(2019) 12:205
Umemiya‑Shirafuji et al. Parasites Vectors
https://doi.org/10.1186/s13071-019-3469-9

Parasites & Vectors
Open Access

RESEARCH

Intracellular localization of vitellogenin
receptor mRNA and protein during oogenesisof a parthenogenetic tick, Haemaphysalis
longicornis
Rika Umemiya‑Shirafuji1†, Ryo Mihara1†, Kozo Fujisaki1,2 and Hiroshi Suzuki1*

Abstract
Background: Vitellogenin (Vg), a key molecule for oocyte development synthesized in the fat body during bloodfeeding, is released into the hemolymph and then taken into the oocytes via Vg receptor (VgR) in ticks. Previously,
we showed that VgR mRNA is expressed in the ovary at the adult stage of parthenogenetic Haemaphysalis longicornis
ticks and its expression increases after blood-feeding. However, intracellular localization of VgR mRNA and protein ateach developmental stage of oocytes during oogenesis remains largely unclear.
Methods: mRNA and protein expression profiles of H. longicornis VgR (HlVgR) in the oocytes from the unfed to ovi‑
position periods were analyzed by real-time PCR, in situ hybridization, and immunostaining. To elucidate the timing of
the onset of Vg uptake, RNA interference (RNAi)-mediated gene silencing of HlVgR was performed.
Results: In situ hybridization revealed that HlVgR mRNA was detected in the cytoplasm of stage I-III oocytes, andweaker positive signals for HlVgR mRNA were found in the cell periphery of stage IV and V oocytes. Likewise, HlVgR
protein was detected by immunostaining in the cytoplasm of stage I-III oocytes and in the cell periphery of stage IV
and V oocytes. Each developmental stage of the oocytes showed distinct patterns of mRNA and protein expression of
HlVgR. Moreover, RNAi of HlVgR caused delayed or arrested development in the oocytes. The ovaries of control ticks
showed all developmental stages of oocytes, whereas stage I-III oocytes were found in the ovaries of HlVgR-RNAi ticks
at 5 days after engorgement.
Conclusions: These results suggest that active uptake of Vg is required for development from stage III to stage IV
during oogenesis. Our data clearly revealed an apparent shift in the intracellular localization of VgR for both mRNAand protein level in oocytes during oogenesis.
Keywords: Tick, Haemaphysalis longicornis, Vitellogenin receptor, Ovary, Oocyte*Correspondence: hisuzuki@obihiro.ac.jp†
Rika Umemiya-Shirafuji and Ryo Mihara contributed equally to this work
1
National Research Center for Protozoan Diseases, Obihiro Universityof Agriculture and Veterinary Medicine, Inada-Cho, Obihiro, Hokkaido
080‑8555, Japan
Full list of author information is available at the end of the article© The Author(s) 2019. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License(http://creat​iveco​mmons​.org/licen​ses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium,provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license,
and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creat​iveco​mmons​.org/
publi​cdoma​in/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

BackgroundTicks, vectors of harmful pathogens for human and animals, absolutely require a blood meal for their development and reproduction. Nutrients derived from a blood
meal allow the synthesis of vitellogenin (Vg), an essential molecule for oocyte development, in the fat body offemale ticks, being stimulated by ecdysteroid hormones[1–10]. The regulatory network underlying the synthesisof Vg (vitellogenesis) in ticks is being clarified by RNA
interference (RNAi)-mediated gene knockdown experiments and inhibitor assays. A GATA factor was foundto be a transcriptional activator of the Vg gene in thefemale Haemaphysalis longicornis [11]. Subsequent studies revealed that vitellogenesis is controlled by the activation of the target of rapamycin (TOR), a key moleculeof a nutrient-sensing pathway in eukaryotic cells, throughecdysteroid hormone stimulation [12]. TOR phosphorylates S6 kinase and regulates the transcription and intracellular localization of a GATA factor in the fat body. Our
previous data suggested that the serine/threonine protein
kinase Akt is an upstream factor of TOR and essential
not only for controlling gene transcription but also for
regulating vitellogenesis [13].
Following vitellogenesis, Vg is released into thehemocoel filled with hemolymph and then is taken
into developing oocytes in ticks [7, 10]. The absorptionof Vg into the oocytes occurs via receptor-mediatedendocytosis, and the Vg receptor (VgR) is a member ofthe low-density lipoprotein receptor (LDLR) superfamily [14]. In hard ticks, homologues of VgR have been
identified from Amblyomma hebraeum (AhVgR) [15],Dermacentor variabilis (DvVgR) [16], H. longicornis(HlVgR) [17], Rhipicephalus microplus (RmVgR) andR. appendiculatus (RaVgR) [18]. These tick VgRs havecommon structural elements similar to those in insects:
LDLR class A repeats, LDLR class B repeats, epidermal growth factor (EGF) precursor homology domains
containing YWXD repeats, an O-linked carbohydrate
domain, a transmembrane domain and a cytoplasmic
domain [14–18]. The predicted molecular weight oftick VgR based on the amino acid sequences is approximately 200 kDa [15–18]. These studies showed thatknockdown of VgR in female ticks led to the failureof Vg uptake in the oocytes and the subsequent oviposition, even though the ticks engorged completely.
Immunoelectron microscopy revealed that VgR is localized on the external surface of the plasma membrane
in developing tick oocytes [17]. In addition, whole tissue in situ hybridization using the ovaries of engorgedfemales at 10 days following engorgement revealed
that VgR mRNA was detected in the smallest oocytes,
which had not yet begun to fill with yolk particles [15].

