and characterization of differentially expressed mRNAs in uninfected and infected Dermacentor variabilis. Insect Mol. Biol. 12, 185–193 44 Nene, V. et al. (2004) Genes transcribed in the salivary glands of female Rhipicephalus appendiculatus ticks infected with Theileria parva. Insect Biochem. Mol. Biol. 34, 1117–1128 45 Pal, U. et al. (2004) TROSPA, an Ixodes scapularis receptor for Borrelia burgdorferi. Cell 119, 457–468 46 Ramamoorthi, N. et al. (2005) The Lyme disease agent exploits a tick protein to infect the mammalian host. Nature 436, 573–577 47 Sukumaran, B. et al. (2006) An Ixodes scapularis protein required for survival of Anaplasma phagocytophilum in tick salivary glands. J. Exp. Med. 203, 1507–1517 48 de la Fuente, J. et al. (2006) Reduction of tick infections with Anaplasma marginale and A. phagocytophilum by targeting the tick protective antigen subolesin. Parasitol. Res. 100, 85–91 49 Almaza´n, C. et al. (2005) Vaccination with recombinant tick antigens for the control of Ixodes scapularis adult infestations. Vaccine 23, 5294– 5298 50 Pedra, J.H. et al. (2006) Disruption of the salivary protein 14 in Ixodes scapularis nymphs and impact on pathogen acquisition. Am. J. Trop. Med. Hyg. 75, 677–682 51 Boldbaatar, D. et al. (2006) Molecular cloning and functional characterization of an aspartic protease from the hard tick Haemaphysalis longicornis. Insect Biochem. Mol. Biol. 36, 25–36
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Special issue: Tick–host–pathogen interactions in the post-genomic era
RNA interference for the study and genetic manipulation of ticks Jose´ de la Fuente1,2, Katherine M. Kocan1, Consuelo Almaza´n1,3 and Edmour F. Blouin1 1
Department of Veterinary Pathobiology, Center for Veterinary Health Sciences, Oklahoma State University, Stillwater, OK 74078, USA 2 Instituto de Investigacio´n en Recursos Cinege´ticos IREC (CSIC-UCLM-JCCM), Ronda de Toledo s/n, 13071 Ciudad Real, Spain 3 Present address: Facultad de Medicina Veterinaria y Zootecnia, Universidad Auto´noma de Tamaulipas, Km. 5 carretera Victoria-Mante, CP 87000 Victoria, Tamaulipas, Mexico
Ticks are ectoparasites of wild and domestic animals, and humans. A more comprehensive understanding of tick function and the tick–pathogen interface is needed to formulate improved tick-control methods. RNA interference (RNAi) is the most widely used gene-silencing technique in ticks where the use of other methods of genetic manipulations has been limited. In the short time that RNAi has been available, it has proved to be a valuable tool for studying tick gene function, the characterization of the tick–pathogen interface, and the screening and characterization of tick protective antigens. This review considers the applications of RNAi to tick research and the potential of this technique for tick functional studies, and to elucidate the tick–pathogen and tick–host interface. It is probable that the knowledge gained from this experimental approach will contribute to development of vaccines to control tick infestations and the transmission of tickborne pathogens. Introduction Ticks are obligate hematophagous ectoparasites of wild and domestic animals, and humans. They are considered to be second to mosquitoes as vectors of human diseases worldwide [1] and the most important vectors affecting the cattle industry worldwide [2]. Ticks are classified in the subclass Acari, order Parasitiformes, suborder Ixodida and are distributed worldwide from Arctic to tropical regions [3]. Despite efforts to control tick infestations, these ectoparasites remain a serious problem for human and animal health [4,5]. RNA interference (RNAi) [6] is a nucleic acid-based reverse genetic approach that involves disruption of gene expression to determine gene function or its effect on a metabolic pathway. Small interfering RNAs (siRNAs) are the effector molecules of the RNAi pathway, which is initiated by double-stranded RNA (dsRNA) and results in a potent sequence-specific degradation of cytoplasmic mRNAs containing the same sequence as the dsRNA trigger [7–9]. Post-transcriptional gene silencing mechanisms Corresponding author: de la Fuente, J. ([email protected]). Available online 25 July 2007. www.sciencedirect.com
