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OPEN RNA activation in ticks
Kofi Dadzie Kwofie 1,2, Emmanuel Pacia Hernandez 1,3, Anisuzzaman 4, Hayato Kawada 1,5, 
Yuki Koike 5, Sana Sasaki 5, Takahiro Inoue 5, Kei Jimbo 5, Fusako Mikami 1, 
Danielle Ladzekpo 2,6, Rika Umemiya‑Shirafuji 7, Kayoko Yamaji 8, Tetsuya Tanaka 9, 
Makoto Matsubayashi 10, Md Abdul Alim 4, Samuel Kweku Dadzie 2, Shiroh Iwanaga 11,12, 
Naotoshi Tsuji 1,5 & Takeshi Hatta 1,5*
RNA activation (RNAa) is a burgeoning area of research in which double‑stranded RNAs (dsRNAs) 
or small activating RNAs mediate the upregulation of specific genes by targeting the promoter 
sequence and/or AU‑rich elements in the 3′‑ untranslated region (3’‑UTR) of mRNA molecules. So 
far, studies on the phenomenon have been limited to mammals, plants, bacteria, Caenorhabditis 
elegans, and recently, Aedes aegypti. However, it is yet to be applied in other arthropods, including 
ticks, despite the ubiquitous presence of argonaute 2 protein, which is an indispensable requirement 
for the formation of RNA‑induced transcriptional activation complex to enable a dsRNA‑mediated 
gene activation. In this study, we demonstrated for the first time the possible presence of RNAa 
phenomenon in the tick vector, Haemaphysalis longicornis (Asian longhorned tick). We targeted the 
3ʹ‑UTR of a novel endochitinase‑like gene (HlemCHT) identified previously in H. longicornis eggs 
for dsRNA‑mediated gene activation. Our results showed an increased gene expression in eggs 
of H. longicornis endochitinase‑dsRNA‑injected (dsHlemCHT) ticks on day‑13 post‑oviposition. 
Furthermore, we observed that eggs of dsHlemCHT ticks exhibited relatively early egg development 
and hatching, suggesting a dsRNA‑mediated activation of the HlemCHT gene in the eggs. This is the 
first attempt to provide evidence of RNAa in ticks. Although further studies are required to elucidate 
the detailed mechanism by which RNAa occurs in ticks, the outcome of this study provides new 
opportunities for the use of RNAa as a gene overexpression tool in future studies on tick biology, to 
reduce the global burden of ticks and tick‑borne diseases.
The manipulation of gene expression is instrumental to understanding disease mechanisms and developing novel 
 therapeutics1–8. In the last decade, this technology has focused mainly on gene silencing through the process of 
RNA interference (RNAi), based on the primary understanding of noncoding small RNAs (ncRNAs) such as 
double-stranded RNAs (dsRNAs) and short interfering RNAs (siRNAs)4,5,9, where gene function is assessed by 
the suppression of the gene expression. Recent reports have revealed a new role for ncRNAs in a phenomenon 
known as RNA activation (RNAa)2,10–13. In many genes, ncRNAs are reported to overlap the 3′- untranslated 
region (3ʹ-UTR) which plays a pivotal role in cellular regulation and disease p athology14. Although the role of 
terminus-associated ncRNAs is not understood, their association with the 3ʹ-UTR suggests that they may affect 
gene  regulation15. It was observed that ncRNAs targeting the 3ʹ-UTR of the PR gene caused either transcriptional 
gene silencing or activation by interacting with an overlapping noncoding sense  transcript16. In RNAa, dsRNAs, 
also referred to as small activating RNAs (saRNAs), target the promoter sequence and/or AU-rich elements in 
1Department of Parasitology and Tropical Medicine, Kitasato University School of Medicine, Sagamihara, 
Kanagawa  252-0374,  Japan.  2Department of Parasitology, Noguchi Memorial Institute for Medical Research, 
College of Health Sciences, University of Ghana, P.O. Box LG 581, Legon, Accra, Ghana. 3Department of Veterinary 
Paraclinical Sciences, College of Veterinary Medicine, University of the Philippines at Los Baños, College, 
4031  Laguna,  Philippines.  4Department  of  Parasitology,  Faculty  of Veterinary Science,  Bangladesh Agricultural 
University, Mymensingh 2202, Bangladesh. 5Department of Molecular and Cellular Parasitology, Graduate School 
of Medical Sciences, Kitasato University, Sagamihara, Kanagawa 252-0374, Japan. 6Department of Environmental 
Parasitology, Tokyo Medical and Dental University, Bunkyo-ku, Tokyo 113-8510, Japan. 7National Research Center 
for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Hokkaido 080-8555, 
Japan.  8Department  of  Tropical  Medicine  and  Center  for  Medical  Entomology,  The  Jikei  University  School  of 
Medicine,  Minato-ku,  Tokyo  105-8461,  Japan.  9Laboratory  of  Infectious  Diseases,  Joint  Faculty  of  Veterinary 
Medicine, Kagoshima University, Kagoshima 890-0065, Japan. 10Department of Veterinary Immunology, Graduate 
School of Veterinary Sciences, Osaka Metropolitan University,  Izumisano, Osaka 598-8531, Japan. 11Department 
of Molecular Protozoology, Research Institute for Microbial Diseases, Osaka University, Yamadaoka, Suita, 
Osaka  565-0871,  Japan.  12Center for Infectious Disease Education and Research (CIDER), Osaka University, 
Yamadaoka, Suita, Osaka  565-0871, Japan. *email: htakeshi@med.kitasato-u.ac.jp
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3ʹ-UTR of mRNA molecules and regulate gene expression at the transcriptional/epigenetic  level16. It provides 
a tool for the specific activation of endogenous and exogenous g enes17. Even though both RNAi and RNAa 
utilize ncRNAs, there are key differences. While siRNAs are loaded to argonaute (AGO) proteins 1–4 in  RNAi1, 
saRNAs specifically bind to AGO 2 protein, which is an indispensable requirement for dsRNA-mediated gene 
 activation1,11,18,19. In addition, unlike RNAi, RNAa occurs in the nucleus of cells, where transcriptional elements 
are recruited to upregulate gene  expression11. The recruitment of these elements results in the formation of 
RNA-induced transcriptional activation (RITA) c omplex18,19, which either binds directly to the DNA or binds 
to a newly formed transcript in the r egion20–22.
RNAa was first discovered, when saRNAs designed to have complementary sequences to the promoter regions 
of E-cadherin, p21, and vascular endothelial growth factor (VEGF), were observed to mediate sequence-specific 
mRNA transcription activation of the respective targeted genes in mammalian  cells11. Subsequently, the phenom-
enon has been applied to several other cells of mammals, plants, bacteria, and Caenorhabditis elegans10,13,23–27 
suggesting that this pathway is highly regulated and evolutionarily  conserved10. Recently, the presence of RNAa 
has been demonstrated and reported in Aedes aegypti, the arthropod vector of Dengue  fever17. Interestingly, 
there has not been any report of RNAa in any other arthropod vector, despite the ubiquitous presence of AGO 
2 in other  arthropods28, including t icks29,30.
Arthropod chitinases have been shown to be essential components in the hydrolysis of chitin, present in 
the exoskeleton and peritrophic membrane (PM) in the gut of arthropods, to allow degradation of the cuticle 
and development during the immature s tages31,32. In developmental events such as, embryogenesis, egg hatch-
ing, blood feeding, engorgement, and molting at various tick stages, chitinolytic enzymes such as chitinase are 
essential to remove old cuticle and allow synthesis of new cuticle for continued growth and development. Over 
the years, chitinases have been identified in different ticks species, including Haemaphysalis longicornis (Asian 
longhorned tick)31–34.