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However, little is known about intracellular localizationof VgR mRNA and protein at each developmental stage,
namely stages I to V, of oocytes throughout oogenesis.
Transmission of some pathogens is tightly linked to
the reproduction system in female ticks. After ingesting a blood meal containing some pathogens, ticks
may give rise to progeny with these pathogens. An
ixodid female tick generally lays a few thousand eggs
[19]; therefore, the basis of oogenesis is an important
key for the development of novel control strategies forpathogen transmission by ticks. For instance, it has
been reported that VgR might be involved in transovarial transmission of Babesia parasites by a vector tickH. longicornis [17]. This finding led to the hypothesisthat Babesia parasites bind to tick VgR and invade thedeveloping oocytes of ticks and that some surface molecules of parasites have ligand-binding activity for VgR.
Therefore, it is important to determine the expression
pattern and intracellular localization of VgR to elucidate the interactions between Babesia parasites andtick VgR during oogenesis.
In the present study, we examined mRNA and protein
expression profiles of HlVgR in the oocytes from theunfed to the oviposition periods by real-time PCR andin situ hybridization and immunostaining using ovariantissue sections. Furthermore, to determine the timingof the onset of Vg uptake during oogenesis, oocytes of
HlVgR-RNAi ticks were observed histologically on thebasis of the classification criteria [20] for developing
oocytes in the parthenogenetic H. longicornis.

Results
Expression of HlVgR mRNA in the ovary

Real-time PCR with specific primers was conducted
to determine the relative expression level of HlVgR in
the ovary from the unfed to the oviposition periods.
Average value of HlVgR mRNA expression was 0.005
in the unfed period. By contrast, the expression at theslow feeding was more than 1000-fold higher than thatat the unfed period (t(5) = − 6.957, P = 0.0009) (Fig. 1).
The expression levels of HlVgR in the rapid feeding period were higher than those at previous periods (t(5) = − 8.798, P = 0.0003) and were the highest atengorgement (t(5) = − 13.761, P < 0.0001). These results
suggest that HlVgR was upregulated in the ovary during blood-feeding. In addition, expression levels of
HlVgR at 4 days after engorgement (4dAE) (t(10) = 4.487,
P = 0.0012) and at the beginning of the oviposition
period (0dAO) (t(10) = 4.707, P = 0.0008) decreased significantly compare with those at engorgement. At 10
days after the beginning of oviposition (10dAO), HlVgR
expression appeared to be increased again.

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

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Fig. 1 mRNA expression of HlVgR in the ovaries of H. longicornis from unfed to oviposition periods. Total RNA was extracted from each ovary
sample. DNase-treated RNA was subjected to cDNA synthesis. Data were normalized by a real-time PCR analysis of HlP0 mRNA levels in the cDNA
samples. Data represent the mean ± SD of triplicate samples. Asterisk and hash symbols indicate significant downregulation (Student t-test) and
upregulation (Welchʼs t-test), respectively, at P < 0.01. Abbreviations: 4dAE, 4 days after engorgement; 0dAO, the beginning of oviposition; 10dAO, 10
days after the beginning of oviposition

Localization of HlVgR mRNA in the ovary

HlVgR mRNA was undetected by in situ hybridization in
the ovaries in the unfed period (Fig. 2). The cytoplasm ofoocytes at stages I and II was stained with the antisense
probe of HlVgR in the slow feeding, rapid feeding, andengorgement periods. The expression of HlVgR mRNA in
oocytes of all stages (stages I to V) was detected at 4dAE,
0dAO, and 10dAO. HlVgR mRNA was localized throughout the cytoplasm in the oocytes of stages I to III. Positive signals for HlVgR mRNA were found in the external
region of the oocytes that were at stages IV and V, but
the signals seemed to be weaker than those of the former
stages. The expression of HlVgR mRNA was also detectedin pedicel cells and the ovarian wall (Fig. 2).
Localization of HlVgR protein in the ovary

