Investigation of vertical and horizontal transmission of Spiroplasma in ticks under laboratory conditions

Scientific Reports volume 13, Article number: 13265 (2023) Cite this article

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Many arthropods harbour bacterial symbionts, which are maintained by vertical and/or horizontal transmission. Spiroplasma is one of the most well-known symbionts of ticks and other arthropods. It is still unclear how Spiroplasma infections have spread in tick populations despite its high prevalence in some tick species. In this study, Ixodes ovatus, which has been reported to harbour Spiroplasma ixodetis at high frequencies, was examined for its vertical transmission potential under experimental conditions. Next, two isolates of tick-derived Spiroplasma, S. ixodetis and Spiroplasma mirum, were experimentally inoculated into Spiroplasma-free Haemaphysalis longicornis colonies and the presence of Spiroplasma in their eggs and larvae was tested. Our experimental data confirmed that S. ixodetis was transmitted to eggs and larvae in a vertical manner in the original host I. ovatus. In the second experiment, there was no significant difference in engorged weight, egg weight, and hatching rate between Spiroplasma-inoculated and control H. longicornis groups. This suggested that Spiroplasma infection does not affect tick reproduction. Spiroplasma DNA was only detected in the eggs and larvae derived from some individuals of S. ixodetis-inoculated groups. This has demonstrated the potential of horizontal transmission between different tick species. These findings may help understand the transmission dynamics of Spiroplasma in nature and its adaptation mechanism to host arthropod species.

Ixodid ticks are blood-feeding ectoparasites of vertebrates with approximately 700 species distributed worldwide1. They serve as vectors for many pathogens and cause significant global public health and veterinary problems2,3. Ixodid ticks attach to their hosts for days or weeks while feeding on the blood. Depending on the number of hosts that they infest during their life cycle, they are divided into one-, two-, and three-host ticks. One-host ticks attach to a single host during their life cycle, while two- and three-host ticks drop off the host after feeding to moult in the environment between stages4,5,6,7,8.

The term ‘symbiosis’ was first defined in 1879 to describe the condition where different species live together in close association9. Symbiotic relationships are important in biological processes and ecological systems10. In nature, symbiotic relationships between bacteria and arthropods are well known and have been extensively studied10. Symbionts use disparate transmission strategies to allow their survival in their hosts11. Some symbionts such as Wolbachia and Arsenophonus are transmitted vertically in host arthropods12. Meanwhile, others use horizontal transmission routes, for instance, by being acquired from environmental or infected conspecifics or other species13,14. In some cases, a combination of multiple infection routes creates complex transmission dynamics in nature15,16. The dominant transmission form of endosymbionts is vertical transmission, which occurs primarily from the mother to offspring17. Some insect symbionts display specialised transmission strategies, for instance, via parental post-oviposition secretions18.

Members of the genus Spiroplasma are helical bacteria that lack cell walls. It is estimated that 5–10% of arthropods harbour Spiroplasma as their symbionts19,20. Many species of Spiroplasma maintain their infection in their hosts through vertical transmission21. In Drosophila, Spiroplasma uses yolk uptake machinery to move into the germline for vertical transmission22. It has recently been found that some of these vertically transmitted Spiroplasma confer protection against nematodes, parasitoid wasps, and fungi to their hosts21. Although vertical transmission of Spiroplasma has been confirmed in some species21, phylogenetic studies have reported poor clustering of Spiroplasma including from the same host species. This suggests that horizontal transmission between unrelated hosts occurs frequently21,23,24.

Ticks generally harbour maternally inherited bacterial endosymbionts25,26. Some of these endosymbionts, such as Coxiella and Francisella are likely essential for the life cycle of ticks27. Ticks have also been shown to harbour Spiroplasma28,29. How Spiroplasma infection spreads among tick populations is still unclear despite high infection rates being observed in some tick species, such as Ixodes ovatus and Haemaphysalis kitaokai30.

