SEARCH

SEARCH BY CITATION

Keywords:

  • EcR;
  • Ornithodoros moubata;
  • ecdysteroid;
  • vitellogenesis;
  • tick

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Molecular mechanisms of ecdysteroid regulation in development and reproduction have been thoroughly investigated in Diptera and Lepidoptera, but few studies report the molecular actions of ecdysteroids in hemimetabolous insects and more primitive arthropods. Ecdysteroids appear to be the main hormones regulating development and vitellogenesis in ticks. An ecdysteroid receptor that showed high homology with EcRs of other arthropods was isolated from Ornithodoros moubata (OmEcRA). OmEcR expression patterns coincided with ecdysteroid titres in the haemolymph during moulting and vitellogenesis and differed between mated and virgin females. Therefore, OmEcR appears to mediate the regulation of moulting and vitellogenesis by ecdysteroids in O. moubata females as seen in other arthropods.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Development and reproduction as well as many other processes are regulated by ecdysteroids in invertebrates. Ecdysteroids are released into the haemolymph and stimulate various tissues with specifically timed responses. Gene regulatory mechanisms of ecdysteroids have been thoroughly characterized in higher insect orders such as Diptera and Lepidoptera. Two nuclear receptors, the ecdysteroid receptor (EcR) and ultraspiracle (USP), an orthologue of the retinoid X receptor (RXR), are required for gene regulation by ecdysteroids in insects (Yao et al., 1992, 1993; Thomas et al., 1993; Hall & Thummel, 1998). However, EcR requires RXR as a heterodimer partner to function as an EcR in crustaceans (Durica & Hopkins, 1996; Durica et al., 2002; Wu et al., 2004) and ticks (Mao & Kaufman, 1998; Palmer et al., 2002). The heterodimer of EcR and USP or RXR accepts ecdysteroid and the ecdysteroid/receptor complex binds to ecdysteroid response elements (EcREs) of target genes to regulate gene transcription. The precursor egg yolk protein vitellogenin (Vg) has been extensively studied as a target gene activated by ecdysteroids in mosquitoes. Female Aedes aegypti release ecdysteroids into the haemolymph and synthesize Vg in the fat body after blood feeding (Raikhel & Dhadialla, 1992). The Vg gene contains the regulatory EcRE sequence in the 5′ upstream region and Vg gene transcription is directly regulated by the ecdysteroid/EcR/USP complex (Wang et al., 2000a; Kokoza et al., 2001; Martin et al., 2001; Raikhel et al., 2002). This complex appears in a tissue- and stage-specific manner in A. aegypti (Kapitskaya et al., 1996; Martin et al., 2001) and interacts with early ecdysteroid genes such as broad complex, E74 and E75 to regulate Vg transcription (Raikhel et al., 2002).

EcR genes have been isolated from several species of arthropods (Henrich & Brown, 1995; Bonneton et al., 2003; King-Jones & Thummel, 2005) and have a common structure. The N terminus of a nuclear receptor is a variable region because of alternate splicing (A/B domain) and this region determines the transactivation function of the receptor. The second region is the C domain or DNA binding domain (DBD) that binds the receptor to the promoter region of target genes. The DBD is followed by a hinge area (D domain), and the fourth region is the E domain or ligand binding domain (LBD) for ligand reception. In addition, Dipteran and Lepidopteran insects have a variable F domain at the carboxyl terminus, the function of which is unknown (Koelle et al., 1991; Cho et al., 1995; Fujiwara et al., 1995; Swevers et al., 1995; reviewed in Bonneton et al., 2003). Insects produce several isoforms of EcR with different A/B domains and the number of isoforms differs between insect species (Koelle et al., 1991; Talbot et al., 1993; Swevers et al., 1995; Jindra et al., 1996). Each isoform is expressed in a tissue- and stage-specific manner and responds to different ecdysteroid doses and transcriptional regulation (Talbot et al., 1993; Wang et al., 2000b, 2002; Sullivan & Thummel, 2003) indicating different functions for each isoform. Only recently have molecular mechanisms of ecdysteroid gene regulation been reported in hemimetabolous insects (Saleh et al., 1998; Cruz et al., 2006) and more primitive arthropods (Chung et al., 1998; Kim et al., 2005). These studies indicate further research on ecdysteroid regulation of gene transcription in lower arthropods is needed to better understand the evolution and mode of action of ecdysteroids.