Positive signals of HlVgR were not detected by immunofluorescence staining in the ovary in the unfed period(Fig. 3a) but were found in the cytoplasm of oocytes
in the slow feeding period. In the rapid feeding andengorgement periods, HlVgR signals were detected
throughout the cytoplasm of stage I and II oocytes. The
HlVgR signals were also detected in stage II to V oocytes
at 4dAE, 0dAO, and 10dAO (Fig. 3b). HlVgR signals were

scarcely observed in stage I oocytes during these periods.
HlVgR signals were detected throughout the cytoplasmof the oocytes of stage II. Notably, the signals were localized throughout the cytoplasm as well as external regionof the stage III oocytes, whereas the signals were detected
at the chorion and the cell periphery in the oocytes ofstages IV and V. In addition, HlVgR signals were localized
in the cytoplasm surrounding the yolk granules. HlVgR
signals were not detected in the pedicel cells and ovarian
wall at any period examined in our study.
Oocyte development in HlVgR‑RNAi ticks

To clarify the timing of the onset of Vg uptake during oogenesis, oocytes in the ovaries were observed
after RNAi of the HlVgR gene. Real-time PCR revealed
decreased expression levels of HlVgR mRNA at engorgement (t(2) = 16.327, P = 0.0037) and at 5 days after
engorgement (t(2) = 12.749, P = 0.0061) compared withthat in the control ticks (Fig. 4a, c). There was no difference in the body weight of engorged ticks between the
HlVgR-RNAi and control groups (Table 1). The oocytes
were typically colorless and white in color in both the
HlVgR-RNAi and control ticks at engorgement (Fig. 4b).
At this period, oocytes of stages I and II were observed

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

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Fig. 2 Expression sites of HlVgR mRNA in H. longicornis ovary from unfed to oviposition periods. Paraffin sections of the ovaries were treated withsense probe (S) or antisense probe (AS). Arrowheads show pedicel cells. Abbreviations: W, ovarian wall; I, stage I oocyte; II, stage II oocyte; III, stage III
oocyte; IV, stage IV oocyte; V, stage V oocyte. Scale-bars: 50 µm

in both ovaries on the histological sections. Control ticks
had amber-colored oocytes in the ovaries and began oviposition at 5 days after engorgement, whereas the ovaries
in the HlVgR-RNAi ticks possessed amberlite and whitecolored oocytes, which were classified into stages I to III(Fig. 4d). No yolk granules were found in the cytoplasmof any of the oocytes of the HlVgR-RNAi ticks. Although
HlVgR-RNAi ticks began oviposition at 12 days after
engorgement (Table 1), the eggs were not in amber colorand not coated with waxy components as shown previously [17].

Discussion
Expression levels of HlVgR mRNA in the ovary of H. longicornis increased with the progress of blood-feeding andreached the highest level at engorgement, suggesting thatblood-feeding is a triggering process for the upregulationof HlVgR mRNA (Fig. 1). Because ticks have a panoistic
ovary, the ratio of each oocyte stage is asynchronous during oogenesis [20]. Subsequently, the spatial patterningof HlVgR mRNA expression in each developmental stageof the oocytes was examined by in situ hybridization.
The cytoplasm of oocytes at stages I and II was intensely
stained with the antisense probe of HlVgR at the slow
feeding, rapid feeding, and engorgement periods (Fig. 2).
At these periods, the ovaries of parthenogenetic H. longicornis possess stage I and II oocytes, and the ratio of stage
II to all oocytes in the rapid feeding period increased atengorgement [20]. In addition, the HlVgR mRNA level
was significantly decreased at 4dAE and at 0dAO compared with that at engorgement (Fig. 1). At 4dAE and
0dAO, more than half of all oocytes were classified
into stages III, IV or V [20]. By linking mRNA expression patterns with morphological features of oocytes, it
was obvious that HlVgR was highly expressed in stage
I and II oocytes compared to the other stages (Figs. 1
and 2 and [20]). These findings substantially correspondwith the findings that a high degree of positive staining
was detected in the smallest oocytes of A. hebraeum bywhole tissue in situ hybridization [15]. Moreover, HlVgR

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

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Fig. 3 Localization of HlVgR protein in the ovary of H. longicornis. Positive signals are displayed in yellow color in the ovarian sections from unfed toengorgement periods (a) and from 4 days after engorgement to oviposition periods (b). Stage I or II oocytes at unfed to engorgement periods wereobserved in areas surrounded by dashed circles. Abbreviations: PC, phase contrast; I, stage I oocyte; II, stage II oocyte; III, stage III oocyte; IV, stage IV
oocyte; V, stage V oocyte. Scale-bars: 50 µm

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

Fig. 4 HlVgR-knockdown by RNA interference (RNAi) in H. longicornis.
a, c Total RNA was purified from engorged ticks in each group.
The gene silencing of HlVgR was assessed by real-time PCR. The
HlVgR expression was normalized to the HlP0 mRNA expression.
Data represent the mean ± SD of triplicate samples. HlVgR mRNA
expressions of RNAi-treated ticks (HlVgR RNAi) were significantly lower
than those of control ticks (Control) at both engorgement (a) and at 5days after engorgement (c) (*P < 0.01; Welchʼs t-test). b, d Phenotypesof the ovaries of HlVgR RNAi-treated and control ticks. Note that
HlVgR-knockdown caused delayed development in the oocytes
at both engorgement (b) and at 5 days after engorgement (d).
Abbreviations: I, stage I oocyte; II, stage II oocyte; III, stage III oocyte;
IV, stage IV oocyte; V, stage V oocyte. Scale-bars: b, 1 mm for ovaries
and 100 µm for ovarian sections; d, 2 mm for ovaries, and 200 µm forovarian sections