H. longicornis is a three-host tick, native to countries within eastern Asia, such as Korea, Japan and 
 China32,35–37. It is also a highly invasive tick species, well established as exotic species in Australia, New Zealand 
and recently, in the United States of A merica32,35–37. In addition, it is a zoonotic vector of Dabie bandavirus38, 
Anaplasma, Babesia, Borrelia, Ehrlichia and Rickettsia pathogen  species23,35–37. Their vectoral capacity is greatly 
influenced by their high reproductive potential in laying several thousand eggs, which hatch between 21 and 
25 days post-oviposition (dpo) at a temperature of 25 °C39. Chemical acaricides have been the primary control 
method. However, limitations in using acaricides have raised suggestions on applying gene expression manipu-
lation technologies such as RNAi, CRISPR-Cas, and RNAa in the control of t icks40–42. We recently identified a 
novel endochitinase-like (HlemCHT) gene tag, S03044-17L21 from the expressed sequence tag (EST) database 
derived from H. longicornis egg-specific complementary DNA (cDNA) l ibrary43. In trying to elucidate the func-
tion of this newly identified gene with RNAi, we coincidentally encountered the phenomenon of RNAa in the 
tick egg. In this study, we therefore report our findings and the potential use of RNAa in vector control. This is 
the first attempted evidence of RNAa in ticks.
Results
Sequence analysis of the full‑length HlemCHT. To obtain the full-length open reading frame (ORF) 
of the putative endochitinase-like gene (S03044-17L21 identified in the cDNA library, namely, HlemCHT), we 
employed the strategy summarised in Fig. 1a. Briefly, predicted exons and introns on the upstream sequence 
of the HlemCHT gene were retrieved from Chromosome 3 after a nucleotide BLAST search of a partial ORF 
sequence against both local (using BioEdit 7.2) and National Center for Biotechnology Information (NCBI) 
non-redundant (nr) protein databases of Haemaphysalis longicornis genome (GenBank accession number JAB-
STR000000000). Forward and reverse primers (Red arrows) were then designed to target sequences before the 
predicted starting methionine, and sequences at about 150 bp downstream from the end of the partial ORF 
fragment respectively. Amplified cDNA was sequenced after cloning into a vector and the assembly of newly 
sequenced 5ʹ-cDNA fragment and already existing cDNA fragment into a contig sequence was done using CAP3 
Sequence Assembly Program to obtain the full-length ORF. We finally obtained a cDNA of full length, 3768 bp 
(Fig. 1b), with an ORF of 1710 bp long. Analysis using the Compute pI/MW function of the Expasy web tool 
(www. expasy.o rg/r esou rces/c ompu te- pi- mw) revealed that the ORF encoded 569 amino acid polypeptides, with 
a predicted molecular weight of 64.7 kDa. The 5ʹ- and 3ʹ- UTRs of the cDNA consisted of 215 bp and 1843 bp 
respectively (Fig. 1b). The full-length ORF sequences obtained in this study have been deposited in the DDBJ, 
EMBL, and GenBank databases. (Accession number LC744416).
BLAST analysis using the nucleotide BLAST (BLASTn) program on the web tool (https:// blast.n cbi.n lm.n ih. 
gov/B last.c gi?P ROGRA M=b lastp &P AGE_T YPE=B lastS earch &L INK_L OC=b lasth ome) with HlemCHT (ORF) 
sequence as a query, showed homologs of HlemCHT in other tick species, such as Rhipicephalus sanguineus 
(Accession No. XP_049270163.1), Rhipicephalus microplus (Accession No. XP_037285628.1), Dermacentor 
andersoni (Accession No. XP_050044043.1), Dermacentor silvarum (Accession No. KAH7965576.1) and Ixodes 
scapularis (Accession No. XP_040067710.3) with sequence similarities of 88.10%, 89.35%, 87.64%, 88.66%, 
88.05%, and 91.60% respectively. HlemCHT was also observed to have a conserved motif, TTC GACG GC TTC 
GAC CTG GACT GGG AGTA. This motif was predicted by the web program Interpro scan (https://w ww. ebi.a c. 
uk/i nterp ro/r esult/I nterP roSca n/i prsca n5-R 20221 202-1 02605-0 383-5 28420 65-p 1m/) to code for an active site, 
characteristic of the protein family of glycoside hydrolases (Glycosyl hydrolases family 18 (GH18)) of which 
endochitinases belong.
Expression profile of HlemCHT transcripts in different life cycle and feeding stages. To deter-
mine the normal expression of HlemCHT gene within the life cycle of H. longicornis, we investigated its expres-
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Figure 1.  Full-length ORF of HlemCHT cDNA. (a) Strategy for obtaining full-length ORF of HlemCHT. 
(1) Predicted exons and introns on the upstream sequence of the putative endochitinase-like gene (S03044-
17L21 identified in the cDNA library, namely, HlemCHT) were retrieved from Chromosome 3 (Chr #3) after a 
nucleotide BLAST search of a partial ORF sequence against both local and NCBI nr databases of Haemaphysalis 
longicornis genome. (2) Forward and reverse primers (Red arrows) were designed to target sequences before 
the predicted starting methionine, and sequences at about 150 bp downstream from the end of the partial ORF 
fragment respectively. (3) Amplified cDNA was sequenced after cloning into a vector. (4) Assembly of newly 
sequenced 5ʹ-cDNA fragment and already existing cDNA fragment into a contig sequence was done using 
CAP3 Sequence Assembly Program to obtain the full-length ORF. (b) HlemCHT gene structure showing the 
positions the RNAi target site (Green) within the full-length ORF (Orange) and the RNAa target site (Yellow) at 
the 3′-UTR. Also, within the ORF is a conserved sequence (Blue) in other ixodid tick species’ endochitinase-like 
genes.
sion in the different life stages (12/15 days embryonated eggs, larvae. nymphs and adults) of H. longicornis par-
thenogenetic ticks at different feeding stages (unfed, partially fed and fully engorged) using reverse-transcription 
PCR (RT-PCR). The statutory expressed H. longicornis 40S Ribosomal S3a protein was used as an internal con-
trol gene, while a no template control served as a negative control. In our results, we observed HlemCHT to be 
solely present in the 12 and 15 dpo eggs (Fig. 2a), suggesting that the expression of HlemCHT is embryo-specific. 
In addition, we investigated the expression profile of HlemCHT in tick eggs during embryogenesis. To do this, 
eggs from fully-engorged maternal ticks were divided into eight (1-, 4-, 7-, 10-, 13-, 16-, 19- and 21-days post-
infection) groups, according to which, they were harvested and frozen. Total RNA was then extracted from each 
group of tick eggs (50–100 /group) and quantitative reverse-transcription PCR (RT-qPCR) analysis was per-
formed. H. longicornis 40S ribosomal gene S3a was used to as a reference gene to normalize the HlemCHT gene 
expression data. From our results, the relative expression of HlemCHT increased significantly (P < 0.005) with 
embryo development until day 13 dpo, which recorded the highest expression (Fig. 2b). Relative expression val-
ues per the days assessed are as follows: day 1 = no expression determined, day 4 = 1.00 ± 0.00, day 7 = 2.27 ± 0.04, 
day 10 = 6.25 ± 0.27 and day 13 = 7.95 ± 0.16. Beyond 13 dpo, the relative expression sharply declined as follows: 
day 16 = 0.24 ± 0.01, day 19 = 0.19 ± 0.02 and day 21 = 0.46 ± 0.03, suggesting that HlemCHT is highly expressed 
in H. longicornis eggs during mid-phase (day 12–15 dpo) embryogenesis.