Ticks are ectoparasitic arthropods that require a blood meal for development and reproduction and in the process of obtaining a blood meal they transmit various pathogens. Despite their importance as vectors of disease, little is known of the mechanisms regulating development and reproduction in ticks. Ecdysteroid regulation of vitellogenesis has been investigated in various species of ticks (reviewed by Taylor & Chinzei, 2002). Ecdysteroids are released into the haemolymph after blood feeding and appear to regulate development, ecdysis and vitellogenesis (Diel et al., 1982; Germond et al., 1982; Stauffer & Conneat, 1990; Palmer et al., 2002; Taylor & Chinzei, 2002; Rees, 2004; Ogihara et al., 2007). Chinzei et al. (1991) and Taylor et al. (1991) reported that juvenile hormone (JH) does not stimulate vitellogenesis in O. moubata and O. parkeri. In addition Neese et al. (2000) reported the absence of JH and JH precursors in both hard and soft ticks. In addition, injection of ecdysteroids into unfed females induced Vg synthesis (Taylor et al., 1997). These results are consistent with the hypothesis that ecdysteroids play a primary role in the regulation of development and reproduction in ticks and studies on the ecdysteroid regulatory mechanisms of ticks can provide an excellent system for elucidating these mechanisms in primitive arthropods.

The hard tick Amblyomma americanum has three EcRs (AamEcRA1, A2 and A3) and two RXRs (Guo et al., 1997, 1998). The three AamEcRs are expressed with high titres of ecdysteroids during moulting in larval and nymphal stages (Palmer et al., 2002). In addition, AamEcR and AamRXR form a heterodimer and accept ecdysteroids for subsequent binding to the Drosophila hsp27 EcRE (Guo et al., 1998; Palmer et al., 2002). In the soft tick O. moubata, mated females synthesis large amounts of Vg and produce mature eggs but virgin females do not synthesize adequate amounts of Vg to produce mature eggs. These differences in Vg synthesis and egg maturation between mated and virgin females appear to be regulated by ecdysteroids because only mated females show high titres of ecdysteroids in the haemolymph after engorgement (Ogihara et al., 2007). However, the molecular mechanisms involving the EcR receptors have not yet been reported in these ticks. Therefore, we isolated EcR from O. moubata and determined the OmEcR expression patterns from engorgement of the last nymphal (female) instar through the reproductive cycle of adult females to clarify the roles of ecdysteroids in the regulation of reproduction of soft ticks.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Isolation and characteristics of OmEcR gene

Based on partial sequences of OmEcR (Taylor et al., 2000) the complete open reading frame (ORF) of O. moubata EcR was obtained by 5′ and 3′ rapid amplification of cDNA ends (RACE) (Fig. 1). 5′ RACE of OmEcR showed several bands, indicating O. moubata may have additional isoforms as shown in other arthropods. However, we were unable to confirm the presence of other isoforms in O. moubata. The identified sequence was 2140 nucleotides with 568 deduced amino acid residues (AB191193). The sequence contained the A/B region, a DBD, hinge region, LBD but no carboxy terminal F domain as also seen in the hard tick A. americanum, crustaceans and lower insects. The DBD is located at residues 197–273 and contains eight cysteines to form a C2C2 zinc finger for DNA binding. The LBD is located at residues 346–567.

image

Figure 1. Nucleotide and deduced amino acid sequences of OmEcRA. The sequence was determined by 5′ RACE and 3′ RACE and translated into amino acids. The numbers indicate nucleotide length. A single underline indicates the DNA binding domain and a double underline indicates the ligand binding domain. Arrowheads indicate the cysteine residues of the zinc fingers. Asterisk (*) indicates the translation stop codon. Start codon is indicated with a single underline.

Download figure to PowerPoint

The homology of each domain was determined with the EMBOSS needle program (global pairwise alignment) and a comparison of the identities and similarities of OmEcR with other arthropods is shown in Table 1. The A/B isoform specific domain of OmEcR showed the highest similarity with AamEcRAs and relatively higher homology with EcRA isoforms when compared with M. sexta and A. aegypti. In insects, the EcRA isoform contains a specific A box upstream of the DBD. The specific A box is also present in OmEcRA but shows divergence from insect EcRAs (Fig. 2) indicating tick EcRs may be more primitive EcRA isoforms. The DBD and LBD showed high homology to EcRs of other arthropods. The highest identities were found in the DBD with an identity of more than 85% compared with other arthropods. OmEcR LBD showed identities of approximately 60% to other EcR LBDs.