Page 6 of 12mRNA was detected in the whole cytoplasm of stage I to
III oocytes and under the oocyte membrane of oocytes atstages IV and V in our study (Fig. 2). The positive signalsof HlVgR mRNA in mature oocytes seemed to be weaker
than those of stage I-III oocytes. Taken together, realtime PCR and in situ hybridization showed that the transcription of HlVgR was activated in stages I and II of theoocytes, namely before the formation of yolk granules,
regardless of the number of days after blood-feeding.
Immunostaining revealed that HlVgR was localized
throughout the cytoplasm of the stage I–III oocytes andin the peripheral region of the stage IV and V oocytes(Fig. 3). As with the results of in situ hybridization, theintracellular localization of HlVgR changed as oocyte
development progressed. Drosophila melanogaster VgR
is uniformly distributed throughout the oocyte during the pre-vitellogenic stages of oogenesis, indicatingthat such oocytes do not take in Vg in the presence ofVgR [21]. After the transition to the vitellogenic stages,
VgR levels increase markedly at the cell periphery in theoocyte. By the end of vitellogenesis, the receptor localizes
predominantly in the external region of the oocyte in the
fruit fly. In the yellow fever mosquito Aedes aegypti, VgR
is internalized, dissociated, and sorted and then recycled
to the plasma membrane during Vg uptake [22]. In ourstudy, we found that HlVgR was distributed throughout
the cytoplasm of stage I and II oocytes (Fig. 3), suggestingthat such oocytes are unlikely to take up Vg from hemolymph. In the stage III oocytes, positive signals weredetected throughout the cytoplasm and in the external
region of the oocytes. HlVgR was localized in the cytoplasm, especially in the peripheral region of the stage
IV–V oocytes, which possess yolk granules in the cytoplasm [20].
Knockdown of HlVgR caused the delay or arrest ofoogenesis in our study (Fig. 4), resulting in the inhibition of oocyte development from stage III to the subsequent stage. Control ticks started to lay eggs at 5 days
after engorgement, whereas the oocytes of the RNAitreated ticks at that timing were still developing to stage
III. The ovaries of the RNAi-treated ticks showed anamberlite color, implying that Vg uptake in the oocytes
starts from stage III. Seixas et al. [18] similarly reported

Table 1 Effects of HlVgR-knockdown by RNA interference (RNAi) in oocyte development of H. longicornisGroupsNo. of ticks

Control20

HlVgR RNAi

15

Body weight of
Typical color of oocytes
engorged ticks (mg)239.0 ± 45.7

208.3 ± 52.4

Developmental stages ofoocytes

Onset of oviposition

E

5dAE

E

5dAE

Colorless and white

Amber

Stages I–II

Stages I–V

At 5 days after engorgement

Colorless and white

Amberlite
and white

Stages I–II

Stages I–III

At 12 days after engorgement

Abbreviations: E, engorgement; 5dAE, 5 days after engorgement

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

that RmVgR-silenced female ticks showed a small degreeof yolk uptake in the oocytes classified into the ovarian
growth phase scores/indexes 2 and 3. They also showed aclear correlation between the amount of RmVgR mRNAand protein in the ovaries: RmVgR protein level wasincreased after the upregulation of mRNA expression
in the ovaries of partially fed female ticks, and the level
peaked before engorgement. Moreover, they describedthat when female ticks drop from the host animal, VgRmRNA expression levels decrease because the receptor
is no longer important once Vg has already been taken
up by the oocytes. However, the Vg concentration in thehemolymph reaches its highest level at 2–5 days after the
beginning of oviposition in hard ticks [23–25]. As shownin our study, the expression level of VgR mRNA depends
on the developmental stages of the oocytes. Therefore,
the decrease of the VgR mRNA level was considered to
be attributed to the decrease in the ratio of stage I and II
oocytes to all oocytes. Oogenesis continues to progress
throughout the oviposition period of around 1 month
until the female ticks die.
Nurse or follicle cells, the providers of nutrients for
insect oocytes, are non-existent in the ovary of ticks.
Instead of such nurse or follicle cells, pedicel cells derived
from ovarian wall epithelial cells are present in ticks.
These pedicel cells are known to attach the oocytes tothe ovarian wall [26, 27]. In addition to their structurally supportive role, it was speculated that tick pedicel
cells have similar functions of nurse or follicle cells in
insects. Ultrastructural observations led to the idea that

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yolk precursors, i.e. Vg, might be synthesized in pedicel
cells of A. triste ticks and would then be transported into
stage II oocytes [26]. In D. melanogaster, VgR mRNA istransported to the oocyte after synthesis in the nurse
cells [21]. We found positive signals for HlVgR mRNA in
both pedicel cells and the ovarian wall at the slow feeding, pre-oviposition, and oviposition periods (Fig. 2).
HlVgR mRNA could be synthesized in the pedicel cells;
however, further experiments are needed to clarify thispoint because HlVgR protein was not detected in pedicel
cells and the ovarian wall by immunostaining.