In this study, the vertical transmission of Spiroplasma was demonstrated using field-collected I. ovatus, which harbours Spiroplasma at high frequencies. In addition, the horizontal transmission potential of Spiroplasma was examined by experimentally inoculating Spiroplasma strains isolated from ticks into Spiroplasma-free laboratory tick colonies.

Out of 30 I. ovatus females (mated beforehand with males) used to infest rabbits, 14 ticks became fully engorged and detached from the animals eight days after infestation. Nine of them laid eggs approximately nine days after detachment, while oviposition was not observed in five engorged ticks.

DNA extracted from three pools of eggs and three pools of larvae per tick were used for detection of Spiroplasma by PCR and Sanger sequencing. Spiroplasma ixodetis DNA was detected in all the egg and larval pools tested. Spiroplasma infection was also confirmed in all the engorged I. ovatus females from which the eggs and larvae tested were obtained.

Females of H. longicornis were injected with either PBS, S. ixodetis, or S. mirum with or without antibiotics (penicillin G sodium salt). Out of 70 ticks used, a total of 63 ticks were alive after the injection. Seven ticks were dead during the incubation period seven days after injection. The average engorgement weight was calculated for 3–7 individuals in each group and fell in the range of 87.4–262.2 mg. The largest engorgement weight was observed for the S. ixodetis (5 × 1011 bacteria) + penicillin G sodium salt injection group (Fig. 1, S. ixodetis H_PG). Meanwhile, the smallest was for the S. mirum (5 × 1011 bacteria) injection group (Fig. 1, S. mirum H_wo). There were no statistically significant differences in engorgement weights between the groups, including those with and without antibiotics, by the Kruskal–Wallis test (Fig. 1). The egg production efficiency was calculated as the laid egg weight divided by the engorged body weight31. The average calculated for 3–7 individuals in each group fell in the range of 33.1–57.1. The largest egg production efficiency was observed in the S. mirum (5 × 108 bacteria) injection group (Table 1, SM_Low_wo), whereas the smallest was observed in the S. ixodetis (5 × 1011 bacteria) injection group (Table 1, SI_High_wo). There were no statistically significant differences in the egg production efficiency between the groups, including those with and without antibiotics. The hatching rate was calculated for 3–7 individuals in each group and fell in the range of 3.5–98.8%. The highest median hatching rate was observed for the S. ixodetis (5 × 108 bacteria) injection group (Table 1, SI_Low_wo). Meanwhile, the lowest was observed for the S. mirum (5 × 1011 bacteria) + penicillin G sodium salt injection group (Table 1, SM_High_PG). There were no statistically significant differences in the hatching rate among the groups, including those with and without antibiotics (Fig. 1, Table 1).

Engorged weight of ticks after the experimental inoculations. PBS_PG is PBS + penicillin G sodium salt. H_wo is 5 × 1011 bacteria. H_PG is 5 × 1011 bacteria + penicillin G sodium salt. L_wo is 5 × 108 bacteria. L_PG is 5 × 108 bacteria + penicillin G sodium salt. The white circle in the figure represents the average engorged weight. The black circle in the figure represents each value.

The presence of Spiroplasma in the eggs was tested by PCR using three egg pools per tick. In total, egg pools from two ticks comprising three pools each in the S. ixodetis (5 × 1011 bacteria) injection group and one tick (two pools) in the S. ixodetis (5 × 1011 bacteria) + penicillin G sodium salt injection group were positive for Spiroplasma infection (Table 1). The rest of the groups tested negative by Spiroplasma-specific PCR. No Spiroplasma was detected in any of the S. mirum injection groups.

The presence of Spiroplasma in the larvae was tested using PCR with three larval pools per individual tick. In total, larvae pools from two ticks comprising three pools each in the S. ixodetis (5 × 1011 bacteria) injection group and one tick (two pools) in the S. ixodetis (5 × 1011 bacteria) + penicillin G sodium salt injection group were positive for Spiroplasma infection (Table 1). The rest of the groups were all negative by Spiroplasma-specific PCR. No Spiroplasma was detected in any of the S. mirum injection groups.

