Arsenophonus nasoniae, a male-killing endosymbiont of chalcid wasps, was recently detected in several hard tick species. Following the hypothesis that its presence in ticks may not be linked to the direct occurrence of bacteria in tick's organs, we identified A. nasoniae in wasps emerging from parasitised nymphs. We confirmed that 28.1% of Ixodiphagus hookeri wasps parasitizing Ixodes ricinus ticks were infected by A. nasoniae. Moreover, in examined I. ricinus nymphs, A. nasoniae was detected only in those, which were parasitized by the wasp. However, in part of the adult wasps as well as in some ticks that contained wasp's DNA, we did not confirm A. nasoniae. We also found, that in spite of reported male-killing, some newly emerged adult wasp males were also infected by A. nasoniae. Additionally, we amplified the DNA of Rickettsia helvetica and Rickettsia monacensis (known to be Ixodes ricinus-associated bacteria) in adult parasitoid wasps. This may be related either with the digested bacterial DNA in wasp body lumen or with a role of wasps in circulation of rickettsiae among tick vectors.
Citation: Bohacsova M, Mediannikov O, Kazimirova M, Raoult D, Sekeyova Z (2016) Arsenophonus nasoniae and Rickettsiae Infection of Ixodes ricinus Due to Parasitic Wasp Ixodiphagus hookeri. PLoS ONE 11(2): e0149950. https://doi.org/10.1371/journal.pone.0149950
Editor: Utpal Pal, University of Maryland, College Park, UNITED STATES
Received: November 10, 2015; Accepted: February 8, 2016; Published: February 22, 2016
Copyright: © 2016 Bohacsova et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the paper.
Funding: This work was supported by APVV-0280-12 from Slovak Research and Development Agency, VEGA Nos. 2/0005/15 from the Scientific Grant Agency of Ministry of Education and Slovak Academy of Sciences, by EU grant FP7-261504 EDENext and "Fondation - IHU Méditerranée Infection."
Competing interests: The authors have declared that no competing interests exist.
Arsenophonus nasoniae is a rod-like Gammaproteobacterium with a large genome and a substantial metabolic capability [1;2]. In laboratory conditions it is cultivable in supplemented cell-free media . Arsenophonus nasoniae harbours unusual combination of pathogenic and symbiotic features. It was originally described as an endosymbiont of the wasp Nasonia vitripennis, yet induces the so-called male killing phenotype—production of female biased secondary sex ratio associated with the death of 80% of male offspring [4;5]. Unlike other intracellular bacteria, A. nasoniae was found only within the somatic tissue and interstitial fluid surrounding the germ cells. These bacteria are ingested by feeding wasp larvae and invade its body through the gut, thus capable to re-infect the reproductive and other tissues of the parasitoid . Arsenophonus nasoniae was described in several parasitoid wasp species which belong to the Chalcidoidea superfamily, e.g.: Nasonia longicornis, Spalangia endius, Spalangia cameronii, Eupelmus vesiculari, Pachycrepoideus vindemmiae and Muscidifurax raptor [7;8].
The chalcid wasp, I. hookeri, was reported in Central Europe for the first time by Bouček and Černý . However, it is distributed worldwide and parasitizes a broad range of hard tick species. This parasitic wasp oviposits to feeding and unfed larval and nymphal tick hosts, but the development of the eggs only occurs in fully engorged nymphs . Once hatched, larvae of the wasp begin to feed on the tick tissue, eventually causing its death. Unlike other parasitic Chalcidoidea, larvae of I. hookeri consume also the vertebrate blood ingested by the nymphs. This diet is unique among the parasitic species of Hymenoptera families . Wasps emerge from their hosts through a single hole gnawed at the posterior end of the abdomen, where the cuticle is the thinnest .
Rickettsiae are well-known tick-associated, obligate intracellular Alfaproteobacteria . Ecological characteristics of the tick vectors of rickettsiae influence the epidemiology and clinical aspects of tick-borne diseases . Ticks may acquire rickettsiae through transovarial transmission (the transfer of bacteria from the adult female ticks to the subsequent generation of ticks via the eggs) and/or transstadial transmission, i.e. the transfer of bacteria from stage to stage .