Induction of RNAa in H. longicornis using dsRNA. To investigate the potential occurrence of the 
RNAa phenomenon in eggs of H. longicornis, we targeted the 3ʹ-UTR of the HlemCHT gene because, among 
the many observations made concerning the genomic target site of saRNA, some have suggested that saRNAs 
demonstrate optimal function when they target sequences downstream from the 3′-UTR of the intended tar-
get  gene16. Therefore, adult female ticks were injected with synthesized dsRNAs targeting either the 3ʹ-UTR of 
HlemCHT gene (dsHlemCHT) or a non-target control, Escherichia coli malE (Ec-malE) gene (dsEc-malE). RNA 
was then extracted from day 13 eggs derived from the dsRNA-injected maternal ticks because HlemCHT was 
observed to be highly expressed in tick eggs at 13 dpo (Fig. 2b), and as such, any changes in expression pattern, 
be it activation or suppression, would be clearly shown. The expression of HlemCHT was quantified by RT-
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Figure 2.  Normal expression of HlemCHT in H. longicornis. (a) RT-PCR analysis of HlemCHT in different 
developmental stages of H. longicornis. Analysis was performed using total RNA from eggs (12 and 15 Dpo), 
the whole body of larvae (unfed, partially fed, engorged), nymph (unfed, partially fed, engorged), and adult 
(unfed, partially fed, engorged). H. longicornis 40S ribosomal protein S3a was used as an internal control. PCR 
products were analysed on 1.5% agarose gel and stained with ethidium bromide for viewing. Original gels are 
presented in Supplementary Fig. S1. (b) RT-qPCR analysis of HlemCHT during embryogenesis of tick eggs. 
The expression of each mRNA was normalized to the expression of 40S ribosomal protein S3a measured in 
same RNA preparation. The relative expression values are represented as “means ± SD”. Error bars represent 
the SEM of 4 replicates. *Indicates a difference at P < 0.05; **designates P < 0.005; ***designates P < 0.001. The 
results shown are from a single experiment and are representative of three independent experiments. n.d. not 
determined.
qPCR analysis. In our results, targeting the 3ʹ-UTR, showed a 19 ± 8.20—fold significant (P = 0.0145) increase 
in HlemCHT expression in dsHlemCHT-derived eggs relative to dsEc-malE-derived eggs (Fig. 3a). In addition, 
morphological examination of the dsHlemCHT-derived eggs at 13 dpo, showed a normal phenotype character-
istic of stage 12 embryo described in previous s tudies44,45 with evidence of limb formation, whitish strands of 
malpighian tubules and a distinctly formed bright white hindgut, similar to dsEc-malE-derived eggs (Fig. 3b). 
Moreover, dsHlemCHT- derived eggs were observed to begin hatching within 20 days post-oviposition, about 
24 h earlier than dsEc-malE -derived eggs (Fig. 3c). Furthermore, at 21 days post-oviposition, when dsEc-malE-
derived eggs began hatching, the hatching rate was 32.9 ± 32.6%. In contrast, the hatching rate for dsHlemCHT-
derived eggs was relatively higher at 52.95 ± 13.50%, although not significant (P = 0.5066) (Fig. 3c,d). On the 
other hand, when the ORF of the same gene was targeted, the relative expression of HlemCHT was observed 
to be 0.09 ± 0.02 in dsHlemCHT-derived eggs as opposed to 1.00 ± 0.04 in dsEc-malE-derived eggs, suggesting 
a significant (P = 0.0001) downregulation of the gene in dsHlemCHT-derived eggs at 13 dpo (Fig. 4a). Interest-
ingly, phenotypic observation of day 13 dsHlemCHT-derived eggs also revealed a normal phenotype as observed 
when the 3ʹ-UTR was targeted, with no difference observed between the dsEc-malE-derived and dsHlemCHT-
derived eggs (Fig. 4b). Consequently, hatching of dsRNA-derived eggs begun on day 22 post-oviposition, with no 
significant (P = 0.5306) difference observed between hatching rates of dsHlemCHT-derived eggs (27.73 ± 4.00%) 
and dsEc-malE-derived eggs (32.26 ± 5.51%) (Fig. 4c,d). These results, therefore, indicate the importance of the 
3ʹ-UTR as an essential target for dsRNA-mediated activation of the HlemCHT gene in tick eggs.
Discussion
RNAa is the targeted upregulation of a specific g ene21. The use of RNAa was limited to only a few major mam-
malian  models3,11,13,46, but it has now been reported in other organisms such as plants, bacteria, C. elegans, and 
recently, mosquito arthropods. It is currently being explored as a potential tool for the specific activation of genes, 
towards the development of novel therapeutics for undruggable disease and vector c ontrol14,17.
In the present study, we targeted the 3ʹ-UTR of a novel endochitinase-like gene (HlemCHT), in H. longicornis 
eggs for dsRNA-mediated gene activation and showed an increased expression of the gene on day 13 post-
oviposition. In a simultaneous experiment, when we targeted sequences within the ORF of the same gene, we 
observed a downregulation of the gene (Fig. 4), suggesting the possibility of a dsRNA-mediated gene activation, 
which may be based on the association of the dsRNA (saRNA) with the 3ʹ-UTR of the gene.
Different dsRNA-mediated gene activation mechanisms have been proposed to involve the 3ʹ-UTR. For 
instance, it has been suggested that the saRNAs that target the 3ʹ-UTR trigger a looping mechanism bringing 
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Figure 3.  Confirmation of HlemCHT activation by dsRNA targeting the 3ʹ-UTR. (a) Relative gene expression 
of HlemCHT at 13 days post-oviposition (dpo). Data is represented by a histogram. Relative expression of the H. 
longicornis 40S ribosomal gene was used to normalize the HlemCHT gene expression data. Error bars represent 
the SEM of 4 replicates. *P = 0.0145. (b) Representative stereomicroscopic images of dsRNA-derived eggs at 
13 dpo. (c) Representative stereomicroscopic images of dsRNA-derived eggs hatching at 20 and 21 dpo. (d) 
Histogram showing the comparison of the hatching rate of dsHlemCHT-derived eggs, as against eggs of dsEc-
malE at 21 dpo. Error bars represent the SEM of 4 replicates. The results shown are from a single experiment and 
are representative of three independent experiments.
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Figure 4.  Confirmation of HlemCHT suppression by dsRNA targeting the ORF. (a) Relative gene expression 
of HlemCHT at 13 days post-oviposition (dpo). Data is represented by a histogram. Relative expression of 
the H. longicornis 40S ribosomal gene S3a was used to normalize the HlemCHT gene expression data. Error 
bars represent the SEM of 4 replicates. *P = 0.0001. (b) Representative stereomicroscopic images of dsRNA-
injected tick eggs at 13 dpo. (c) Histogram showing the comparison of the hatching rate of dsHlemCHT-derived 
eggs, as against eggs of dsEc-malE at 22 dpo. Error bars represent the SEM of 4 replicates (d) Representative 
stereomicroscopic images of dsRNA-derived eggs hatching at 22 dpo. The results shown are from a single 
experiment and are representative of three independent experiments.
the 3′-terminus and promoter into proximity to allow the recruitment of additional proteins and modulation of 
promoter a ctivity16. It has also been proposed that saRNA molecules may target AU-rich elements in 3ʹ-UTR 
of mRNA molecules and induce gene expression at the transcriptional level. For example, two studies showed 
miR-369-3 to positively regulate mRNA translation by targeting AU-rich elements in 3′-UTRs in stressed  cells47. 