Table 1.  Comparison of the functional domains of OmEcR with other EcRs
EcR genes % Identity (similarity) of each domain
A/BCDE
AamEcRA1Tick56 (65)   
AamEcRA2 44 (55)100 (100)62 (69)99 (99)
AamEcRA3 20 (22)   
UpEcRCrab26 (38) 96 (100)46 (55)67 (81)
LmEcROrthoptera31 (39) 97 (100)49 (63)68 (82)
BgEcRAOrthoptera30 (37) 99 (100)54 (70)66 (82)
TmEcRColeoptera21 (28) 99 (100)47 (58)65 (80)
PmEcRAHymenoptera24 (32) 94 (99)53 (64)68 (82)
MsEcRALepidoptera28 (38) 88 (97)36 (47)57 (70)
MsEcRB1 20 (30)   
AaEcRADiptera29 (36) 86 (96)36 (45)61 (73)
AaEcRB1 22 (28)   
image

Figure 2. Multiple alignment of OmEcRA and other arthropod EcRs. The specific A box in the A/B domain, functional P and D boxes in the DNA binding domain, T and A boxes in the hinge area, and the AF2 box in the ligand binding domain are enclosed with solid line boxes. DBD start and end as well as LBD start and end are indicated. AamEcR is from Amblyomma americanum, BgEcRA from Blattella germanica, AaEcRA from Aedes aegypti and BgEcRB1 from Bombyx mori.

Download figure to PowerPoint

Multiple alignments of the EcRs were performed for a more detailed analysis (Fig. 2). The DBD contains functional sequences called P and D boxes for specific DNA recognition and T and A boxes in the hinge area to affect DNA recognition. All residues of the P box are the same as other EcRs but the residues of the D box of arthropods separate into two groups. OmEcR has the same amino acid sequence as AamEcR, BgEcR, UpEcR in the D box and these EcRs form a heterodimer with RXR. Homologies of the T and A boxes in the hinge area are also highly conserved in OmEcR. The LBD functions for heterodimerization and ligand binding. The 3′ area of the LBD sequence is called the AF2 region and effects the formation of the heterodimer. OmEcR has residues similar to the AF2 region of other arthropods indicating the heterodimerization of OmEcR is similar to other EcRs. Therefore, OmEcR is likely able to form a heterodimer with RXR and bind to the EcRE to regulate gene transcription as reported in other arthropods.

Phylogenetic trees for EcR DBD and LBD are shown in Fig. 3(A) and Fig. 3(B), respectively. The DBD of OmEcR grouped with the clade that includes the hard tick A. americanum, crustaceans and lower insects, whereas Lepidopteran and Dipteran EcR DBDs clustered in a different clade. On the other hand, the LBD of ticks and a crab were included in a branch separate from the insects and other arthropods. The phylogenetic tree as well as homology in the LBD suggest the ligand responses and heterodimerization of ticks and insects differ.

image

Figure 3. Phylogenetic tree of EcR genes constructed by the neighbour-joining method with the Clustal X program. Numbers at the branches indicate the bootstrap values with 1000 replicates. Human LXRα is the liver X receptor of Homo sapiens and is used as the outgroup. The sequences used were Manduca sexta, Bombyx mori, Aedes aegypti, Drosophila melanogastar, Uca pugliator, Locusta migratoria, Pheidole megacephara, Amblyomma americanum, Tenebrio molitor and Blattella germanica. (See Experimental procedures for gene accession numbers.)

Download figure to PowerPoint

Analysis of the timing of EcR expression

EcR is necessary to mediate ecdysteroid functions. Therefore, OmEcR gene expression was analysed in last instar nymphs and throughout the reproductive cycle of female O. moubata ticks. The total expression of OmEcR was determined with primers designed from the DBD and expression levels of a specific isoform (OmEcRA) were determined with primers designed from the A/B isoform specific domain to further confirm the absence of more than one isoform.