In this study we developed a specific Real-Time PCR tailored for the detection of the I. hookeri DNA and aimed to find (1) an association between the presence of A. nasoniae and I. hookeri in ticks and (2) an influence of the parasitoids on rickettsial infection in ticks. Finally, we hypothesized how the bacteria and parasitoids could act in the developmental cycle of ticks to which the ontogenesis of wasps is connected.
Material and Methods
Host-seeking nymphal I. ricinus were collected by blanket dragging from vegetation in the campus of the Slovak Academy of Sciences (SAS) in Bratislava, 48.17°N, 17.07°E, altitude about 210 m, in April-May 2014 and at the beginning of September 2014. The SAS campus is a fenced area of 32 ha located on the south-western foothills of the Small Carpathians. It is characterized by patches of the original oak-hornbeam forest with admixture of beech, ash, black locust, maple, lime tree, elm, alder, and common hazel which are fragmented by roads, pavements, and built-up areas. No specific permissions were required for questing tick collections in this location as the SAS campus is not a protected area. The field study did not involve any endangered nor protected species.
Parasitoid wasp eclosion
Nymphs collected in April-May 2014 were subsequently fed on laboratory Balb/c mice. Thirty I. ricinus nymphs were placed in a retaining chamber and glued on the shaved back of one mouse. Engorged nymphs were collected to 15 ml tubes closed with nylon mesh and perforated lid. Tubes were incubated in a desiccator at room temperature and 85% relative humidity. Engorged nymphs were evaluated for the presence of parasitoid wasps, and/or moulting to adult ticks. The time period from the detachment of engorged nymphs to the emergence of parasitoid wasps ranged from 20 to 30 days. The parasitoids that emerged were separated into test tubes and deep-frozen at -80°C until subsequent analyses. The sex ratio (males to females) of adult wasps was evaluated by microscopic examination.
Extraction of DNA
DNA was extracted from emerged I. hookeri wasps and host-seeking I. ricinus nymphs collected in September 2014, using an EZ1 Advanced XL automated extractor (Qiagen), and Qiagen manufacturer’s kit, according to supplier’s instructions. DNA was stored at 4°C until further use.
Polymerase chain reaction
Based on cytochrome oxidase subunit I sequences of I. hookeri available in GenBank (JQ315225.1), we designed primers and a Taqman probe for Real-Time PCR to specifically detect I. hookeri: Iphag583f 5′-TTGCTGTTCCAACAGGAGTAAA-3′ and Iphag820r 5′-CAAAAAATTGCAAAAACTGC-3′ and probe Iphag612s 6FAM®-AGATGATAAGCTTCAATAAATGGAA-TAMRA®. DNA extracted from I. hookeri (obtained from parasitized I. ricinus nymphs in 2013) served as a positive control when using the primers and probe targeting parasitoid DNA in ticks. The set of primers/probe was verified for specificity with 30 negative controls (DNAs extracted from 20 bacterial, 5 arthropod and 5 vertebrate species).
PCRs were carried out in a CFX 96 Real-Time system C 1000 Thermal Cycler controlled by the vendor software (BioRad). The 20 μl of PCR mixture included 10 μl of the Takyon No ROX Probe 2x MasterMix UNG (Eurogentec), 0.5 μl (20 pmol.μl-1) of each primer, 0.5 μl (20 pmol.μl-1) of probe, 3.5 μl of milliQ water and 5 μl of extracted DNA. The amplification conditions were as follows: an initial denaturation step at 95°C for 3 min, followed by 40 cycles of denaturation at 95°C, annealing and elongation at 60°C for 60 s, with fluorescence acquisition in single mode. To avoid false-negative results, each sample was run in technical triplicate. A mean cycle threshold (Ct) value below 35 indicated the sample as positive, and a Ct value above 35 indicated the sample as positive only if another two sets were positive .
Arsenophonus nasoniae was detected by rpoB gene-based Real-Time PCR  in emerged parasitoid wasps and unfed I. ricinus nymphs. As a positive control we used DNA extracted from A. nasoniae cultured on Columbia agar .