Conclusions
Our study revealed that an apparent shift in the intracellular localization of HlVgR mRNA and protein was
associated with the developmental stages of oocytes inparthenogenetic H. longicornis ticks. This study showedthat Vg uptake via VgR is a critical process for the transition of oocytes from stage III to IV in the tick. Taken
together with findings from our previous studies [11–13, 17, 20, 28], we propose a model of Vg uptake during oogenesis in parthenogenetic H. longicornis (Fig. 5).
To date, multiple Vgs (HlVg-1, HlVg-2, and HlVg-3)
were identified from the parthenogenetic H. longicornis[28]. This study revealed that the HlVg-1 transcript was
detected only in the midgut of this tick species. The
HlVg-2 was expressed in the ovary and fat body, while
the HlVg-3 was found only in the fat body. “Vgs” in Fig. 5
includes HlVg-2 and HlVg-3 proteins based on our previous results. In other tick species, DvVg1 mRNA wasFig. 5 A schema of Vg uptake during oogenesis of parthenogenetic H. longicornis. Synthesized Vgs (Vg-2 and Vg-3) are released from the fat body
into the hemocoel filled with hemolymph. The Vgs are taken up by oocytes of stages III, IV and V. Abbreviations: I, stage I oocyte; II, stage II oocyte; III,stage III oocyte; IV, stage IV oocyte; V, stage V oocyte; Vgs, vitellogenins; VgR, vitellogenin receptor

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

detected in the midgut, fat body, and ovary of D. variabilis [29, 30]. However, DvVg2 mRNA was detected inboth fat body and midgut but not found in the ovary.
mRNA expression of AhVg1was observed in the fat body,
midgut, and ovary, while AhVg2 was detected in both fat
body and midgut in A. hebraeum [31]. In R. microplus,
Vg-1 and Vg-2 are synthesized in the ovary [32]. The contribution of each Vg to vitellogenesis remains unclear;
however, ovarian molecules are considered as targets
for anti-tick vaccine development as they can affect tickbiology by decreasing oogenesis and embryogenesis [33].
After blood-feeding, immunoglobulins pass through the
midgut epithelium and then arrive in the hemolymph ofarthropods while retaining their immunological characters [34]. It is believed that cell membrane receptors
serve as vaccine candidate molecules in blood-feedingarthropods. Further understanding of the molecularmechanisms regulating oogenesis will provide valuable
information for the control of ticks. Moreover, our current findings may help to clarify the interaction between
transovarially transmitted pathogens and ovarian molecules, especially Vg or VgR, in ticks. The potential of
HlVgR as an antigen for the development of an anti-tick
vaccine could induce infertility in ticks and thus inhibit
the transmission of tick-borne diseases.

Methods
Ticks and animals

The parthenogenetic H. longicornis ticks (Okayama
strain) were maintained at the National Research Center
for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan. The female
ticks were fed on Japanese white rabbits (Japan SLC, Shizuoka, Japan) as described previously [35]. Eight-weekold female BALB/c mice (CLEA Japan, Tokyo) were used
to produce anti-HlVgR polyclonal antibody. The rabbits
were reared in a temperature- and humidity-regulatedroom under controlled lighting, water and commercial
regular chow. The mice were maintained in a temperature- and humidity-regulated P2 room under same conditions as rabbits.
Reverse transcription and real‑time PCR

For collection of the ovaries, 30 ticks at the unfed, 20
ticks at slow feeding (3 days after attachment), 5 ticks at
rapid feeding (4–6 days after attachment), and 3 ticks at
engorgement (5–7 days after attachment) periods weredissected. Twelve engorged females in individual glass
tubes were placed in a container. The container was thenplaced in dark incubator at 25 °C and saturated humidity. The ovaries were also sampled from 4 ticks at 4 days
(4dAE; pre-oviposition), 4 ticks at 7 days (0dAO; beginning of oviposition) and 4 ticks at 17 days (10dAO) after