The last instar nymphs of O. moubata moult at approximately 10 days after engorgement, so total OmEcR and OmEcRA expression in the last instar nymphs was analysed from 0 to 9 days after engorgement (Fig. 4). The expression level of total OmEcR increased immediately after engorgement, decreased 1 day after engorgement, then subsequently increased from 2 to 5 days after engorgement and again just before moulting. EcR regulation appears to start soon after engorgement and play an important role in the regulation of moulting. The expression patterns of OmEcRA showed a pattern almost identical to total OmEcR expression. In addition, analysis of absolute gene copy numbers showed no significant differences (t-test, P > 0.05) between total OmEcR and OmEcRA in last instar nymphs at the highest level of expression 5 days after engorgement. Therefore, OmEcRA appears to be the main OmEcR isoform expressed during moulting.

image

Figure 4. Analysis of OmEcR gene expression in last instar nymphs of O. moubata. (A) Total OmEcR expression was determined with primers constructed from the DNA binding domain, and (B) OmEcRA expression determined with primers constructed from the A/B domain of OmEcRA. Total RNA was isolated from last instar nymphs everyday after engorgement until moulting. Unfed values of total OmEcR and OmEcRA expression were standardized as 1 unit. cDNA from nymphs each day after engorgement were used for real-time PCR. The values presented are mean ± SE. Vertical arrow indicates the time of ecdysis.

Download figure to PowerPoint

OmEcR and OmEcRA expression in newly emerged females was compared between unfed mated (Fig. 5A,B) and virgin females (Fig. 5C,D). In mated females, total OmEcR and OmEcRA expression was observed immediately after emergence and 5–15 days after emergence. Virgin females did not show total OmEcR and OmEcRA expression immediately after emergence, but higher expression from 1 to 10 days after emergence. OmEcR expression differs in unfed virgin and mated females. Mated females show repression of OmEcR expression at the moult from the last instar nymph to adult female with subsequent activation 5 days after emergence, whereas virgin females show higher levels of OmEcR expression during the period before feeding.

image

Figure 5. Analysis of OmEcR gene expression in newly emerged adult females of O. moubata. (A) Total OmEcR and (B) OmEcRA expression in mated females. Total RNA was isolated from emerged adults approximately 6–12 h after emergence to allow males to mate with them. (C) Total OmEcR and (D) OmEcRA expression in virgin females. Total RNA was isolated from emerged adults as above except males were not added to the individual containers. Values for total OmEcR or OmEcRA expression of unfed mated females was standardized as 1 unit. The values presented are mean ± SE. Vertical arrow indicates the time of ecdysis.

Download figure to PowerPoint

Shortly after engorgement, females of O. moubata begin vitellogenesis and oviposit eggs approximately 10 days after feeding. Both mated and virgin females were fed and the expression levels of total OmEcR and OmEcRA analysed throughout the reproductive cycle (Fig. 6). Engorged mated females showed a rapid increase in total OmEcR expression from soon after blood feeding and maintained a high level of expression until 7 days after engorgement (Fig. 6A). OmEcRA expression from 0 to 6 days after engorgement was similar to total OmEcR expression (Fig. 6B). Total OmEcR and OmEcRA expression levels in virgin females similarly increased soon after engorgement to much higher levels than seen in mated females but rapidly decreased to lower levels 2 days after engorgement (Fig. 6C,D). In addition, analysis of absolute gene copy numbers showed no significant differences (t-test, P > 0.05) between total OmEcR and OmEcRA in mated and virgin females at the highest level of expression 1 day after engorgement. OmEcRA appears to be the main isoform for regulation of vitellogenesis and mating appears to synchronize the expression of the EcR receptor in mated females.

image

Figure 6. Analysis of OmEcR gene expression in mated and virgin females of O. moubata. (A) Total OmEcR and (B) OmEcRA expression in mated females after engorgement. (C) Total OmEcR and (D) OmEcRA expression in virgin females after engorgement. Total RNA was isolated from mated and virgin females immediately and everyday for 20 days after engorgement. Unfed values of total OmEcR or OmEcRA expression were standardized as 1 unit. The values presented are mean ± SE. Vertical arrows indicate the start of Vg synthesis and oviposition in mated females.