However, in this study, characteristic AU-rich elements were not found in the 3′-UTR of HlemCHT, which sug-
gests that the gene activation observed in our study may probably be due to a looping mechanism triggered by 
saRNAs targeting the 3′-UTR, and bringing the 3′ terminus and promoter into proximity for the activation of 
the gene, leading to a 19-fold increase in gene expression. Further studies are however required to elucidate the 
exact mechanism of RNAa in ticks.
This study also established a positive relationship between dsRNA-mediated gene expression and egg hatching, 
concerning hatching start time and percentage of hatched eggs. We observed that eggs of dsHlemCHT began 
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hatching about 24 h earlier than control eggs, as well as a slightly higher percentage of hatched eggs by 21 dpo, 
suggesting that the possible dsRNA-mediated gene activation of HlemCHT observed on day 13, drove the early 
hatch of dsHlemCHT -derived eggs. Indeed, several reports on the role of chitinase in eggs of arthropods and 
nematodes, have indicated that chitinase (-like) genes are vital for successful egg development and  hatching48–53. 
For example, knockdown of Tribolium (flour beetle) chitinase-like gene, TcCHT10 in adult females, resulted in 
none of the eggs hatching, despite the presence of fully developed embryos in these  eggs52. In our study, a single 
injection of dsRNA targeting the ORF, into unfed adult female ticks resulted in no phenotypic changes, despite 
the downregulation of the HlemCHT gene observed (Fig. 4). However, a repeated introduction (once daily for 
3 consecutive days) of dsRNA into fully-engorged adult female ticks before the start of oviposition resulted in 
a downregulation of the gene and the subsequent delay or reduction of hatching rate of H. longicornis eggs. The 
results for this modified RNAi method will be reported in another article being prepared currently.
Overall, this study’s results provide some evidence of the presence of RNAa phenomenon in the tick vector, 
H. longicornis for the first time. More studies are however required to provide further insight into the detailed 
mechanisms by which the expression of genes is activated by dsRNA (saRNAs) in ticks. Currently, no estab-
lished gene manipulation technology permits stable gene knock-ins for applications such as gene replacement 
and over-expression in t icks41. Therefore, the discovery of RNAa phenomenon in ticks has the potential to 
provide new technology for the application of RNAa in the over-expression of both endogenous and exogenous 
genes in ticks. For this purpose, we will investigate other H. longicornis genes for the established phenomenon 
of RNAa in subsequent studies. Furthermore, reports of RNAa in Ae. aegypti17 and H. longicornis (this study), 
show the potential of  saRNAs40, considering their small size, versatility, and safety, to reverse phenomena, such 
as the emergence of acaricide/insecticide resistance due to the downregulation of critical g enes54,55. Finally, this 
report will hopefully provide new opportunities, based on the usefulness of RNAa as a tool for future research 
in arthropod biology, to modulate gene responses, to restrict the expansion of tick populations and transmission 
of pathogens to reduce the global burden of ticks and tick-borne diseases.
Materials and methods
Ticks and animals. The parthenogenetic Okayama strain of the ixodid tick H. longicornis bred by feed-
ing on mice as described  previously56, was maintained at the Department of Parasitic and Tropical Medicine, 
Kitasato University School of Medicine (KUSM), Sagamihara, Kanagawa, Japan. Its ability to reproduce par-
thenogenetically by feeding on the blood of several hosts including mice makes it easier to maintain in the lab. 
Therefore has become an ideal model tick species for tick experiments in many labs globally as can be seen in 
various p ublications27,37,57–66. Fifteen-week-old BALB/c CrSlc mice (SLC Japan, Shizuoka, Japan) were adapted 
to standard animal husbandry conditions (25 °C, 60% Relative Humidity (RH)) for 7 days, and subsequently to 
tick maintenance condition (25 °C, 70% RH) for 2 days before the experiment.
Ethical statement. Mice maintenance and tick feeding experiments were carried out under protocols 
approved by the Animal Care and Use Committee, KUSM (Approval no. 2022.073), which are based on the 
Law for the Humane Treatment and Management of Animals, Standards Relating to the Care and Management 
of Laboratory Animals Relief of Pain (the Ministry of the Environment), Fundamental Guidelines for Proper 
Conduct of Animal Experiment Related Activities in Academic Research Institutions (the Ministry of Educa-
tion, Culture, Sports, Science Technology), as well as, the Guidelines for Proper Conduct of Animal Experiments 
(the Science Council of Japan), and are reported in accordance with the ARRIVE g uidelines67,68. At the end of 
the experiments, the animals were euthanized by exsanguination under anesthesia with isoflurane followed by 
cervical dislocation.
Cloning and sequencing of full‑length HlemCHT. The plasmid (pCR-Blunt II-TOPO/S03044-17L21) 
containing partial cDNA encoding the HlemCHT transcript was first isolated from a previously established 
cDNA  library43. A BLASTn search of the partial sequence of the 5ʹ-terminal (ATG ACG TAT GATC TG CGT 
GGG AAC TGG GCT GGC TTT ACGG) of this cDNA was performed locally against Haemaphysalis longicornis 
genome database (GenBank accession number JABSTR000000000) with the p-value of <  10–6 by using BioEdit 
7.2 (https:// bioed it. softwa re. infor mer.c om/7. 2/). According to this analysis, the partial sequence was identified 
in Chromosome No.3 with an expected value of 1.00E−12 and an identity of 97%. By using this partial genome 
sequence, the BLASTN program on the web tool (https:// blast. ncbi.n lm. nih.g ov/B last. cgi? PAGE_ TYPE= Blast 
Search) was done again against the NCBI nr database and the predicted HlemCHT gene (GenBank accession 
number currently not available due to ongoing updates of genome annotations) located on the chromosome 
No. 3 of H. longicornis was identified. According to the previous (11th February 2022) annotated information, 
the upstream sequence including predicted exons and introns was retrieved to design the forward primer on 
the most upstream region before starting methionine. Reverse primer was also designed at about 150 bp down-
stream of the edge of known cDNA fragment. Both forward and reverse primers on the 5ʹ-UTR were designed 
to amplify the unknown fragment flanked by the starting methionine and the partial cDNA sequence. Primers 
used in this study were purchased from Eurofins Genomics K.K., Tokyo, Japan, and are listed in Table 1. RT-PCR 
was then performed to amplify the unknown target sequence. The amplification was performed using a PCR 
program of 94 °C for 2 min, 35 cycles of 98 °C for 30 s, either 64–67 °C for 30 s, and 68 °C for 30 s, followed by 
elongation for 1 min at 68 °C. The PCR products were purified from agarose gel using VIOGENE Gel/PCR DNA 
isolation system Extraction Kit (VIOGENE BIOTEK, TAIWAN) according to the manufacturer’s instructions. 