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engorgement. Total RNA was extracted from the collected ovaries using TRI® reagent (Sigma-Aldrich, St.
Louis, MO, USA) and incubated with DNase (TURBO
DNA-free™ Kit; Life Technologies, Frederick, MD, USA)to remove genomic DNA. Single-strand cDNA was synthesized using purified total RNA and a ReverTra Ace®
qPCR RT Kit (Toyobo, Osaka, Japan) according to aprevious report [36]. Subsequently, real-time PCR was
conducted using a 7300 Real-Time PCR System (LifeTechnologies) and THUNDERBIRD® SYBR qPCR Mix(Toyobo) according to the manufacturer’s protocol. The
primers used in our study are listed in Table 2. Standard
curves used for the calculation of relative gene expression were created from five-fold serial dilutions of cDNA
derived from the ovaries of engorged females. The PCR
conditions were as follows: 95 °C for 10 min, 40 cycles ofdenaturation at 95 °C for 15 s, and annealing/extension
at 60 °C for 60 s. Dissociation curves for individual wells
were created to confirm the single PCR product. Data
were analyzed by the 7300 system SDS software version
1.4 for Windows 7. The control amplification was carriedout using the primers designed for H. longicornis ribosomal protein P0 (HlP0) (GenBank: EU048401). All reactions were run in duplicate, and data obtained were used
to calculate the mean ± standard deviation (SD). The
differences in relative gene expression levels of HlVgR
between each sample were compared using the Student
t-test (vs Unfed) or Welchʼs t-test (vs Engorgement),
depending on the data distribution. Statistical significance was set at P < 0.01.
Preparation of histological sections of ovary

Thirty-two ovaries in total collected at the same periods
as described in the previous section were fixed with 4%paraformaldehyde-0.1% glutaraldehyde in phosphatebuffered saline (PBS) at 4 °C, overnight. The ovaries (10µm thick) were sectioned with a cryostat (CM3050 S,
Leica Biosystems, Wetzlar, Germany) as described previously [37]. To prepare paraffin sections, a total of 55
ovaries were immersed in 4% paraformaldehyde-2.5%
glutaraldehyde in PBS at 4 °C, overnight. After washingwith PBS, the ovaries were pre-embedded in 3% agarose(Sigma-Aldrich) in PBS and then embedded in paraffin(Sakura Finetek Japan, Tokyo, Japan). The sections (5 µm
thick) were cut on a REM-700 microtome (Yamato Kohki
Industrial, Saitama, Japan).
In situ hybridization

In situ hybridization was carried out according to the
manufacturer’s protocol of the in situ hybridization reagents starting kit (Nippon Gene, Tokyo, Japan). RNAprobes were generated from PCR products with T7 or
SP6 ends produced by amplification of ovary cDNA usingUmemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

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Table 2 Primers used in this study
Experiment

Description

Sequence (5′–3′)Real-time PCR

HlP0-Fwd

CTC​CAT​TGT​CAA​CGG​TCT​CA

In situ hybridization

Expression of recombinant HlVgR protein

HlP0-Rev

TCA​GCC​TCC​T TG​AAG​GTG​AT

HlVgR-Fwd

GAC​CGT​TCT​TGC​T TC​TGT​CC

HlVgR-Rev

GCG​ACC​ATC​ACA​CCA​GTA​GA

T7-HlVgR-1-Fwd

TAA​TAC​GAC​TCA​C TA​TAG​GCT​GTC​CCT​CGG​ATG​ACT​TCT​CCT​G

HlVgR-1-Rev

CCC​CAC​AGT​CGA​TCA​TGC​CA

HlVgR-1-Fwd

CTG​TCC​C TC​GGA​TGA​C TT​C TC​C TG​

SP6-HlVgR-1-Rev

ATT​TAG​GTG​ACA​C TA​TAG​CCC​CAC​AGT​CGA​TCA​TGC​C

XhoI-HlVgR-Fwd

CCG​C TC​GAG​ATC​TAC​TGG​TGT​GAT​GGT​

EcoRI-HlVgR-Rev

GGA​ATT​CAC​AGT​GGC​TGC​T TT​CGT​

gene-specific primers (Table 2). The PCR was performed
by a GeneAmp® PCR System 9700 (Life Technologies)
and PrimeSTAR® MAX DNA Polymerase (Takara Bio,
Shiga, Japan). The cycling conditions were as follows:
30 cycles of denaturation at 98 °C for 10 s and annealing/extension at 68 °C for 60 s. The PCR products (534bp) were purified using the QIAquick Gel Extraction Kit(Qiagen, Hilden, Germany). Sense and antisense RNAprobes were generated using purified template DNA,
T7, and SP6 RNA polymerase, respectively, and digoxigenin-11-UTP (Roche Diagnostics GmbH, Mannheim,
Germany) for 60 min at 37 °C (for T7 RNA polymerase)and 40 °C (for SP6 RNA polymerase). Synthesized RNAprobes were confirmed by electrophoresis on a 1.5%
agarose gel and were then treated with alkali hydrolysis
buffer for 44 min at 60 °C.
Paraffin sections of the ovaries were deparaffinized
and treated with 5–8 µg/ml proteinase K (Nippon Gene)for 20 min at 37 °C. They were acetylated for 15 min atroom temperature by adding 1 ml of acetic anhydride
into 400 ml of acetylation buffer and then pre-hybridized
in hybridization solution for 30 min at 42 °C. Hybridization was performed for 16 h at 42 °C after adding 1 µg/
ml of sense or antisense probe per section. Hybridized
sections were treated with blocking reagent (Roche Diagnostics GmbH) for 30 min at room temperature, treatedwith anti-digoxigenin-AP Fab fragments (Roche Diagnostics GmbH) for 1 h at room temperature, and thenreacted with BCIP®/NBT liquid substrate system (SigmaAldrich) for 24 h at room temperature. Following staining of hybridized probes, nucleus staining was performed
with 1% methyl green (Muto Pure Chemicals, Tokyo,
Japan) for 2 min at room temperature. The sections werethen observed under a BZ-9000 light microscope (Keyence, Osaka, Japan). To determine expression sites of
HlVgR mRNA in the ovary, the developmental stages ofeach oocyte were classified into five stages as described
previously [20].
Expression of recombinant HlVgR protein and productionof mouse anti‑HlVgR polyclonal antibody