Download figure to PowerPoint

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Molecular studies on the functions of ecdysteroids have focused on Dipteran and Lepidopteran species and have only recently included hemimetabolous insects and other arthropods. These recent studies suggest the mechanisms and receptors regulating ecdysteroid functions differ between recent and primitive arthropods. Ticks provide an excellent system in a more primitive group to determine the evolutionary development of ecdysteroid regulatory mechanisms because ecdysteroids play an important part in the development and reproduction of ticks. In this study, we isolated EcR from the soft tick O. moubata (OmEcRA) and showed that expression in engorged last instar nymphs and unfed and fed mated and virgin adult females coincide with the ecdysteroid titres that regulate ecdysis and vitellogenin synthesis.

The DBD of OmEcRA showed over 85% identity with other arthropods and 100% identity with AamEcRs (Table 1). Guo et al. (1998) and Palmer et al. (2002) showed that AamEcRs can bind to the EcRE of Drosophila and activate gene transcription. The 100% homology between the DBDs of OmEcRA and AamEcRs suggest OmEcRA may also function in the regulation of gene transcription. The LBD of OmEcRA showed approximately 60% identity to EcRs of other arthropods. The phylogenetic tree also shows the LBD of OmEcRA is different from insect EcRs (Fig. 3). Ecdysteroids have several chemical conformations and the EcR isoforms show different responses to the various ecdysteroids (Hiruma et al., 1997; Wang et al., 2000a). Divergence of EcR LBDs among arthropods suggests differences in the partner for heterodimerization and ligand response.

Analysis of OmEcR expression in last instar nymphs revealed that the highest expression occurred 5 days after engorgement and just before ecdysis (Fig. 4). These expression peaks coincided with increases in ecdysteroid titres in the haemolymph (Ogihara et al., 2007). Ecdysteroids function in larval tissue degeneration and adult tissue construction in holometabolous insects such as D. melanogaster (Kozlova & Thummel, 2002). Several genes that are related to programmed cell death are regulated by the ecdysteroid and EcR/USP complex (Cakouros et al., 2002; Lee et al., 2002; Yin & Thummel, 2004). Therefore, OmEcR expression several days before ecdysis in the last instar nymphal stage suggests ecdysteroids function to prepare the female tissues for development in addition to the regulation of ecdysis. OmEcR expression in newly emerged females is clearly different between mated and virgin females indicating mating has an effect on EcR expression, but further research is needed to clarify these effects.

After engorgement females enter a vitellogenic cycle in which Vg synthesis begins approximately 3 days after engorgement and Vg is incorporated into the oocytes. Egg laying starts approximately 10 days after engorgement. Ecdysteroid titres increase in mated females from soon after engorgement to 6 days after engorgement (Ogihara et al, 2007). OmEcR expression was observed in mated females from soon after engorgement to 8 days after engorgement (Fig. 6A,B). Therefore, OmEcR expression and high titres of ecdysteroids appear together in mated females before and during Vg synthesis. Virgin females synthesize Vg at much lower levels than mated females and eggs do not mature and are not oviposited. This absence of increased Vg synthesis and egg development appears to be due to the ecdysteroid titres remaining at low levels (Ogihara et al., 2007). OmEcR expression was approximately two times higher in virgin females than mated females (Fig. 6). Unliganded EcR/USP heterodimer has been shown to repress the transcription of target genes (Schubiger et al., 2005). Therefore, the low ecdysteroid titre and the high OmEcR expression may result in a lower Vg titre and lack of egg maturation in virgin females.