initiated by dsRNA have been discovered in all eukaryotes studied thus far, and RNAi has been rapidly developed in a variety of organisms as a tool for functional genomics studies and other applications [10]. RNAi has become the most widely used gene-silencing technique in ticks and other organisms where alternative approaches for genetic manipulation are not available or are unreliable [5,11]. The genetic characterization of ticks has been limited until the recent application of RNAi [12,13]. In this paper, we review current research using RNAi in ticks and discuss future applications of RNAi to tick research. Mechanism of RNAi in ticks Little is known about the mechanism of RNAi in ticks. Most of our understanding of RNAi is derived from studies on Drosophila melanogaster, Homo sapiens, Caenorhabditis elegans, Arabidopsis thaliana and Neurospora crassa cell systems [7–10]. The most closely related organism to ticks (in terms of evolution) in which the mechanism of RNAi has been characterized is D. melanogaster. However, recent results have provided evidence of both similarities and differences between the processes of RNAi in D. melanogaster and in Anopheles gambiae, suggesting that although many of the basic mechanisms of RNAi are evolutionarily conserved, the role of particular protein components of the RNAi machinery differs [14]. The proteins involved in the process of RNAi in ticks have not been identified. However, herein we propose a model for RNAi in ticks based on current information on RNAi in D. melanogaster [8] and mosquitoes [14] (Figure 1). This model is consistent with the results of experiments characterizing viral suppressors of RNAi in tick cells, which suggest that ticks utilize a mechanism of dsRNA-mediated RNAi similar to that described in other eukaryotes [15,16]. Exogenous or viral dsRNA enter the cytoplasm, where it is first processed into double-stranded siRNAs 21–23 nucleotides in length. The key protein for this specific degradation in D. melanogaster is Dicer-1. This RNase III-like dsRNAspecific ribonuclease contains RNase III, helicase, PAZ (Piwi/Argonaute/Zwille) domains that are involved in protein–protein interactions, a dsRNA-binding domain
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Figure 1. Postulated model of molecular events in tick dsRNA-mediated RNAi. The systemic RNAi pathway might use a protein similar to Sid-1 for dsRNA cell uptake. Exogenous and viral dsRNA enter the cytoplasm, where it is first processed into double-stranded siRNAs 21–23 nucleotides in length. The key protein for this specific degradation, the D. melanogaster RNase III-like dsRNA-specific ribonuclease, Dicer-1, has not been identified in ticks. Dicer-1 then presents the siRNAs to the RNA-induced silencing complex (RISC), which incorporates the siRNAs and targets, and degrades any mRNA with cognate sequences. None of the RISC components have been identified in ticks.
and a DEAD (i.e. the single letter code for Asp–Glu–Ala– Asp)-box helicase domain [8]. Dicer-1 then presents the siRNAs to the RNA-induced silencing complex (RISC), which incorporates the siRNAs and targets, and degrades any mRNA with cognate sequences [7]. Other protein components of RISC, such as the Argonaute (Ago) family, might be required for RNAi [7]. A sequence with identity to Ago-2, found in the Boophilus microplus expressed sequence tag (EST) database (TC984; http://compbio.dfci.harvard.edu/ tgi/cgi-bin/tgi/gimain.pl?gudb=b_microplus), is the only tick sequence associated with the RNAi machinery currently identified. In plants and C. elegans, RNAi is amplified by a process that is dependent on a RNA-dependent RNA polymerase (RdRP), thus resulting in a systemic RNAi effect that can spread from the target tissue to other tissues [7]. However, although RdRP-like activity has been detected in Drosophila embryos [17], a homolog has not been identified in the genomic sequences of higher organisms, such as D. melanogaster and H. sapiens. Furthermore, RdRP-dependent RNAi is not present in D. melanogaster [18] and the mechanism described for RNAi in mosquitoes suggests that RdRP-mediated transitive amplification is absent in this species [14]. Taken together, these observations suggest that RdRP, if present, has a minor role in RNAi in higher eukaryotes. The application of dsRNA via body cavity injection, feeding or soaking leads to global and persistent gene silencing in treated ticks and their progeny [19–22]. This systemic RNAi involves the uptake of dsRNA and might be evolutionarily conserved [23]. At least one protein, systemic RNA interference-deficient-1 (Sid-1), has been identified as necessary and sufficient for systemic RNAi induction in C. elegans and the grasshopper (Schistocerca americana) [23–25]. However, the sid-1 gene has not been identified in ticks [19]. Additional evidence suggests that the basic mechanism of RNAi in ticks is similar to that reported in other www.sciencedirect.com