To screen wasps and nymphs for the presence of all spotted fever group rickettsiae (SFG), we used a previously published PCR assay with Rickettsia-specific gltA gene–based RKND03 system . If the first screening was positive, a second directed step of molecular screening was performed to target rickettsiae at the species level using various sets of primers and probes [16;19]. DNA extracted from the cell culture supernatant of Rickettsia montanensis served as a positive control when using the primer and probe set targeting SFG Rickettsia; DNA extracted from the cell-culture supernatant of each particular Rickettsia species served as a positive control for the corresponding primer and probe set.
The usage of animals in the experiment was approved by the State Veterinary and Food Administration of the Slovak Republic (permit number 1335/12-221). The mice were euthanized at the end of the study by cervical dislocation. The experiments were performed under standard conditions in the experimental animal facility of the Institute of Virology, Slovak Academy of Sciences (Permit number: SK P 01014). No specific permissions were required for locations where collecting of questing ticks was carried out. The field studies did not involve any endangered nor protected species.
We screened for bacterial positivity I. hookeri adults that emerged from engorged I. ricinus nymphs, collected in questing stage from vegetation in April-May 2014 and subsequently fed on laboratory mice, and in unfed host-seeking nymphs collected in September 2014.
Engorged tick nymphs and emerged I. hookeri wasps
A total of 360 I. ricinus nymphs (Fig 1), used for infestation of laboratory mice, were engorged and detached. Of those 50 nymphs (13.8%) were parasitized by I. hookeri. Numbers of wasps emerging from each individual nymph were not exactly evaluated but, they were ranging from 2–20. We collected the adult wasps soon after emerging and stored at -80°C for further purposes.
A total of 360 I. ricinus nymphs were fed on 12 Balb/c mice. Of those, 50 engorged nymphs were parasitized by I. hookeri wasps. The obtained 96 parasitoids were subsequently screened for the presence of A. nasoniae and rickettsiae by PCR. The DNA of A. nasoniae was found in 27 wasps (28.1%), Rickettsia sp. in 22 wasps (22.9%). Eight wasps were positive for both bacteria—A. nasoniae and Rickettsia sp.
Determination of the sex of the wasps was done by microscopic examination. Emerged females were mainly black coloured and their length was around 1 mm. Males resembled females beside differences in genitalia and antennae (Fig 2). The sex ratio (males to females) of adult wasps was 1:3.6. We identified 19 males, 59 females, but were not able to assign the sex of 8 wasps (Fig 1), which was likely due to improper handling.
Ixodiphagus hookeri harbours the endosymbiotic bacterium Arsenophonus nasoniae, and Rickettsia sp.
All I. hookeri obtained from parasitized nymphs were screened for the presence of A. nasoniae using specific primers for its rpoB gene. DNA of this bacterium was detected by Real-Time PCR, in 28.1% of the parasitoids, mainly females. Surprisingly, A. nasoniae was also identified in four male wasps, despite of its well-known reproductive parasitism causing male offspring mortality.
Furthermore, all 96 wasps were tested for the presence of Rickettsia species. We found that 22.9% of examined wasps contained DNA of rickettsiae (Fig 1). After amplification with specific primers, we detected Rickettsia helvetica in 13.5% (4 males, 8 females, 1 sex non identifiable) and Rickettsia monacensis in 9.4% (2 males, 6 females, 1 sex non identifiable) of wasps. Interestingly, eight parasitoids were positive for the presence of both A. nasoniae and Rickettsia sp. (R. helvetica or R. monacensis).
Bacterial infection rates in the natural population of I. ricinus nymphs due to presence of I. hookeri
DNA was extracted from 41 host-seeking I. ricinus nymphs, collected from vegetation in September 2014 (Fig 3). They were directly tested for the presence of the bacteria, A. nasoniae and/or Rickettsia sp., as well as for the presence of I. hookeri DNA by Real-Time PCR. Out of all investigated host-seeking nymphs, 14.6% contained wasp’s DNA, which was comparable to the value, obtained for engorged nymphs (13.8%) that originated from the same collection site. Strikingly, we identified A. nasoniae in a lower percentage of questing nymphs (9.8%) than in the emerged parasitic wasps (28.1%).