A partial fragment (936 bp) corresponding to second
ligand-binding domain with class A repeats of HlVgR(GenBank: AB299015) [17] was amplified by PCR withthe forward primer containing a recognition site for
XhoI and the reverse primer with a recognition site for
EcoRI (Table 2). The PCR was conducted by a GeneAmp®
PCR System 9700 (Life Technologies) and PrimeSTAR®
MAX DNA Polymerase (Takara Bio). The cycling conditions were as follows: 30 cycles of denaturation at 98 °C
for 10 s and annealing/extension at 68 °C for 60 s. The
PCR product (952 bp) containing partial HlVgR and
restriction enzyme sites purified by Wizard® SV Gel
and PCR Clean-up System (Promega, WI, USA) was
inserted into the pCold™ ProS2 DNA plasmid vector(Takara Bio), which had been digested with XhoI andEcoRI, with a Ligation-Convenience Kit (Nippon Gene)according to the manufacturer’s protocol. Recombinant
plasmids were transformed into ECOS™-competent
Escherichia coli DH5α (Nippon Gene). After transformation, the plasmids were confirmed by sequencing andused to transform ECOS™-competent E. coli BL21 (Nippon Gene). Expression of recombinant histidine-tagged
HlVgR (rHlVgR; approximately 60 kDa) was inducedwith 0.1 mM isopropyl-β-D (-)-thiogalactopyranosideat 15 °C overnight. The rHlVgR was solubilized withbinding buffer (pH 8.0) containing 8 M urea. The purification of rHlVgR was performed by His GraviTrap (GEHealthcare, Chicago, IL, USA) using elution buffer (pH8.0) containing 8M urea under the denaturing conditions. After concentration of the purified rHlVgR using
Pierce® Concentrator 9K MWCO (Thermo Fisher Scientific, Waltham, MA, USA), urea in the rHlVgR solution

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

was removed using Slide-A-Lyzer™ G2 Dialysis Cassette
7K MWCO (Thermo Fisher Scientific). Fifty microgramsof soluble rHlVgR or PBS in Freund’s complete adjuvant (Sigma-Aldrich) was intraperitoneally injected into
8-week-old female BALB/c mice. The same antigen or
PBS in Freund’s incomplete adjuvant (Sigma-Aldrich)
was repeatedly injected intraperitoneally into the mice ondays 14, 28 and 42 after the first inoculation. At 7 days
after the last immunization, sera against rHlVgR werecollected from mice. Anti-rHlVgR IgG in the sera waspurified using the Melon™ Gel IgG Spin Purification Kit(Thermo Fisher Scientific) and was stored at − 30 °C until
use.
Immunostaining

Frozen sections were incubated with Liberate Antibody
Binding solution (Polysciences, Warrington, PA, USA)for 5 min at room temperature for antigen retrieval.
After washing with PBS, the sections were blocked in
10% goat serum in PBS at 4 °C overnight. The sections
were then incubated with anti-rHlVgR IgG that had been
diluted in 5% goat serum in PBS (1:10) for 1 h at 37 °C.
Subsequently, the sections were washed with PBS andthen reacted with Alexa Fluor® 594 goat anti-mouse IgG(H + L) (1:4000; Life Technologies) for 1 h at 37 °C. The
sections were then mounted in ProLong® Gold Antifade
Mountant with DAPI (Life Technologies) after washing with PBS. Fluorescent signals were observed under a
TCS SP5 confocal laser microscope (Leica Microsystems,
Wetzlar, Germany), and the signals were displayed in yellow color to allow clearer detection of signals.
Histological observation of ovary in HlVgR‑RNAi ticks