Spiroplasma has been identified as one of the core microbial taxa in the tick microbiome25. However, its transmission dynamics in nature remains unknown in ticks. This study used field-collected I. ovatus, a species harbouring Spiroplasma at high frequencies, for experimental infestation under laboratory settings. It was confirmed that Spiroplasma is transmitted to eggs and larvae. This study has provided the first experimental evidence that S. ixodetis is vertically transmitted in ticks.

Previously, it was suggested that horizontal transmission of Spiroplasma between mites and Drosophila (Drosophila nebulosa and Drosophila willistoni) could occur under experimental conditions32. Spiroplasma ixodetis has been detected in several arthropods, including some species of ticks. Phylogenetic analysis has suggested horizontal transmission between ticks and other arthropods24. In this study, S. ixodetis and S. mirum were experimentally inoculated into Spiroplasma-free H. longicornis. Spiroplasma was detected only in the S. ixodetis injection groups, but not in the S. mirum injection groups (Table 1). This result provides the first experimental evidence that S. ixodetis can be maintained in the non-native tick species, supporting the horizontal transmission potential of Spiroplasma among different tick species. Having said that, the lower frequency of Spiroplasma transmission may have resulted from the poor adaptation of the isolates to the new host tick species. Spiroplasma strains introduced into other species are often poorly transmitted from the mother to offspring in Drosophila33. Spiroplasma citri, which normally infects leafhoppers, grows well in D. melanogaster haemolymph but cannot access the oocyte and, therefore, is not vertically transmitted34. Although a number of interspecific transfers of Spiroplasma between Drosophila species have been confirmed35, the transfer of Spiroplasma from Drosophila to other arthropod species has rarely been documented. The vertical transmission rate of Spiroplasma in Drosophila was found to be severely affected by temperature36, indicating that environmental factors under laboratory settings, such as temperature, may have affected the results.

Spiroplasma poulsonii in Drosophila colonises host oocytes at specific stages, coinciding with vitellogenesis, and requires yolk transport and uptake machinery to achieve efficient vertical transmission22. Spiroplasma citri has been observed entering the salivary cells of the beet leafhopper by receptor-mediated endocytosis37. These findings suggest that Spiroplasma may have a general capacity to interact with the host endocytic machinery to ensure vertical transmission. Vitellogenin uptake through the vitellogenin receptor in the oocyte is an essential event for the progress of oogenesis in ticks, including H. longicornis38,39. Therefore, Spiroplasma in ticks may use the same mechanism as that observed in other arthropods. To further confirm this hypothesis, the localization of S. ixodetis in I. ovatus after the blood feeding up to oviposition needs to be clarified.

In this study, Spiroplasma was inoculated with antibiotics in several injection groups. However, no significant difference was observed in terms of engorged weight, egg weight, and hatching rate between the antibiotic-treated and non-treated groups. When the symbiont density in host insects has been experimentally decreased using antibiotics, host reproduction ability is reduced because of symbiont sorting upon vertical transmission40. In the present study, antibiotic treatment had no effect on the vertical transmission of Spiroplasma in ticks. Antibiotic treatment for ticks has been associated with several reproductive dysfunctions, such as reduced engorged and egg weight and hatching rate. This is because of dysbiosis of microbiota and reduction of endosymbionts41,42. In our preliminary experiments, ticks injected with 0.5 units of penicillin G sodium salt died within 24 h, indicating an adverse effect of antibiotics on tick physiology (data not shown). The lack of a pronounced effect from the antibiotic treatment in the current experiment may be partially explained by the lower concentration of antibiotics where the microbiota was not affected in the antibiotic-treated groups. Therefore, it is necessary to optimise the concentration and type of antibiotics to be administered as well as inoculation routes in the future to understand the interaction between Spiroplasma and tick microbiota.