Overall 41 host-seeking I. ricinus nymphs were examined for the presence of A. nasoniae and Rickettsia sp. as well as for I. hookeri by PCR. The DNA of A. nasoniae was successfully amplified in 4 nymphs (9.8%) and the presence of rickettsiae was confirmed in 5 nymphs (12.2%). Six nymphs were parasitized by I. hookeri wasps (14.6%). As shown in the Venn-diagram, all nymphs that were positive for A. nasoniae were simultaneously parasitized by I. hookeri. Co-infection by both bacteria occurred only in one I. ricinus nymph which also contained wasp DNA.
All nymphs that were positive for A. nasoniae also hosted I. hookeri, suggesting that the presence of the bacterium depends not on the developmental stage of the tick but on the parasitisation of I. ricinus by the wasp. Two nymphs that were positive for I. hookeri were negative for A. nasoniae. Summing up, not all wasps harboured A. nasoniae but this bacterium was detected exclusively in those I. ricinus nymphs that carried I. hookeri. Thus, we may hypothesise, that the presence of I. hookeri in I. ricinus nymphs is correlated with the presence of Arsenophonus nasoniae.
The prevalence of I. ricinus nymphs infected by rickettsiae was 12.2% (Fig 3). One of these nymphs was found to harbour both A. nasoniae and Rickettsia sp., and was simultaneously parasitized by I. hookeri.
In this study we focused our attention to three parasitic organisms of ticks: the wasp I. hookeri, and the bacteria A. nasoniae and Rickettsia sp.
Among arthropod vectors, ticks harbour the largest diversity of microorganisms, ranging from viruses (tick-borne encephalitis), to bacteria (Rickettsia sp., Borrelia spp., species from Anaplasmataceae family, etc.) and/or eukaryotes (Babesia sp., Theileria sp.) [20;21]. Ticks are also hosts of macroparasites; including wasps such as I. hookeri (Table 1), which carry their own microbiome . Normally, many insect species are simultaneously infected by multiple microbial symbionts, which in turn interact with each other, an co-regulate the biological processes of the host .
The parasitisation rate of the I. ricinus nymphs, fed on laboratory Balb/c mice, by I. hookeri was 13.8%. Similarly, we found that 14.6% of questing I. ricinus nymphs contained wasps DNA. This agrees with the parasitation rates reported by Hu et al.  and Stafford et al. , but is higher compared to the investigated occurrence in Germany  and Italy . The parasitisation rate of engorged nymphs might be even higher, since ticks are more likely parasitized while feeding on their vertebrate hosts ; wasp females visually evaluate and choose feeding nymphs over questing nymphs, because a feeding nymph may be an immediately available source of a meal for parasitoid larvae .
We discovered that natural populations of the wasp I. hookeri are infected at a 28.1% prevalence by A. nasoniae. Remarkably, A. nasoniae has never been reported to be associated with I. hookeri wasps before. This bacterium is well-known for its unique evolutionary evolved male-killing phenomenon, eliminating the male offspring of the wasps [3;6]. This is achieved by inhibition of the production of maternal centrosomes, organelles required specifically for early male embryonic development—male arise from unfertilized, haploid eggs and obtain their centrosomes maternally . In that context we also found that the sex ratio of emerging adult parasitoids was strongly female biased (19♂:69♀). The obtained data were similar to those reported by Davis et al. , or Collatz et al. . Mostly female wasps (77.78%, 1:5.25 ratio of infected males to females, in two instances sex non identifiable) were positive for A. nasoniae. We were able to identify only four adult males to be positive for A. nasoniae using Real-Time PCR.
Arsenophonus nasoniae has been identified in seven hard tick species (Table 2), but thus far there has been no clear understanding of the nature of the bacterium-host relationship in ticks. Using primers specific to I. hookeri and A. nasoniae, we showed that in an I. ricinus population, all individual ticks harbouring A. nasoniae were parasitized by I. hookeri, while ticks without wasps were Arsenophonus-free. Keeping in mind, that the simple detection of a vector-borne bacterial agent in a parasite does not demonstrate vector competency , it seems that the presence of A. nasoniae may be linked to the presence of wasp parasitoids in those ticks.