Cloning of the target sequence into pCR-Blunt II-TOPO vector was also done according to the manufacturer’s 
instruction and transformed into DH5α E. coli competent cells (Nippon gene DO., LTD., Tokyo, Japan). Posi-
tive colonies were sub-cultured and plasmid purification was done using PureLink™ Quick Plasmid Miniprep 
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Primers Position Primer sequence Tm ( °C)
Forward TGTA TGC GTT CAG TG CTA CCG GGCG TA 65.1
HlemCHT 5’-complete ORF
Reverse TAAA GC CAG CCCA GT TCC CAC GCAG AT 65.1
Forward TGG TTG GGC TGA GGG CGGA AA GAAG TA 65.1
HlemCHT 5 ’-TOPO colony PCR
Reverse TGT AGA CGAA AC TTG GCG ACGG GC ACA 65.1
Forward TGG TTG GGC TGAG GGC GGA AAG AA GTA 65.1
HlemCHT 5’-complete ORF sequencing
Reverse TGT AGA CGAA AC TTG GCGA CG GGCA CA 65.1
Forward ACG AGCT GT GCG TGC TTG CCA AAG TCT 65.1
Full HlemCHT ORF
Reverse TGC AGG CTA GTT TTT CCG GCT AGC CACTC 66.2
Forward CGG GGA GGT GCA TAC TCTG AC AGG GCTAA 67.6
Full HlemCHT ORF sequencing
Reverse CGG TGTT GC CCT TGT CACT CA GCGT GT 66.6
Forward *gaaattaatacgactcactataggTCA CAT GCT CAC GATC ACC CG CATA CC 65.1
HlemCHT (3ʹ-UTR) dsRNA synthesis (T7)
Reverse *gaaattaatacgactcactataggGGA CGTG TC CTT GCA CGT TGGC AT ACA 65.1
Forward TCAC ATG CTC AC GAT CAC CCG CATA CC 65.1
HlemCHT (3ʹ-UTR) dsRNA synthesis (Gene-specific)
Reverse GGA CGTG TCC TTG CAC GTT GGC ATA CA 65.1
Forward *gaaattaatacgactcactataggCAA GGGC AA CACC GGA CT CAGG GC 66.6
HlemCHT (ORF) dsRNA synthesis (T7) TTA 
Reverse *gaaattaatacgactcactataggGAAC AT CACC AT GGG CGT CCC GTA CAC 66.6
Forward CAAG GG CAAC ACC GGA CT CAG GGC TTA 66.6
HlemCHT (ORF) dsRNA synthesis (Gene-specific)
Reverse GAA CAT CAC CATG GG CGT CCC GTA CAC 66.6
Forward CGA CCG CTT TGG TGG CTA CGC TCAA TC 66.6
MalE gene
Reverse CCG CAGT ACG CAC GGC AT ACCA GA AAG 66.6
Forward *gaaattaatacgactcactataggGTTG GCT GA AAT CAC CCC GGA CAA 65.1
MalE dsRNA synthesis (T7) AGC 
Reverse *gaaattaatacgactcactataggTTAA TA CCT GCGC TCA GC ACG CCAA CG 65.1
Forward GTT GGC TGA AATC AC CCCG GAC AAA GC 65.1
MalE dsRNA synthesis (Gene-specific)
Reverse TTA ATAC CT GCG CTCA GCA CGC CA ACG 65.1
Forward CATA CC GCAC GA GAAC GA CTG CAC CAA 65.1
Full HlemCHT ORF-TOPO colony PCR / HlemCHT RT-qPCR
Reverse CAC AGTT GCC GT TGTC CTG GT TCCA CA 65.1
Forward CCTC AT CCG CAA GGT GAA GGT GCT CAA 65.1
40S ribosomal S3a RT-qPCR
Reverse AAG CGCC CT CAGA CA TTC TCCA GCA CA 65.1
Table 1.  List of primers. *gaaattaatacgactcactatagg—T7 promoter sequences.
Kit (Invitrogen by ThermoFisher Scientific K.K., Tokyo, Japan), after which samples were sequenced by Sanger 
technology in an independent external company (Eurofins Genomics K.K., Tokyo, Japan). Starting methionine 
and 5’UTR sequence were confirmed accordingly. The newly sequenced 5ʹ-cDNA fragment and already exist-
ing cDNA fragment were assembled into a contig sequence using CAP3 Sequence Assembly Program on the 
web tool (https://d oua. prabi. fr/s oftw are/ cap3)69. Full-length ORF predicted to encode this novel endochitinase-
like protein was isolated and confirmed from another set of RT-PCR with the Full HlemCHT ORF primer set 
(Table 1) and sequence analysis at the company (Eurofins Genomics K.K., Tokyo, Japan). The strategy for cloning 
and sequencing of full-length HlemCHT is illustrated in Fig. 1a.
Expression pattern of HlemCHT in different life cycle and feeding stages. To determine the nor-
mal expression of HlemCHT in different developmental and feeding stages of H. longicornis, reverse transcrip-
tion-polymerase chain reaction (RT-PCR) was employed. To do that, larval (150), nymphal(100) and adult (7) 
stages of H. longicornis ticks were allowed to feed on the shaven back of BALB/c mice (3 mice per each devel-
opmental stage). At the beginning of the tick expansion period (3–5 days after tick attachment), About 50 par-
tially engorged larvae, 50 partially engorged nymphs and 5 partially engorged adult female ticks, were detached 
carefully with fine tip forceps without breaking the hypostome (in the case of the adults) or gently passing a 
soft-bristled paint brush over the attached ticks to detach them (in the case of larvae and nymphs). The rest of 
the ticks were allowed to fully engorge and drop off by themselves. Two fully engorged adult ticks were allowed 
to lay eggs under the suitable temperature of 25 °C and > 70% humidity. Eggs laid by each engorged tick were 
further divided into two groups (50–100/group) and incubated for either 12 or 15 days post oviposition. Either 
tick developmental stages (unfed, partially fed and fully engorged (after egg laying)) or egg samples collected at 
their respective times were immediately flash frozen in liquid nitrogen and stored at − 80 °C until used. For RNA 
extraction, frozen samples were ground to a fine powder, using a mortar and pestle kept in liquid nitrogen. Total 
RNA was isolated from the resulting powder with the RNeasy mini kit (Qiagen, Valencia, CA, USA) according 
to the manufacturer’s instructions. The quantity and integrity of the RNA was measured spectrophotometri-
cally by ultraviolet light absorbance using NanoDrop™ One Microvolume UV–Vis Spectrophotometer (Thermo 
Scientific, Rockford, IL, USA) and stored at − 80 °C until use. Complementary DNA (cDNA) was synthesized 
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from about 150 ng total RNA extracted from the tick/egg samples using ReverTrace™ qPCR RT Master Mix 
with gDNA remover (Toyobo, Japan) kit according to manufacturer’s instructions. Polymerace chain reactions 
(PCRs) were performed to amplify a 110 bp region of HlemCHT and a 192 bp region of 40S Ribosomal protein 
S3a using primers purchased from Eurofins Genomics K.K., Tokyo, Japan. Primers are shown in Table 1. The 
reaction mixtures for PCR were prepared using the KOD Plus Neo ™ PCR kit (Toyobo, Japan) and band amplifi-
cation was done using a Bio-Rad T100™ Thermal Cycler, with the following cycling conditions; initial denatura-
tion at 94 °C for 2 min, followed by 35 cycles of denaturation at 98 °C for 10 secs, annealing at 66 °C for 30 secs 
(for both genes) and extension at 68 °C for 10 secs, with a final extension performed at 68 °C for 1 min. The PCR 
products were separated by a 1.5% agarose gel electrophoresis and stained with ethidium bromide for viewing.
Expression pattern of HlemCHT in tick eggs during embryo development. By the same method 
of blood feeding and egg laying described above, eggs were collected from two different fully engorged maternal 
ticks, and each set was divided into eight (1-, 4-, 7-, 10-, 13-, 16-, 19- and 21-days post-infection) groups, accord-
ing to which, eggs were harvested and frozen. Total RNA from tick eggs (about 50–100/group) was extracted, 
followed by subsequent cDNA synthesis as have already been described in this study. RT-qPCR analysis was 
then performed using TB Green® Premix DimerEraser™ kit (Takara, Japan) according to the manufacturer’s 
instructions with the same primer sets (Table 1) already described for either HlemCHT or H. longicornis 40 s 
ribosomal protein S3a to determine their expression levels. Each experiment was done with two different RNA 
samples per each group and were run in duplicates. The experiment was repeated three times, and the data were 
analyzed with the 2 −ΔΔCt  method70 using Microsoft Excel version 2019 and expressed as mean ± standard devia-
tion. Statistical significance was determined using an unpaired two-tailed student’s t-test, with a value of P < 0.05 
considered significant after the normal distribution and homogeneity of variance were confirmed by Kolmogo-
rov–Smirnov and F-tests respectively.