Ixodes ovatus were collected by flagging the vegetation in Sapporo city, Japan (N 43.02 E 141.29) in May 2021 and identified morphologically under a stereomicroscope according to the standard morphological keys43. Field-collected ticks were transferred to Petri dishes and preserved in an incubator at 16 °C until use. Parthenogenetic laboratory colonies of H. longicornis Okayama strain (Fujisaki, 1978)44, maintained at the National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, Japan, were used in Spiroplasma infection experiments. This tick colony was confirmed to be Spiroplasma-free by Spiroplasma-specific PCR prior to the experiment.

Female and male ticks were placed in the same Petri dish a week before the experiment. Thirty female I. ovatus were attached to the ears of Japanese white rabbits (Slc: JW/CSK; Japan SLC, Shizuoka, Japan) using earbags. The attached ticks were allowed to feed until they became engorged and detached naturally. The fully engorged ticks were then collected and incubated in the dark at 25 °C with saturated humidity for oviposition (Fig. 2).

Overview of experimental flow of experimental infestation of field-collected Ixodes ovatus on rabbits.

The pools of eggs (n = 10) and larvae (n = 10) were collected from each tick and subjected to DNA extraction and Spiroplasma-specific PCR, as described below.

In this study, two species of Spiroplasma: S. ixodetis (strain 135) and S. mirum (strain Q35) were used. Spiroplasma ixodetis strain 135 was isolated from male Ixodes monospinosus using ISE6 cells in a previous study45. The isolate was grown in ISE6 cells received from the CEH Institute of Virology and Environmental Microbiology (Oxford, UK) with L-15B medium supplemented with 10% foetal bovine serum and 5% tryptose phosphate broth (Sigma-Aldrich, St. Louis, MO, USA) at 32 °C as described previously30 . Spiroplasma mirum strain Q35 was isolated from a female Ixodes pavlovsky collected at Urausu town, Japan (N 43.46, E 141.76) by co-culturing the tick homogenate with ISE6 cells. The bacterial species was identified by amplifying and sequencing the 16S rDNA sequence as previously reported (unpublished)45. The isolate was thereafter cultured using modified SP4 medium at 32 °C46. The culture medium was changed when the colour changed from red to yellow. The titration of Spiroplasma was conducted by counting the bacterial cells under a dark-field microscope. The final concentrations of Spiroplasma solution were adjusted to 1 × 109 and 1 × 1012 bacteria/µL using culture media prior to inoculation into the ticks.

Females of H. longicornis were attached to glass slides and injected with 0.5 µL of PBS or Spiroplasma solution through the fourth coxae using an IM 300 Microinjector (Narishige, Tokyo, Japan). There were 10 injection groups, namely the PBS groups, PBS only, and PBS + penicillin G sodium salt (Sigma-Aldrich, St. Louis, MO, USA); S. ixodetis injection groups, S. ixodetis (5 × 108 bacteria and 5 × 1011 bacteria) only and S. ixodetis (5 × 108 bacteria and 5 × 1011 bacteria) + penicillin G sodium salt; S. mirum injection groups, S. mirum (5 × 108 bacteria and 5 × 1011 bacteria) only and S. mirum (5 × 108 bacteria and 5 × 1011 bacteria) + penicillin G sodium salt. The final concentration of penicillin G sodium salt in the solution was 100 U/mL.

After inoculation, the ticks injected were left for seven days at 25 °C in an incubator. For rabbit infestation, a total of 70 ticks per injection group were attached to separate ears of Japanese white rabbits. The attached ticks were allowed to feed until they became engorged and detached naturally. The fully engorged ticks were then weighed and incubated in the dark at 25 °C with saturated humidity for oviposition. After oviposition, the eggs were weighed and incubated under the same conditions until hatching. The hatching rate was calculated by counting the number of hatched larvae among the selected pools of eggs (50–150 eggs/pool). Three pools of eggs (100 each) and larvae (100 each) from each tick were subjected to DNA extraction and Spiroplasma-specific PCR (Fig. 3).