arthropods. Based on the known tolerance for mismatches and gaps in sequence base-pairing with target RNA during RNAi [26], experiments were designed to assay the effect of the injection of cross-species subolesin (4D8) dsRNAs on the reduction of tick weight, oviposition and survival in Ixodes scapularis and Amblyomma americanum (Figure 2). A reduction in tick weight, oviposition and survival was observed with the injection of heterologous subolesin dsRNA, probably owing to the 72% identity between I. scapularis and A. americanum subA sequences [27]. Recently, Nijhof et al. [19] and Kocan et al. [28] demonstrated in B. microplus, A. americanum, Dermacentor variabilis and I. scapularis the effect of systemic RNAi on tick progeny when dsRNA injected in the hemolymph of replete female ticks caused silencing in the next generation larvae. However, silencing of gene expression was not observed in adult ticks [19]. This result can be explained by slow dilution of the gene silencing factors and argues against amplification of RNAi in ticks, a mechanism responsible for long-term gene silencing by RNAi in C. elegans [29]. Although indirect, these results suggest a conserved RNAi mechanism in ticks. However, sequence information and functional studies are required to identify and characterize components of the RNAi machinery in ticks. Methods for RNAi in ticks Four different methods have been used to deliver dsRNA for RNAi in ticks: (i) injection; (ii) soaking; (iii) feeding; and (iv) virus production of dsRNA (Table 1). (i) Injection of dsRNA. The injection of dsRNA into ticks has been done manually with Hamilton (Reno, NV, USA) syringes (‘injection’ in Table 1) or by using microinjectors (‘microinjection’ in Table 1). Aljamali et al. [12] were the first to report the application of RNAi to ticks. Although not described in detail, they manually injected dsRNA into tick hemolymph to induce RNAi. Subsequently, de la Fuente et al.
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Table 1. Summary of RNA interference experiments in ticks Tick species Target gene a Study of tick gene function Histamine-binding Amblyomma protein (HBP) americanum
Delivery of dsRNA b
Phenotype
Refs
Soaking of salivary glands Injection in ticks
Reduction of histamine-binding activity in salivary glands and aberrant tick feeding pattern Inhibition of PGE2-stimulated anticoagulant protein secretion by isolated salivary glands Reduction in anticoagulant protein release by salivary glands and aberrant tick feeding Inhibition of tick feeding and reduced tick weight Reduction in tick survival, weight and oviposition Reduction in anticoagulant activity in salivary glands and reduction in tick weight Reduction in mRNA levels but no effect on tick feeding Inhibition of tick feeding and reduction in tick weight Reduction in tick weight and oviposition Reduction in tick survival, weight and oviposition Suppression of RNAi induced by SFV replicon
[12,13]
A. americanum
Synaptobrevin
Soaking of salivary glands
A. americanum
nSec1
Soaking of salivary glands Injection in ticks
A. americanum
Synaptobrevin cystatin
A. americanum
Subolesin (4D8)
Soaking of salivary glands Injection in ticks Injection in ticks
Ixodes scapularis
Salp14
Microinjection in ticks
I. scapularis
Salp14
I. scapularis
Actin
I. scapularis I. scapularis
Subolesin (4D8) 50 nucleotidase (4F8)
Microinjection into nymphal ticks Microinjection and injection in ticks Injection in ticks Injection in ticks
I. scapularis
Semliki Forest virus (SFV) replicon
I. scapularis Haemaphysalis longicornis H. longicornis
Subolesin (4D8) Cubilin-related serine proteinase (HISP) Follistatin-related protein (FRP) Longepsin Subolesin (4D8) Rs86 (homologue of Bm86) Subolesin (4D8)
Injection in ticks Injection in ticks
Subolesin (4D8)
Injection in ticks
Subolesin (4D8)
Injection in unfed and replete ticks
B. microplus
Bm86
B. microplus
Bm91
Injection in unfed and replete ticks Injection in unfed and replete ticks
H. longicornis Rhipicephalus sanguineus R. sanguineus Dermacentor variabilis
Dermacentor marginatus Boophilus microplus
Characterization of tick–pathogen interface TROSPA I. scapularis
Infection of ISE6 cells with recombinant SFV expressing viral suppressors Soaking of IDE8 cells Microinjection in ticks
[22]
[34] [27] [32]
[50] [30,32] [27,30] [30] [16]
c
Reduction in mRNA levels Reduction in tick weight
[38]
Microinjection in ticks
Reduction in tick oviposition
[40]
Injection in ticks Injection in ticks
Reduction in mRNA and protein levels Reduction in tick survival, weight and oviposition Reduction in tick weight and oviposition Reduction in tick survival, weight and oviposition. Diminished reproductive performance and absence of viable offspring Reduction in tick survival, weight and oviposition Reduction in tick survival, weight and oviposition. Abnormal egg development. Systemic RNAi on tick progeny Systemic RNAi on tick progeny