Double‑stranded RNA (dsRNA) synthesis. To investigate the occurrence of RNAa in ticks, the 3ʹ-UTR 
of the HlemCHT gene was targeted. A template partial-length cDNA fragment at the 3ʹ-UTR of the HlemCHT 
transcript (497 bp) was therefore amplified from a pCR-Blunt II-TOPO/S03044-17L21 plasmid by PCR, using 
primer sets flanked by a T7 promoter and gene-specific primer sets (Table 1) which were designed using Primer 
3 (v. 0.4.0) program (https:// bioin fo. ut.e e/ primer 3-0. 4.0/) and purchased from Eurofins Genomics K.K., Tokyo, 
Japan. To provide further evidence that the association of dsRNA to the 3ʹ-UTR is vital for the occurrence of 
RNAa, we also synthesized dsRNA targeting the ORF in a parallel experiment. A template partial-length cDNA 
fragment within the ORF of the HlemCHT transcript (586 bp) was therefore amplified from the same plasmid 
using T7 promoter-flanked primers, as well as gene-specific primer sets designed and purchased as already 
described (Table 1). As a negative control (also a non-target control), dsRNA targeting Escherichia coli malE (Ec-
malE) was also synthesized. Ec-malE is absent in ticks and dsRNA targeting Ec-malE has been shown in several 
s tudies61–63,71 not to affect normal tick physiology, including egg laying and the normal expression of tick-specific 
genes. Moreover, it has been shown to give comparable results to a no injection (non-manipulated) control in 
most of the studies referred to above. Briefly, the Ec-malE DNA fragment (848 bp) was first amplified from E. 
coli using primers (Table 1) designed and purchased as already described. The amplified fragments were then 
cloned into the pCR-Blunt II-TOPO vector as earlier described. Positive colonies were sub-cultured and plasmid 
purification was also done as already described. A template DNA fragment of the Ec-malE gene (577 bp) was 
then amplified from the pCR-Blunt II-TOPO/malE plasmid by PCR, using T7 promoter-flanked primers, as 
well as gene-specific primer sets designed and purchased as already described (Table 1). The amplification was 
performed using a PCR program of 94 °C for 2 min, 35 cycles of 98 °C for 30 s, either 66 °C (HlemCHT/3ʹ-UTR), 
66 °C (HlemCHT/ORF) or 67 °C (Ec-malE) for 30 s, and 68 °C for 30 s, followed by elongation for 1 min at 68 °C. 
The PCR products were purified from agarose gel using VIOGENE Gel/PCR DNA isolation system Extrac-
tion Kit (VIOGENE BIOTEK, TAIWAN) according to the manufacturer’s instructions. DsRNAs of HlemCHT 
(3ʹ-UTR), HlemCHT (ORF) and Ec-malE were synthesized by in vitro transcription using the T7 RiboMAX™ 
Express RNAi System (Promega, Madison, WI, USA) according to the manufacturer’s instructions. The purity of 
synthesized dsRNAs was checked by running 1 µL of diluted dsRNA (1:10, 1:100, and 1:1000 with Nuclease-Free 
water) in a 1.5% agarose gel electrophoresis. Concentrations of dsRNAs were also measured by ultraviolet light 
absorbance using NanoDrop™ One Microvolume UV–Vis Spectrophotometer (Thermo Scientific, Rockford, IL, 
USA).
Microinjection of dsRNA into adult female ticks. A volume of 0.5 μl of 2 μg/μl either HlemCHT (3ʹ-
UTR), HlemCHT (ORF) or Ec-malE dsRNA was injected into the hemocoel from the fourth left coxa of unfed 
adult female H. longicornis ticks (10 ticks per each treatment) with a glass needle generated by a micropipette 
PC-10 puller (Narishige International, NY, USA) using an IM-11-2 pneumatic microinjector (Narishige Inter-
national, NY, USA). Injected ticks were left for 18 h at 25 °C in a moist chamber. After 18 h, the dsRNA-injected 
ticks (dsHlemCHT (3ʹ-UTR), dsHlemCHT (ORF) and dsEc-malE) were observed for any mortality due to trau-
matic injury resulting from injection. Live ticks within each treatment group were then placed on a mouse to 
blood feed as earlier described. After 3–4 days of tick attachment, partially fed ticks were removed, leaving 2 ticks 
on each mouse for optimal blood feeding and engorgement. Each engorged tick was allowed to lay eggs after 
detachment from the host mouse.
Maintenance and monitoring of dsRNA‑injected ticks’ eggs. Eggs laid daily by each of the engorged 
ticks were kept separately in the wells of 24-well plates under humidified conditions at 25 °C. Some of the egg 
samples were allowed to develop and hatch, while others were collected on day 13 post-oviposition for con-
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firmation of either the activation or suppression of gene expression by dsRNAs. To monitor the embryonic 
development of eggs till larval hatching, 1% agarose in distilled water, with 2.5 µM amphotericin B (AmpB) was 
dispensed into 3.5 cm petri dishes and allowed to solidify. Eggs were placed on solidified gel and covered slightly 
with halocarbon oil. Petri dishes containing eggs were kept under humidified conditions at 25 °C. Tick eggs 
were monitored daily for any morphological changes under a stereomicroscope (Lecia microsystems, Wetzlar, 
Germany). The hatching rate was calculated as follows: (Number of hatched eggs / total number of eggs moni-
tored) × 100%. The data were analyzed using Microsoft Excel version 2019 and expressed as mean ± standard 
deviation. Statistical significance was determined using an unpaired two-tailed student’s t-test, with a value of 
P < 0.05 considered significant after the normal distribution and homogeneity of variance were confirmed by 
Kolmogorov–Smirnov and F-tests respectively.
Confirmation of dsRNA‑induced gene activation (or silencing) in tick eggs.. To confirm the acti-
vation (or silencing) of HlemCHT by dsRNAs, 50–100 eggs from each of the two sets of samples per dsRNA-
treatment group, were harvested and frozen on day 13 post-oviposition, after which RNA extraction, cDNA syn-
thesis and RT-qPCR analysis were performed as previously described in this study. Each experiment was done 
with two different RNA samples per each group, which were run in duplicates. The experiments were repeated 
three times, and data were analyzed as previously described in this study.
Data availability
The datasets used and/or analyzed during the current study are available from the corresponding author on 
reasonable request. Please note that the details regarding the EST database and cDNA library described in this 
experiment can be referred from Umemiya-Shirafuji et al.43.
Received: 22 February 2023; Accepted: 5 June 2023
References
 1. Zhao, X. Y., Voutila, J., Habib, N. A. & Reebye, V. RNA Activation. Innov. Med. 241–249. https://d oi.o rg/1 0.1 007/9 78-4-4 31-5 5651-
0_ 20 (2015).
 2. Fröhlich, K. S. & Vogel, J. Activation of gene expression by small RNA. Curr. Opin. Microbiol. 12, 674–682 (2009).
 3. Jiao, A. L. & Slack, F. J. RNA-mediated gene activation. Epigenetics 9, 27–36 (2014).
 4. Vaschetto, L. M. RNA activation: A diamond in the rough for genome engineers. J. Cell. Biochem. 119, 247–249 (2018).
 5. Zotti, M. et al. RNA interference technology in crop protection against arthropod pests, pathogens and nematodes. Pest. Manag. 