This assumption was already foreseen in our earlier study, in which the infection rate was comparable, namely, A. nasoniae was identified in 20 I. ricinus individuals; only one adult tick out of 28 and 19 nymphs out of 52 were positive when screened by specific PCR . We proposed that the presence of A. nasoniae in ticks depends on their developmental stage. In the current study we hypothesise that A. nasoniae is present only due to I. hookeri parasitoids.
Several authors already proved [25;42;54] that larvae of the I. hookeri wasp consume the entire tissue contents of the engorged nymphal tick, including the blood meal ingested by the host nymph, which may be infected by a variety of microorganisms. In the present study we obtained important data about the presence of rickettsiae in emerged I. hookeri wasps. We detected R. helvetica, and/ or R. monacensis for the first time in adults of this parasitoid species. Almost 23% Rickettsia-positive individuals were confirmed by Real-Time PCR.
However, we do not know whether the detected rickettsial DNA originated from consumed Rickettsia-infected tick cells or viable rickettsiae (Fig 4). Follow-up studies are needed to clarify the capability of Ixodiphagus wasps to carry viable rickettsiae, thus to transmit them to ticks transovarially.
Adult female I. hookeri oviposits in larvae and nymphs of ixodid ticks, but the wasp eggs start to develop only in fully engorged nymphs. The immature parasitoid wasps consume the nymph’s tissue and its ingested blood meal, causing nymph death. During the tick’s life cycle (eggs, larvae, nymphs, adults), rickettsiae can pass from stage to stage. In our experiments we successfully amplified rickettsial DNA not just in unfed nymphs but also in emerged adult wasps. More experiments will be needed to demonstrate if I. hookeri may act as a biological vector of A. nasoniae and Rickettsia sp.
We detected A. nasoniae in adult I. hookeri wasps for the first time. The natural population of the I. hookeri wasp was infected by A. nasoniae at a 28.1% prevalence. The parasitisation rate of I. ricinus nymphs originating from the same natural site, fed or unfed, by I. hookeri was comparable (13.8% and 14.6%, respectively). Nymphs that were not parasitized by wasps were Arsenophonus-free. Unique in our study was also the molecular detection of rickettsial DNA in 23% of I. hookeri adults. However, the transmission of viable rickettsiae from tick to tick by a wasp deserves further investigation.
We thank Dr. Jean-Michel Bérenger for assistance with microscopy of I. hookeri wasps.
Conceived and designed the experiments: MB OM MK DR ZS. Performed the experiments: MB OM MK ZS. Analyzed the data: MB OM MK DR ZS. Contributed reagents/materials/analysis tools: MB OM MK DR ZS. Wrote the paper: MB OM MK DR ZS.
- 1. Wilkes TE, Darby AC, Choi JH, Colbourne JK, Werren JH, Hurst GDD. The draft genome sequence of Arsenophonus nasoniae, son-killer bacterium of Nasonia vitripennis, reveals genes associated with virulence and symbiosis. Insect Molecular Biology 2010 Feb;19:59–73. pmid:20167018
- 2. Darby AC, Choi JH, Wilkes T, Hughes MA, Werren JH, Hurst GDD, et al. Characteristics of the genome of Arsenophonus nasoniae, son-killer bacterium of the wasp Nasonia. Insect Molecular Biology 2010 Feb;19:75–89.
- 3. Werren JH, Skinner SW, Huger AM. Male-killing bacteria in a parasitic wasp. Science 1986 Feb 28;231(4741):990–2. pmid:3945814
- 4. Gherna RL, Werren JH, Weisburg W, Cote R, Woese CR, Mandelco L, et al. Arsenophonus nasoniae gen. nov., sp. nov., the causative agent of the son-killer trait in the parasitic wasp Nasonia vitripennis. International Journal of Systematic Bacteriology 1991 Oct;41(4):563–5.
- 5. Skinner SW. Son-killer: a third extrachromosomal factor affecting the sex ratio in the parasitoid wasp, Nasonia (= Mormoniella) vitripennis. Genetics 1985 Apr;109(4):745–59. pmid:3988039
- 6. Huger AM, Skinner SW, Werren JH. Bacterial infections associated with the son-killer trait in the parasitoid wasp Nasonia (= Mormoniella) vitripennis (Hymenoptera: Pteromalidae). J Invertebr Pathol 1985 Nov;46(3):272–80. pmid:4067323
- 7. Taylor GP, Coghlin PC, Floate KD, Perlman SJ. The host range of the male-killing symbiont Arsenophonus nasoniae in filth fly parasitioids. J Invertebr Pathol 2011 Mar;106(3):371–9. pmid:21147118
- 8. Balas MT, Lee MH, Werren JH. Distribution and fitness effects of the son-killer bacterium in Nasonia. Evolutionary Ecology 1996 Nov;10(6):593–607.