[51] [27,39]
[19]
Systemic RNAi on tick progeny
[19]
Reduction in infection and transmission of B. burgdorferi Reduction in the capacity of tick-borne B. burgdorferi to infect mice Reduction of survival of A. phagocytophilum in ticks Reduction in mRNA levels but no effect on acquisition of A. phagocytophilum and B. burgdorferi Reduction in tick weight and infection with B. burgdorferi and affected expression of salivary gland and nonsalivary gland proteins Reduction in tick infection by A. marginale
[45]
Microinjection nymphal ticks Microinjection nymphal ticks Microinjection nymphal ticks Microinjection nymphal ticks
into
I. scapularis
Salp15
I. scapularis
Salp16
I. scapularis
Salp14
I. scapularis
Isac
Nymph capillary feeding
D. variabilis
Subolesin (4D8)
Injection in ticks
into into into
Screening and characterization of tick protective antigens cDNA pools Injection in ticks I. scapularis Subolesin (4D8) and Rs86 Injection in ticks R. sanguineus
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[21]
Reduction in tick weight and oviposition Synergistic effect on tick weight and oviposition
[39] [27,31]
[27] [19]
[46] [47] [50]
[20]
[48] [30] [39]
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Table 1 (Continued ) Tick species B. microplus
Target gene a Subolesin (4D8)
Delivery of dsRNA b Injection in unfed and replete ticks
B. microplus
Bm86
B. microplus
Bm91
Injection in unfed and replete ticks Injection in unfed and replete ticks
Phenotype Reduction in tick survival, weight and oviposition. Abnormal egg development. Systemic RNAi on tick progeny Systemic RNAi on tick progeny
Refs [19]
[19]
Systemic RNAi on tick progeny
[19]
a
Abbreviations: Bm86, B. microplus 86; Bm91, B. microplus 91; Isac, I. scapularis anticomplement; nSec1, neuronal secretory protein 1; Rs86, R. sanguineus 86; Salp14/15/16, salivary gland protein 14/15/16. b dsRNA delivered to unfed adult ticks unless indicated otherwise. c Blouin, E.F. et al., unpublished.
[27,30,31] used a similar approach for tick RNAi by injecting dsRNA in the lower right quadrant of the ventral surface of the exoskeleton of unfed female or male ticks using a Hamilton syringe with a one inch, 33-gauge needle (Figure 3). This method has been reliable and reproducible in different tick species [27]. Narasimhan et al. [32] were the first to report the use of microinjection for dsRNA delivery into ticks. The dsRNA was injected into the ventral torso of the idiosoma, away from the anal opening, of adult I. scapularis females using microdispensers (Drummond Scientific, Broomall, PA, USA) and a micromanipulator (Narishige, Tokyo, Japan) connected to a Nanojet microinjector (Drummond Scientific). Both methods deposit dsRNA inside ticks, although microinjection might be a more controlled method owing to its possibility of better controlling the injection volume. However, the survival rate of injected ticks seems to be similar in both methods. Recently, Nijhof et al. [19] developed a method to induce RNAi in B. microplus eggs and larvae by injecting dsRNA into replete female ticks. The dsRNA was injected through the spiracle of the replete tick. This method has been recently reproduced in other tick species [28], thus facilitating gene silencing experiments in ticks and the application of RNAi to a larger number of individuals. However, although successful for tick larvae, the effect of inherited RNAi on nymphs remains to be evaluated. (ii) Soaking or incubation of dsRNA with ticks, tick tissues or cells. Soaking and incubation of dsRNA
with ticks, tick tissues or cells has been used in ex vivo, in vivo and in vitro experiments. Ex vivo soaking of tick-isolated organs in dsRNA solutions have been widely used for the study of A. americanum tick salivary gland physiology [33] (Table 1). In these experiments, salivary glands were incubated for 6 h at 37 8C with dsRNA in cell culture medium to induce RNAi of target genes [13,21,22,34]. This approach for dsRNA delivery might also be useful for RNAi on live ticks. If standardized, soaking of tick larvae into dsRNA could be used as an alternative method to induce RNAi in a large number of ticks and in immature tick stages. Taking advantage of the fact that RNAi is induced in insect cells after incubation with dsRNA [35], in vitro RNAi was obtained in tick IDE8 cells incubated with subolesin dsRNA (E.F. Blouin et al., unpublished) (Table 1). The application of RNAi to cultured tick cells will probably impact on functional genomic studies in tick species from which cell lines have been developed (see Sakyi et al. in this issue). (iii) Feeding of dsRNA to ticks. The feeding of dsRNA to ticks for RNAi was reported by Soares et al. [20], who placed capillary tubes directly over the hypostome of nymphal ticks and the ticks capillary-fed on the dsRNA solution for 2–4 h. They used oral RNAi to demonstrate the role of anticomplement gene (isac) in the nymphal infection by Borrelia burgdorferi [20]. This approach has been used on I. scapularis nymphs only but might prove applicable to other tick species. Capillary feeding is a valuable technique to address a