Sci. 74, 1239–1250 (2018).
 6. Hatta, T. et al. RNA interference of cytosolic leucine aminopeptidase reduces fecundity in the hard tick. Haemaphysalis longicornis. 
Parasitol. Res. 100, 847–854 (2007).
 7. Liao, M. et al. Molecular characterization of Rhipicephalus (Boophilus) microplus Bm86 homologue from Haemaphysalis longicornis 
ticks. Vet. Parasitol. 146, 148–157 (2007).
 8. Galay, R. L. et al. Induction of gene silencing in Haemaphysalis longicornis ticks through immersion in double-stranded RNA. 
Ticks Tick. Borne. Dis. 7, 813–816 (2016).
 9. Kim, D. H. & Rossi, J. J. RNAi mechanisms and applications. Biotechniques 44, 613–616 (2008).
 10. Huang, V. et al. RNAa is conserved in mammalian cells. PLoS ONE 5, 1–8 (2010).
 11. Li, L. C. et al. Small dsRNAs induce transcriptional activation in human cells. Proc. Natl. Acad. Sci. U. S. A. 103, 17337–17342 
(2006).
 12. Place, R. F., Li, L. C., Pookot, D., Noonan, E. J. & Dahiya, R. MicroRNA-373 induces expression of genes with complementary 
promoter sequences. Proc. Natl. Acad. Sci. U. S. A. 105, 1608–1613 (2008).
 13. Janowski, B. A. et al. Activating gene expression in mammalian cells with promoter-targeted duplex RNAs. Nat. Chem. Biol. 3, 
166–173 (2007).
 14. Kwok, A., Raulf, N. & Habib, N. Developing small activating RNA as a therapeutic: Current challenges and promises. Ther. Deliv. 
10, 151–164 (2019).
 15. Ni, W. J., Xie, F. & Leng, X. M. Terminus-associated non-coding RNAs: Trash or treasure?. Front. Genet. 11, 1–10 (2020).
 16. Yue, X. et al. Regulation of transcription by small RNAs complementary to sequences downstream from the 3’ termini of genes. 
Nat. Chem. Biol. 6, 621–629 (2010).
 17. De Hayr, L., Asad, S., Hussain, M. & Asgari, S. RNA activation in insects: The targeted activation of endogenous and exogenous 
genes. Insect Biochem. Mol. Biol. 119, 103325 (2020).
 18. Portnoy, V. et al. SaRNA-guided Ago2 targets the RITA complex to promoters to stimulate transcription. Cell Res. 26, 320–335 
(2016).
 19. Portnoy, V., Huang, V., Place, R. F. & Li, L.-C. Small RNA and transcriptional upregulation. Wiley Interdiscip. Rev. RNA. 2, 748–760 
(2011).
 20. Matsui, M. et al. Activation of LDL Receptor (LDLR) expression by small RNAs complementary to a noncoding transcript that 
overlaps the LDLR promoter. Chem Biol. 17, 1344–1355 (2010).
 21. Meng, X. et al. Small activating RNA binds to the genomic target site in a seed-region-dependent manner. Nucleic Acids Res. 44, 
2274–2282 (2016).
 22. Schwartz, J. C. et al. Antisense transcripts are targets for activating small RNAs. Nat. Struct. Mol. Biol. 15, 842–848 (2008).
 23. Turner, M. J., Jiao, A. L. & Slack, F. J. Autoregulation of lin-4 microRNA transcription by RNA activation (RNAa) in C. elegans. 
Cell Cycle 13, 772–781 (2014).
 24. Gujrati, H., Ha, S., Mohamed, A. & Wang, B. D. MicroRNA-mRNA regulatory network mediates activation of mTOR and VEGF 
signaling in African American prostate cancer. Int. J. Mol. Sci. 23, 2926 (2022).
 25. Park, K. H. et al. Targeted induction of endogenous VDUP1 by small activating RNA inhibits the growth of lung cancer cells. Int. 
J. Mol. Sci. 23, 7743 (2022).
 26. Tan, H. et al. Pan-cancer analysis on microRNA-associated gene activation. EBioMedicine 43, 82–97 (2019).
 27. Yang, K., Shen, J., Tan, F. Q., Zheng, X. Y. & Xie, L. P. Antitumor activity of small activating rnas induced pawr gene activation in 
human bladder cancer cells. Int. J. Med. Sci. 18, 3039–3049 (2021).
 28. Van Rij, R. P. et al. The RNA silencing endonuclease Argonaute 2 mediates specific antiviral immunity in Drosophila melanogaster. 
Genes Dev. 20, 2985–2995 (2006).
Scientific Reports |         (2023) 13:9341  | https://doi.org/10.1038/s41598-023-36523-4 10
Vol:.(1234567890)
www.nature.com/scientificreports/
 29. Feng, C. et al. RNA-dependent RNA polymerases in the black-legged tick produce Argonaute-dependent small RNAs and regulate 
genes. bioRxiv 38, 2021.09.19.460923 (2021).
 30. Kurscheid, S. et al. Evidence of a tick RNAi pathway by comparative genomics and reverse genetics screen of targets with known 
loss-of-function phenotypes in Drosophila. BMC Mol. Biol. 10, 26 (2009).
 31. Arakane, Y. & Muthukrishnan, S. Insect chitinase and chitinase-like proteins. Cell. Mol. Life Sci. 67, 201–216 (2010).
 32. You, M. et al. Identification and molecular characterization of a chitinase from the hard tick Haemaphysalis longicornis. J. Biol. 
Chem. 278, 8556–8563 (2003).
 33. You, M. & Fujisaki, K. Vaccination effects of recombinant chitinase protein from the hard tick Haemaphysalis longicornis (Acari: 
Ixodidae). J. Vet. Med. Sci. 71, 709–712 (2009).
 34. Assenga, S. P. et al. The use of a recombinant baculovirus expressing a chitinase from the hard tick Haemaphysalis longicornis and 
its potential application as a bioacaricide for tick control. Parasitol. Res. 98, 111–118 (2006).
 35. Tufts, D. M. et al. Distribution, host-seeking phenology, and host and habitat associations of Haemaphysalis longicornis ticks, Staten 
Island, New York. USA. Emerg. Infect. Dis. 25, 792–796 (2019).
 36. Jia, N. et al. Haemaphysalis longicornis. Trends Genet. 37, 292–293 (2021).
 37. Duncan, K. T., Sundstrom, K. D., Saleh, M. N. & Little, S. E. Haemaphysalis longicornis, the Asian longhorned tick, from a dog in 
Virginia, USA. Vet. Parasitol. Reg. Stud. Reports 20, 100395 (2020).
 38. Casel, M. A., Park, S. J. & Choi, Y. K. Severe fever with thrombocytopenia syndrome virus: Emerging novel phlebovirus and their 
control strategy. Exp. Mol. Med. 53, 713–722 (2021).
 39. Yano, Y., Shiraishi, S. & Uchida, T. A. Effects of temperature on development and growth in the tick, Haemaphysalis longicornis. 
Exp. Appl. Acarol. 3, 73–78 (1978).
 40. Ghanbarian, H., Aghamiri, S., Eftekhary, M., Wagner, N. & Wagner, K. D. Small activating rnas: Towards the development of new 
therapeutic agents and clinical treatments. Cells 10, 1–13 (2021).
 41. Nuss, A., Sharma, A. & Gulia-Nuss, M. Genetic manipulation of ticks: A paradigm shift in tick and tick-borne diseases research. 
Front. Cell. Infect. Microbiol. 11, 1–7 (2021).