- 9. Bouček Z, Černý V. Cizopasník klístat, chalcidka Hunterellus hookeri How. v CSR. Folia Zoologica Entomologica 1954;3(18):109–11.
- 10. Hu R, Hyland KE. Effects of the feeding process of Ixodes scapularis (Acari: Ixodidae) on embryonic development of its parasitoid, Ixodiphagus hookeri (Hymenoptera: Encyrtidae). J Med Entomol 1998 Nov;35(6):1050–3. pmid:9835701
- 11. Davis AJ. Bibliography of the Ixodiphagini (Hymenoptera, Chalcidoidea, Encyrtidae), parasites of ticks (Acari, Ixodidae), with notes on their biology. Tijdschrift Voor Entomologie 1986 Dec 15;129(6):181–90.
- 12. Davis AJ, Campbell A. Ixodiphagus texanus Howard (Hymenoptera: Encyrtidae), a parasite of the rabbit tick in Nova Scotia. Canadian Journal of Zoology 1979;57(5):1164–6.
- 13. Bergey's Manual. Bergey's Manual of Systematic Bacteriology. 2nd ed. New York: Springer; 2005.
- 14. Parola P, Raoult D. Tick-borne bacterial diseases emerging in Europe. Clin Microbiol Infect 2001 Feb;7(2):80–3. pmid:11298147
- 15. Parola P, Paddock CD, Socolovschi C, Labruna MB, Mediannikov O, Kernif T, et al. Update on tick-borne rickettsioses around the world: a geographic approach (vol 26, pg 657, 2013). Clinical Microbiology Reviews 2014 Jan;27(1):166.
- 16. Renvoise A, Rolain JM, Socolovschi C, Raoult D. Widespread use of real-time PCR for rickettsial diagnosis. FEMS Immunol Med Microbiol 2012 Feb;64(1):126–9. pmid:22092999
- 17. Mediannikov O, Subramanian G, Sekeyova Z, Bell-Sakyi L, Raoult D. Isolation of Arsenophonus nasoniae from Ixodes ricinus ticks in Slovakia. Ticks Tick Borne Dis 2012 Dec;3(5–6):367–70. pmid:23182269
- 18. Socolovschi C, Mediannikov O, Sokhna C, Tall A, Diatta G, Bassene H, et al. Rickettsia felis-associated uneruptive fever, Senegal. Emerg Infect Dis 2010 Jul;16(7):1140–2. pmid:20587190
- 19. Sekeyova Z, Fournier PE, Rehacek J, Raoult D. Characterization of a new spotted fever group rickettsia detected in Ixodes ricinus (Acari: Ixodidae) collected in Slovakia. J Med Entomol 2000 Sep;37(5):707–13. pmid:11004782
- 20. Jongejan F, Uilenberg G. The global importance of ticks. Parasitology 2004;129 Suppl:S3–14. pmid:15938502
- 21. Plantard O, Bouju-Albert A, Malard MA, Hermouet A, Capron G, Verheyden H. Detection of Wolbachia in the tick Ixodes ricinus is due to the presence of the hymenoptera endoparasitoid Ixodiphagus hookeri. PLoS One 2012;7(1):e30692. pmid:22292021
- 22. Chaves S, Neto M, Tenreiro R. Insect-symbiont systems: from complex relationships to biotechnological applications. Biotechnol J 2009 Dec;4(12):1753–65. pmid:19844913
- 23. Ramos RA, Campbell BE, Whittle A, Lia RP, Montarsi F, Parisi A, et al. Occurrence of Ixodiphagus hookeri (Hymenoptera: Encyrtidae) in Ixodes ricinus (Acari: Ixodidae) in Southern Italy. Ticks Tick Borne Dis 2015 Jan 8.