Figure 2. Sequence requirements for tick RNAi. I. scapularis and A. americanum female ticks were injected with homologous (endogenous; E) or heterologous (H) subolesin (4D8) dsRNA and the percent reduction of tick weight (black bars), oviposition (white bars) and survival (grey bars) was determined in dsRNA-injected ticks when compared to mock saline-injected controls. Two experiments were conducted with similar results. Bars represent mean + SD. All differences were statistically significant with respect to saline-injected controls (P < 0.05; Student’s t-Test). Subolesin dsRNAs were prepared and injected, and ticks were analyzed as previously described [27]. www.sciencedirect.com
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Figure 3. Typical in vivo RNAi experiment. Subolesin (4D8) dsRNA, or injection buffer alone as a control (C), was injected into the lower right quadrant of the ventral surface of the exoskeleton of a female A. americanum tick with a one inch, 33-gauge needle and a Hamilton syringe. After feeding, tick survival, engorgement weight and oviposition were evaluated, and compared between dsRNA-injected and control ticks. Experimental procedures and results of this experiment were published previously [27].
variety of biological questions including the study of tick–pathogen [36] and tick–host [37] interfaces. (iv) Virus production of dsRNA. The infection of tick cells with vector-borne RNA viruses, such as Semliki Forest virus (SFV), triggers the RNAi pathway and can be used for the characterization of dsRNAmediated gene silencing in ticks [16]. The RNAi methods discussed above have advantages and disadvantages, which depend on the experimental design and objectives. Injection of dsRNA into ticks might prove to be the most universal method for in vivo RNAi in ticks, particularly with the possibility of generating a high number of treated individuals through inherited RNAi [19,28]. Inherited RNAi and feeding of dsRNA solutions to ticks are relatively easy ways to deliver dsRNA into immature tick stages for the study of tick–pathogen interactions [19,20,28]. However, incubation with dsRNA solutions might be the best approach for ex vivo studies of gene expression in isolated tick organs [33]. Finally, in vitro RNAi experiments with tick cell lines might be the only approach for high throughput screening and functional genomics studies in ticks. RNAi for the study of tick gene function As a reverse genetic approach, RNAi has been used mostly for the analysis of gene function in ticks (Table 1). Functional studies have been conducted in seven tick species of the www.sciencedirect.com
genera Amblyomma, Ixodes, Haemaphysalis, Dermacentor, Rhipicephalus and Boophilus (Table 1). Typically, in vivo RNAi experiments involve the analysis of tick survival, engorgement and oviposition to analyze the effect of gene silencing on tick feeding and reproduction (Figure 3). These experiments have provided valuable information about the function of genes involved in the regulation of tick feeding [12,13,19,21,22,27,30,32,34,38,39] and reproduction [19,27,30,31,39,40]. In the near future, the availability of genome sequence information for several tick species [41,42] would enable the application of RNAi for functional genomewide studies to address fundamental questions on tick physiology, development and gene regulation. RNAi for the characterization of tick–pathogen interface The highest impact of tick infestations on animal and human populations is through the pathogens they transmit [1,2]. Differential gene expression has been characterized in D. variabilis ovaries in response to rickettsial infection [43] and in salivary glands of female Rhipicephalus appendiculatus infected with Theileria parva [44], suggesting that pathogens modify the expression of tick genes that are required for pathogen infection and are involved in tick defense mechanisms to infection. However, the function of these genes in tick–pathogen interactions is largely unknown.