 42. Zhang, Y. et al. Liposome mediated double-stranded RNA delivery to silence ribosomal protein P0 in the tick Rhipicephalus 
haemaphysaloides. Ticks Tick. Borne. Dis. 9, 638–644 (2018).
 43. Umemiya-Shirafuji, R. et al. Data from expressed sequence tags from the organs and embryos of parthenogenetic Haemaphysalis 
longicornis. BMC Res. Notes 14, 14–17 (2021).
 44. Santos, V. T. et al. The embryogenesis of the Tick Rhipicephalus (Boophilus) microplus: The establishment of a new chelicerate 
model system. Genesis 51, 803–818 (2013).
 45. Hernandez, E. P. et al. Expression analysis of glutathione S-transferases and ferritins during the embryogenesis of the tick Haema-
physalis longicornis. Heliyon 6, e03644 (2020).
 46. Kulka, M., Alexopoulou, L., Flavell, R. A. & Metcalfe, D. D. Activation of mast cells by double-stranded RNA: Evidence for activa-
tion through Toll-like receptor 3. J. Allergy Clin. Immunol. 114, 174–182 (2004).
 47. Vasudevan, S. & Steitz, J. A. AU-rich-element-mediated upregulation of translation by FXR1 and Argonaute 2. Cell 128, 1105–1118 
(2007).
 48. Ward, K. A. & Fairbairn, D. Chitinase in developing eggs of Ascaris suum (Nematoda). J. Parasitol. 58, 546–549 (1972).
 49. Abriola, L. et al. Development and optimization of a high-throughput screening method utilizing Ancylostoma ceylanicum egg 
hatching to identify novel anthelmintics. PLoS ONE 14, 1–11 (2019).
 50. Moreira, M. F. et al. A chitin-like component in Aedes aegypti eggshells, eggs and ovaries. Insect Biochem. Mol. Biol. 37, 1249–1261 
(2007).
 51. Habeeb, S. M., Ashry, H. M. & Saad, M. M. Ovicidal effect of chitinase and protease enzymes produced by soil fungi on the camel 
tick Hyalomma dromedarii eggs (Acari:Ixodidae). J. Parasit. Dis. 41, 268–273 (2017).
 52. Zhu, Q., Arakane, Y., Beeman, R. W., Kramer, K. J. & Muthukrishnan, S. Functional specialization among insect chitinase family 
genes revealed by RNA interference. Proc. Natl. Acad. Sci. U. S. A. 105, 6650–6655 (2008).
 53. Mkandawire, T. T., Grencis, R. K., Berriman, M. & Duque-Correa, M. A. Hatching of parasitic nematode eggs: A crucial step 
determining infection. Trends Parasitol. 38, 174–187 (2022).
 54. Tan, C. P., Sinigaglia, L., Gomez, V., Nicholls, J. & Habib, N. A. RNA activation—a novel approach to therapeutically upregulate 
gene transcription. Molecules 26, 6530 (2021).
 55. Wilson, T. G. & Ashok, M. Insecticide resistance resulting from an absence of target-site gene product. Proc. Natl. Acad. Sci. U. S. 
A. 95, 14040–14044 (1998).
 56. Fujisaki, K. Maintenance of parthenogenetic Okayama strain of Haemaphysalis longicornis. Natl. Inst. Anim. Health Quart. (Yat-
abe) 18, 27–38 (1978).
 57. Umemiya-Shirafuji, R. et al. Transovarial persistence of Babesia ovata DNA in a hard tick, Haemaphysalis longicornis, in a semi-
artificial mouse skin membrane feeding system. Acta Parasitol. 62, 836–841 (2017).
 58. Umemiya-Shirafuji, R. et al. Hard ticks as research resources for vector biology: From genome to whole-body level. Med. Entomol. 
Zool. 70, 181–188 (2019).
 59. Neelakanta, G. & Sultana, H. Transmission-blocking vaccines: focus on anti-vector vaccines against tick-borne diseases. Arch. 
Immunol. Ther. Exp. (Warsz) 63, 169–179 (2015).
 60. Umemiya, R. et al. Haemaphysalis longicornis: Molecular characterization of a homologue of the macrophage migration inhibitory 
factor from the partially fed ticks. Exp. Parasitol. 115, 135–142 (2007).
 61. Anisuzzaman, et al. Longistatin in tick saliva blocks advanced glycation end-product receptor activation. J. Clin. Invest. 124, 
4429–4444 (2014).
 62. Yamaji, K. et al. HlCPL-A, a cathepsin L-like cysteine protease from the ixodid tick Haemaphysalis longicornis, modulated midgut 
proteolytic enzymes and their inhibitors during blood meal digestion. Infect. Genet. Evol. 16, 206–211 (2013).
 63. Anisuzzaman et al. Longistatin, a plasminogen activator, is key to the availability of blood-meals for ixodid ticks. PLoS Pathog. 7, 
e1001312 (2011).
 64. Maeda, H. et al. Establishment of a novel tick-Babesia experimental infection model. Sci. Rep. 6, 37039 (2016).
 65. Hatta, T. et al. Semi-artificial mouse skin membrane feeding technique for adult tick, Haemaphysalis longicornis. Parasites Vectors 
5, 263 (2012).
 66. Hatta, T. et al. Quantitative PCR-based parasite burden estimation of Babesia gibsoni in the vector tick, Haemaphysalis longicornis 
(acari: Ixodidae), fed on an experimentally infected dog. J. Vet. Med. Sci. 75, 1–6 (2013).
 67. Kilkenny, C., Browne, W. J., Cuthi, I., Emerson, M. & Altman, D. G. Improving bioscience research reporting: The ARRIVE 
guidelines for reporting animal research. Vet. Clin. Pathol. 41, 27–30 (2012).
 68. Percie du Sert, N. et al. The ARRIVE guidelines 2.0: Updated guidelines for reporting animal research. Br. J. Pharmacol. 177, 
3617–3624 (2020).
 69. Huang, X. & Madan, A. CAP3: A DNA sequence assembly program. Genome Res. 9, 868–877 (1999).
 70. Livak, K. J. & Schmittgen, T. D. Analysis of relative gene expression data using real-time quantitative PCR and the 2-ΔΔCT method. 
Methods 25, 402–408 (2001).
 71. Kocan, K. M., Blouin, E. & de la Fuente, J. RNA interference in ticks. J. Vis. Exp. 1, 1–5. https://d oi. org/ 10.3 791/2 474 (2010).
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Acknowledgements
The authors are grateful to the members of the Department of Parasitology and Tropical Medicine (KUSM).
Author contributions
Conceptualization, K.D.K. and T.H.; methodology, K.D.K. and T.H.; tick maintenance, H.K., Y.K., S.S., T.I., 
R.US., K.Y., T.T.,; software, K.D.K. and T.H.; validation, K.D.K. and T.H.; formal analysis K.D.K.; investiga-
tion, K.D.K., F.M., and T.H.; resources, T.H.; data curation, K.D.K.; writing—original draft preparation, K.D.K.; 
writing—review and editing, E.P.H., A., H.K., D.L., M.M., M.A.A., S.K.D., N.T. and T.H.; visualization, K.D.K.; 
supervision, T.H.; project administration, T.H., funding acquisition, T.H. All authors have read and agreed to 
the published version of the manuscript.
Funding
This research was funded by KAKENHI (19F19410, 20H03001, 22H05059 to T.H.) from The Japan Society for 
the Promotion of Science, a grant from Oshimo Foundation on FY2018 to T.H.
Competing interests 
The authors declare no competing interests.
Additional information
Supplementary Information The online version contains supplementary material available at https://d oi. org/ 
10.1 038/s 41598- 023- 36523-4.
Correspondence and requests for materials should be addressed to T.H.
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