- 24. Tijsse-Klasen E, Braks M, Scholte EJ, Sprong H. Parasites of vectors—Ixodiphagus hookeri and its Wolbachia symbionts in ticks in the Netherlands. Parasites & Vectors 2011 Dec 7;4.
- 25. Collatz J, Selzer P, Fuhrmann A, Oehme RM, Mackenstedt U, Kahl O, et al. A hidden beneficial: biology of the tick-wasp Ixodiphagus hookeri in Germany. Journal of Applied Entomology 2011 Jun;135(5):351–8.
- 26. Collatz J, Fuhrmann A, Selzer P, Oehme RM, Hartelt K, Kimmig P, et al. Being a parasitoid of parasites: host finding in the tick wasp Ixodiphagus hookeri by odours from mammals. Entomologia Experimentalis et Applicata 2010 Feb;134(2):131–7.
- 27. Rehacek J. Uzitocny cudzopasnik. Enviromagazin 1998;3(2):19.
- 28. Rehacek J, Kocianova E. Attempt to infect Hunterellus hookeri Howard (Hymenoptera, Encyrtidae), an endoparasite of ticks, with Coxiella burnetti. Acta Virol 1992 Oct;36(5):492. pmid:1364029
- 29. Stafford KC III, Denicola AJ, Kilpatrick HJ. Reduced abundance of Ixodes scapularis (Acari: Ixodidae) and the tick parasitoid Ixodiphagus hookeri (Hymenoptera: Encyrtidae) with reduction of white-tailed deer. J Med Entomol 2003 Sep;40(5):642–52. pmid:14596277
- 30. Knipling EF, Steelman CD. Feasibility of controlling Ixodes scapularis ticks (Acari: Ixodidae), the vector of Lyme disease, by parasitoid augmentation. J Med Entomol 2000 Sep;37(5):645–52. pmid:11004774
- 31. Stafford KC, Denicola AJ, Magnarelli LA. Presence of Ixodiphagus hookeri (Hymenoptera: Encyrtidae) in two Connecticut populations of Ixodes scapularis (Acari: Ixodidae). Journal of Medical Entomology 1996 Jan;33(1):183–8. pmid:8906928
- 32. Mather TN, Piesman J, Spielman A. Absence of spirochaetes (Borrelia burgdorferi) and piroplasms (Babesia microti) in deer ticks (Ixodes dammini) parasitized by chalcid wasps (Hunterellus hookeri). Med Vet Entomol 1987 Jan;1(1):3–8. pmid:2979518
- 33. Doube BM, Heath AC. Observations on the biology and seasonal abundance of an encyrtid wasp, a parasite of ticks in Queensland. J Med Entomol 1975 Oct 31;12(4):433–47. pmid:1195291
- 34. Takasu K, Nakamura S. Life history of the tick parasitoid Ixodiphagus hookeri (Hymenoptera: Encyrtidae) in Kenya. Biological Control 2008 Aug;46(2):114–21.
- 35. Demas FA, Hassanali A, Mwangi EN, Kunjeku EC, Mabveni AR. Cattle and Amblyomma variegatum odors used in host habitat and host finding by the tick parasitoid, Ixodiphagus hookeri. Journal of Chemical Ecology 2000 Apr;26(4):1079–93.
- 36. Mwangi EN, Hassan SM, Kaaya GP, Essuman S. The impact of Ixodiphagus hookeri, a tick parasitoid, on Amblyomma variegatum (Acari: Ixodidae) in a field trial in Kenya. Exp Appl Acarol 1997 Feb;21(2):117–26. pmid:9080682
- 37. Demas FA, Mwangi EN, Hassanali A, Kunjeku EC, Mabveni AR. Visual evaluation and recognition of hosts by the tick parasitoid, Ixodiphagus hookeri (Hymenoptera: Encyrtidae). Journal of Insect Behavior 2002 Jul;15(4):477–94.
- 38. Lopes AJ, Nascimento-Junior JR, Silva CG, Prado AP, Labruna MB, Costa-Junior LM. Parasitism by Ixodiphagus wasps (Hymenoptera: Encyrtidae) in Rhipicephalus sanguineus and Amblyomma ticks (Acari: Ixodidae) in three regions of Brazil. J Econ Entomol 2012 Dec;105(6):1979–81. pmid:23356061
- 39. Howard LO. Another chalcidoid parasite of a tick. Canadian Entomologist 1908 Jul 2;40(7):239–41.