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Recently, tick proteins have been characterized that have a role in the infection and transmission of B. burgdorferi [20,45,46] and Anaplasma phagocytophilum [47] (Table 1). RNAi silencing of genes that encode these proteins results in reduced pathogen infection and/or transmission. One of these proteins, the tick receptor for outer surface protein A (TROSPA) that is required for B. burgdorferi colonization of I. scapularis, exemplifies the association between the tick vector and the pathogen [45]. Pal et al. [45] identified outer surface protein A (OspA) as the bacterial ligand for TROSPA and demonstrated that the blockade of TROSPA by anti-TROSPA antibodies or by the repression of TROSPA expression via RNAi reduced bacterial adherence to the I. scapularis gut in vivo, thereby preventing efficient colonization of the vector, and subsequently reducing pathogen transmission to the mammalian host. Experiments with the tick protective antigen subolesin provided evidence that tick proteins might be targeted to test their role in the vectorial capacity of ticks [48]. Subolesin was recently shown by both RNAi gene silencing and immunization with the recombinant protein to protect against tick infestations, and to cause reduced tick survival and degeneration of gut, salivary gland and reproductive tissues [27,31,37,49]. In addition, targeting subolesin expression by RNAi decreased the ability of ticks to become infected with Anaplasma marginale, the agent of bovine anaplasmosis [48]. Collectively, these results demonstrate that RNAi constitutes an important tool for the study of the tick– pathogen interface, and might contribute to the rapid identification and characterization of potential pathogen transmission-blocking tick vaccine antigens. RNAi for the screening and characterization of tick protective antigens Control of ticks has been attempted primarily by application of acaricides, a method accompanied by serious drawbacks, such as environmental contamination and selection of acaricide-resistant ticks [4,5]. Alternative new approaches have shown promise for tick control, but their use and efficacy have been limited [4]. Vaccines have been shown to be a feasible tick-control method that offers a cost-effective, environmentally friendly alternative to chemical control [4,5]. However, identification of tick protective antigens remains the limiting step in vaccine development. Discovery of protective antigens, along with proper vaccine formulations, field-trial evaluation and commercialization, are all major components required for development of tick vaccines. Recently, de la Fuente et al. [30] demonstrated that RNAi could be used for the screening of tick protective antigens. Through RNAi, it is possible to screen a large number of genes for potential vaccine candidates in a relatively short time and with minimal use of laboratory animals. Selected antigens could then be characterized and evaluated as recombinant proteins in controlled vaccine trials. RNAi might be also used for the preliminary characterization of tick protective antigens and antigen combinations before conducting more expensive vaccine trials for the development of improved vaccine compositions [19,39]. www.sciencedirect.com
Prospects for RNAi applications in ticks The application of RNAi to tick research is still in its infancy. However, RNAi has been shown to be a valuable tool for the study of tick gene function, the characterization of the tick–pathogen interface, and the screening and characterization of tick protective antigens. Tick genomics and proteomics are likely to evolve into projects addressing the sequencing, annotation and functional analysis of tick genomes, providing invaluable information to address basic biological questions and for the development of tick vaccines. For functional analyses, RNAi might become the most valuable tool, used in conjunction with other technologies such as immunomapping and expression library immunization. Methodologically, RNAi will probably evolve into more efficient methods that might enable modification of gene expression in a large number of individuals that would make feasible the application of autocidal control for reduction of tick populations [31]. The mechanism of dsRNA-induced RNAi in ticks should be refined to contribute to a better understanding and utilization of this genetic approach in this species. Finally, RNAi will probably provide comprehensive contributions to the study of the tick– pathogen interface, and might have an impact on the development of vaccines to control tick infestations and the transmission of tick-borne pathogens. Acknowledgements We thank members of our laboratories for fruitful discussions and technical assistance. Part of this work was supported by project no. 1669 of the Oklahoma Agricultural Experiment Station and the Endowed Chair for Food Animal Research (K.M. Kocan), the Instituto de Ciencias de la Salud, Spain (ICS-JCCM) (project no. 06036–00), and the Wellcome Trust under the Animal Health in the Developing World initiative through project 0757990 entitled ‘Adapting recombinant anti-tick vaccines to livestock in Africa’. The writing of this paper has been facilitated through The Integrated Consortium on Ticks and Tick-borne Diseases (ICTTD-3), financed by the International Cooperation Programme of the European Union through Coordination Action Project no. 510561.
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