Overview of experimental flow of Spiroplasma-injection into Haemaphysalis longicornis ticks and their infestation on rabbits.

The eggs, larvae, and post-oviposition females were homogenised using a BioMasher (Nippi, Tokyo, Japan), as described in the manufacturer’s protocol. Genomic DNA was extracted from the homogenates using a NucleoSpin® DNA Insect Kit (Macherey–Nagel GmbH & Co. KG, Düren, Germany), following the manufacturer’s guidelines. To detect Spiroplasma DNA, PCR amplification targeting 1028 bp of 16S rDNA was performed using the primers, spi_f1 (5′-GGGTGAGTAACACGTATCT-3′) and spi_r3 (5′-CCTTCCTCTAGCTTACACTA-3′)30. PCR was conducted in a 20 μL reaction mixture containing 10 μL of 2 × Gflex PCR Buffer (Mg2+, dNTP plus) (TaKaRa Bio Inc., Shiga, Japan), 400 nM of Tks Gflex™ DNA Polymerase, 400 nM of each primer, 1 μL of DNA template, and sterilised water. The reaction was performed at 94 °C for 1 min, followed by 45 cycles at 98 °C for 10 s, 60 °C for 30 s, and 68 °C for 45 s, and a final step at 68 °C for 5 min. PCR products were electrophoresed on a 1.0% agarose gel. The DNA of Spiroplasma species isolated from I. persulcatus in the previous study47 and sterilised water were included in each PCR run as positive and negative controls, respectively. The amplified PCR products were purified using the ExoSAP-IT Express PCR Cleanup Reagent (Thermo Fisher Scientific, Tokyo, Japan). Sanger sequencing was performed using a BigDye Terminator version 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA). Sequencing data were assembled using the ATGC software version 6.0.4 (GENETYX, Tokyo, Japan).

All the statistical analyses were performed using Microsoft 365 Excel. The Kruskal–Wallis test was used to confirm significant differences in engorged tick weight, egg weight, and egg hatching rate per injection group. R software (version 4.2.3) was used to create the graphs48.

All the animal experiments were carried out under the guidance of the Institute for Laboratory Animal Research (ILAR), which was based on Fundamental Guidelines for Proper Conduct of Animal Experiment and Related Activities in Academic Research Institutions under the jurisdiction of the Ministry of Education, Culture, Sports, Science and Technology, Japan. This experimental protocol was approved by the Committee on the Ethics of Animal Experiments of Hokkaido University (Approval No. 22-0030) and the Animal Care and Use Committee of Obihiro University of Agriculture and Veterinary Medicine (Approval Nos. 18-11, 19-74, 20-85, and 21-40). All experiments were performed in accordance with the guidelines of the committee. The study is reported in compliance with the ARRIVE guidelines.

This study is the first to experimentally confirm the vertical transmission of S. ixodetis in ticks using field-collected I. ovatus. In addition, Spiroplasma was detected in eggs and larvae originating from H. longicornis experimentally inoculated with S. ixodetis. This has indicated that S. ixodetis can be transferred into the non-native arthropod species via horizontal transmission. Further experiments are needed to evaluate the viability of the introduced Spiroplasma in the recipient hosts and the transmission efficacy to the next developmental stage or generation. These findings may help understand the transmission dynamics of Spiroplasma in nature, the adaptation mechanism to specific tick species, and ultimately the effects on host tick physiology.

The datasets used and/or analysed during the current study available from the corresponding author on reasonable request.