A 631-bp HlVgR fragment was amplified by PCR using
T7 forward (5’-TAA TAC GAC TCA CTA TAG GCTGTC CCT CGG ATG ACT TCT CCT G-3’) and T7
reverse (5’-TAA TAC GAC TCA CTA TAG GTG
CAG TTG CTC TCG TCC CGG CCG-3’) primers as
described previously [17]. The PCR products were
purified as described above and were then used togenerate double-stranded RNA (dsRNA) using the T7
RiboMax™ Express RNAi System (Promega). Thirtyfive female ticks fixed on a glass slide with adhesive
tape were injected with 0.5 μl of 2 μg/μl (1 μg/tick) of
the dsRNA of HlVgR or firefly luciferase as described
previously [12]. Injected ticks were kept at 25 °C for
18 h in an incubator and then fed on rabbits. Twenty
and fifteen engorged ticks of control and HlVgR RNAi
groups, respectively, were collected from the rabbits
and their body weight was measured. Statistical significance was assessed by the Student t-test. The ticks werethen incubated in a chamber at 25 °C for monitoring ofPage 10 of 12

oviposition. The female ticks were dissected to collect 6
ovaries at the engorgement (5–7 days after attachment)
and 3 ovaries at 5 days after engorgement. The ovaries
immersed in PBS were morphologically observed undera SZX16 stereomicroscope (Olympus, Tokyo, Japan)
with a DP21 camera (Olympus). After observation, theovaries were immersed into 4% paraformaldehyde-0.1%
glutaraldehyde in PBS at 4 °C, overnight, and then subjected to the preparation of frozen sections as describedabove. The frozen sections were stained with hematoxylin and eosin (Muto Pure Chemicals). The stained sections were then observed under a BZ-9000 microscope(Keyence). Moreover, total RNA was purified from theremaining ticks in each group and used to synthesize
cDNA as described above. The gene silencing of HlVgR
was assessed by real-time PCR using a 7300 Real-Time
PCR System (Life Technologies) and THUNDERBIRD®
SYBR qPCR Mix (Toyobo) as described above. The
HlVgR expression was normalized to the HlP0 mRNA
expression. The expressions for control group were set
to 1.0, and relative expressions for HlVgR RNAi groupwere calculated. The data represented the mean ± SD of
triplicate samples. The differences between control and
HlVgR RNAi groups were compared using the Welchʼst-test. Statistical significance was set at P < 0.01.
Abbreviations
AhVgR: Amblyomma hebraeum vitellogenin receptor; dsRNA: double-stranded
RNA; DvVgR: Dermacentor variabilis vitellogenin receptor; HlVgR: Haemaphysalis longicornis vitellogenin receptor; LDLR: low-density lipoprotein receptor;
PBS: phosphate-buffered saline; RaVgR: Rhipicephalus appendiculatus vitel‑
logenin receptor; RmVgR: Rhipicephalus microplus vitellogenin receptor; RNAi:
RNA interference; SD: standard deviation; TOR: target of rapamycin; Vg: Vitello‑genin; VgR: Vitellogenin receptor; 4dAE: 4 days after engorgement; 0dAO: thebeginning of oviposition; 10dAO: 10 days after the beginning of oviposition.
Acknowledgements
We thank Dr. Yasuyuki Abe (Prefectural University of Hiroshima) for assistancewith methodology and Dr. Damdinsuren Boldbaatar (Mongolian University ofLife Sciences) for advice on the manuscript.
Funding
This work was supported by a Grant-in-Aid for Young Scientists (B) from theJapan Society for the Promotion of Science (16K18794) and the Akiyama Life
Science Foundation. This work was partially supported by The Japan Agency
for Medical Research and Development (AMED)/The Japan International
Cooperation Agency (JICA), the Science and Technology Research Partner‑
ship for Sustainable Development (SATREPS), and the Ministry of Education,
Culture, Sports, Science and Technology (MEXT) of Japan as a project for Joint
Usage/Research Center.
Availability of data and materials
All data generated or analyzed during this study are included in this publishedarticle.
Authors’ contributions
RUS, RM, and KF designed this study. RUS and RM performed the experiments,
data analysis, and drafted the manuscript. HS participated in the design,
coordination, and revision of the manuscript. All authors actively contributed
to the interpretation of the findings. All authors read and approved the final
manuscript.

Umemiya‑Shirafuji et al. Parasites Vectors(2019) 12:205

Ethics approval and consent to participate
The care and use of experimental animals in this study were approved by
the Animal Care and Use Committee of Obihiro University of Agriculture andVeterinary Medicine (approval no. 28-35).

Page 11 of 12

14.

Consent for publication
Not applicable.

15.

Competing interests
The authors declare that they have no competing interests.

16.

Publisher’s NoteSpringer Nature remains neutral with regard to jurisdictional claims in pub‑lished maps and institutional affiliations.

17.

Author details
1
National Research Center for Protozoan Diseases, Obihiro University of Agri‑culture and Veterinary Medicine, Inada-Cho, Obihiro, Hokkaido 080‑8555,
Japan. 2 National Agricultural and Food Research Organization, Kannondai
3‑1‑5, Tsukuba, Ibaraki 305‑0856, Japan.

18.

Received: 9 January 2019 Accepted: 29 April 2019

20.

19.

21.
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