- 40. Philip CB. Occurrence of a colony of the tick parasite Hunterellus hookeri Howard in West Africa. Public Health Reports, Washington D C 1931;46(37):2168–72.
- 41. Slovak M. Finding of the endoparasitoid Ixodiphagus hookeri (Hymenoptera, Encyrtidae) in Haemaphysalis concinna ticks in Slovakia. Biologia 2003 Oct;58(5):890.
- 42. Hu R, Hyland KE, Mather TN. Occurrence and distribution in Rhode Island of Hunterellus hookeri (Hymenoptera: Encyrtidae), a wasp parasitoid of Ixodes dammini. J Med Entomol 1993 Jan;30(1):277–80. pmid:8433338
- 43. Ferree PM, Avery A, Azpurua J, Wilkes T, Werren JH. A bacterium targets maternally inherited centrosomes to kill males in Nasonia. Curr Biol 2008 Sep 23;18(18):1409–14. pmid:18804376
- 44. Sparagano O, Giangaspero A. Parasitism in egg production systems: the role of the red mite (Dermanyssus gallinae). Improving the safety and quality of eggs and egg products: Egg chemistry, production and consumption.Cambridge: Woodhead Publishing Limited; 2011. p. 394–414.
- 45. Rynkiewicz EC, Hemmerich C, Rusch DB, Fuqua C, Clay K. Concordance of bacterial communities of two tick species and blood of their shared rodent host. Molecular Ecology 2015 May;24(10):2566–79. pmid:25847197
- 46. Kagemann J, Clay K. Effects of Infection by Arsenophonus and Rickettsia bacteria on the locomotive ability of the ticks Amblyomma americanum, Dermacentor variabilis, and Ixodes scapularis. Journal of Medical Entomology 2013 Jan;50(1):155–62. pmid:23427665
- 47. Ahantarig A, Trinachartvanit W, Baimai V, Grubhoffer L. Hard ticks and their bacterial endosymbionts (or would be pathogens). Folia Microbiologica 2013 Sep;58(5):419–28. pmid:23334948
- 48. Grindle N, Tyner JJ, Clay K, Fuqua C. Identification of Arsenophonus-type bacteria from the dog tick Dermacentor variabilis. Journal of Invertebrate Pathology 2003 Jul;83(3):264–6. pmid:12877836
- 49. Liu L, Li L, Liu J, Hu Y, Liu Z, Guo L, et al. Coinfection of Dermacentor silvarum olenev (acari: ixodidae) by Coxiella-Like, Arsenophonus-like, and Rickettsia-like symbionts. Appl Environ Microbiol 2013 Apr;79(7):2450–4. pmid:23354701
- 50. Dergousoff SJ, Chilton NB. Detection of a new Arsenophonus-type bacterium in Canadian populations of the Rocky Mountain wood tick, Dermacentor andersoni. Exp Appl Acarol 2010 Sep;52(1):85–91. pmid:20186465
- 51. Subramanian G, Sekeyova Z, Raoult D, Mediannikov O. Multiple tick-associated bacteria in Ixodes ricinus from Slovakia. Ticks Tick Borne Dis 2012 Dec;3(5–6):406–10. pmid:23182274
- 52. Liu LM, Liu JN, Liu Z, Yu ZJ, Xu SQ, Yang XH, et al. Microbial communities and symbionts in the hard tick Haemaphysalis longicornis (Acari: Ixodidae) from north China. Parasit Vectors 2013;6(1):310. pmid:24499619
- 53. Clay K, Klyachko O, Grindle N, Civitello D, Oleske D, Fuqua C. Microbial communities and interactions in the lone star tick, Amblyomma americanum. Molecular Ecology 2008 Oct;17(19):4371–81. pmid:19378409
- 54. Mwangi EN, Kaaya GP, Essuman S, Kimondo MG. Parasitism of Ambyomma variegatum by a hymenopteran parasitoid in the laboratory, and some aspects of its basic biology. Biological Control 1994;4(2):101–4.