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We would like to thank all collaborators who supported sample collection. This research was supported by JSPS KAKENHI (19H03118, 20K21358, 20KK0151, and 22H02505), the Japan Program for Infectious Diseases Research and Infrastructure (21fk0108614j0001) from the Japan Agency for Medical Research and Development (AMED), and Cooperative Research Grant(29-joint-1, 30-joint-13, 2020-joint-1, 2021-joint-1, and 2022-joint-12) of the National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine.

Laboratory of Parasitology, Department of Disease Control, Graduate School of Infectious Diseases, Faculty of Veterinary Medicine, Hokkaido University, Sapporo, 060-0818, Japan

Shohei Ogata, Kodai Kusakisako, Keita Kakisaka, Elisha Chatanga, Naoki Hayashi, Yurie Taya, Yuma Ohari, Gita Sadaula Pandey, Abdelbaset Eweda Abdelbaset, Nariaki Nonaka & Ryo Nakao

Laboratory of Molecular Targeted Therapeutics, School of Pharmacy, Nihon University, Chiba, 274-8555, Japan

Shohei Ogata

Division of International Research Promotion, International Institute for Zoonosis Control, Hokkaido University, Sapporo, 001-0020, Japan

Shohei Ogata & Yongjin Qiu

National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, Obihiro, 080-8555, Japan

Rika Umemiya-Shirafuji

Laboratory of Veterinary Parasitology, School of Veterinary Medicine, Kitasato University, Towada, 034-8628, Japan

Kodai Kusakisako

Department of Veterinary Pathobiology, Faculty of Veterinary Medicine, Lilongwe University of Agriculture and Natural Resources, P.O. Box 219, Lilongwe, Malawi

Elisha Chatanga

School of Veterinary Medicine, Rakuno Gakuen University, Ebetsu, 069-8501, Japan

Yuma Ohari

Department of Animal Medicine, Clinical Laboratory Diagnosis, Faculty of Veterinary Medicine, Assiut University, Assiut, 71515, Egypt

Abdelbaset Eweda Abdelbaset

Department of Virology-I, National Institute of Infectious Diseases, Shinjuku-ku, Tokyo, 162-8640, Japan

Yongjin Qiu

Management Department of Biosafety, Laboratory Animal, and Pathogen Bank, National Institute of Infectious Diseases, Shinjuku-ku, Tokyo, 162-8640, Japan

Yongjin Qiu

Division of Risk Analysis and Management, International Institute for Zoonosis Control, Hokkaido University, Sapporo, 001-0020, Japan

Keita Matsuno

One Health Research Center, Hokkaido University, Sapporo, 001-0020, Japan

Keita Matsuno

International Collaboration Unit, International Institute for Zoonosis Control, Hokkaido University, Sapporo, 001-0020, Japan

Keita Matsuno

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Conceptualisation, S.O.; Data curation, R.U.S., K.K., and N.N.; Formal analysis, E.C., N.H., and Y.O.; Investigation, S.O., Y.T., G.S.P., A.E.A., and Y.Q.; Methodology, R.U.S., K.M., and R.N.; Project administration, R.U.S. and R.N.; Funding acquisition, R.N.; Resources, R.U.S., and N.R.; Software, K.M.; Supervision, R.N.; Visualization, S.O.; Writing-original draft, S.O.; Writing-review and editing, R.U.S., K.K., K.K., E.C., N.H., Y.T., Y.O., Y.Q., K.M., N.N., and R.N. All authors have read and agreed to the published version of the manuscript.

Correspondence to Ryo Nakao.

The authors declare no competing interests.

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Ogata, S., Umemiya-Shirafuji, R., Kusakisako, K. et al. Investigation of vertical and horizontal transmission of Spiroplasma in ticks under laboratory conditions. Sci Rep 13, 13265 (2023). https://doi.org/10.1038/s41598-023-39128-z

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Received: 01 May 2023

Accepted: 20 July 2023

Published: 15 August 2023

DOI: https://doi.org/10.1038/s41598-023-39128-z

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