Numerous EcR sequences have been identified from various species. Diptera and Lepidoptera have been reported to have EcRA and EcRB isoforms (Koelle et al., 1991; Talbot et al., 1993; Fujiwara et al., 1995; Swevers et al., 1995; Jindra et al., 1996; Bonneton et al., 2003). Only EcRB isoform has been reported from Leptinotarsa decemlineata (Ogura et al., 2005) and only EcRA from Locusta migratoria (Saleh et al., 1998) and Blattella germanica (Cruz et al., 2006). The hard tick A. americamun has three isoforms but all three isoforms are the EcRA type (Guo et al., 1997). Several reports show that isoforms are expressed with specific patterns and have different functions. EcRA and EcRB isoforms are expressed at different times in the last instar during moulting of M. sexta, D. melanogaster and A. aegypti (Talbot et al., 1993; Truman et al., 1994; Jindra et al., 1996; Margam et al., 2006). These results have contributed to the hypothesis that EcRB1 regulates tissue proliferation and remodelling during metamorphosis and EcRA regulates final differentiation during metamorphosis (Jindra et al., 1996). In this study, OmEcRA was expressed at almost the same level and with similar patterns as total OmEcR indicating O. moubata may express only OmEcRA. Expression of BgEcRA in B. germanica is similar to total BgEcR expression and knockdown of the BgEcRA isoform by RNAi shows the BgEcRA isoform regulates events in both moulting and vitellogenesis (Cruz et al., 2006). Tick development goes through several nymphal stages similar to hemimetabolic insects, so the EcRA isoform may be the primary receptor of ecdysteroids for the regulation of moulting and vitellogenesis in ticks. To further elucidate EcR functions in ticks, clarification that other isoforms do not exist and analysis of their expression if they do exist are necessary. Although, the high homology between AamEcRA and OmEcRA indicate OmEcR can function to bind ecdysteroids and activate gene expression, identification of RXR and confirmation of EcR and RXR heterodimerization in O. moubata are in progress. Further experiments to clarify the stimulation of Vg transcription and ecdysis by ecdysteroids are also needed.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Ticks

Ornithodoros moubata ticks used in this study were from a laboratory colony maintained in incubators at 30 °C ± 1 °C, 70% ± 10% RH and total darkness. Ticks were fed on rabbits (Oryctolagus cunniculus) as described by Chinzei et al. (1983). Virgin females were obtained by keeping fifth (last) instar nymphs isolated in individual wells of 24 well sample plates before and after emergence. Whereas mated females were obtained by adding males to the individual wells of the last instar nymphs just before moulting. The experimental protocols and animal care were approved by the Ethical Committee for Animal Studies at the University of Tsukuba.

Determination of OmEcR nucleotide sequence by rapid amplification of cDNA ends (RACE)

Total RNA was extracted with ISOGEN as described by the manufacturer (Nippon Gene, Tokyo, Japan) from fed females after removal of the gut. For 3′ RACE, first strand cDNA was synthesized with a SuperScript Preamplification System (Gibco BRL, Langley, OK, U.S.A.). Total RNA (5 µg) was mixed with NotI d(T)18 primers and cDNA synthesis was performed as described by the manufacturer. Nested polymerase chain reaction (PCR) for 3′ RACE was carried out with Ex Taq polymerase (TaKaRa Bio, Otsu, Shiga, Japan) using OmEcRC1 Forward derived from the C domain of O. moubata EcR sequence isolated by Taylor et al. (2000) as a sense primer and NotI d(T)18 oligonucleotides as an anti-sense primer for first PCR, and OmEcRC2 Forward as a sense primer and NotI d(T)18 oligonucleotides as an anti-sense primer for a second PCR. The sequences of all primers used in this study are shown in Table 2. The first PCR was performed for 3 min at 94 °C followed by 35 cycles at 94 °C for 30 s, 60 °C for 30 s, 72 °C for 3 min and then held at 72 °C for 7 min. The second PCR was performed for 3 min at 94 °C followed by 25 cycles at 94 °C for 30 s, 60 °C for 30 s, 72 °C for 2 min and then held at 72 °C for 7 min.

Table 2.  Primers used in this study
 NameSequence
RACE adapter5′ RACE Outer primer5′-GCUGAUGGCGAUGAAUGAACACUGCGUUUGCUGGCUUUGAUGAAA-3′
RACE adaptor for sequence5′ RACE Outer primer5′-GCTGATGGCGATGAATGAACACTG-3′
5′ RACE Inner primer5′-CGCGGATCCGAACACTGCGTTTGCTGGCTTTGATG-3′
NotI d(T)185′-d (AACTGGAAGAATTCGCGGCCGCAGGAAT)-3′
Gene-specific primer for sequenceOmEcRA11 Forward5′-TGACTTTCACGATCGTCTGC-3′
OmEcRA11 Reverse5′-GCATGGGTCCCTCTGT-3′
OmEcRA12 Forward5′-GAGTTTAACGTCTCCATCC-3′
OmEcRB1 Forward5′-GAGATGAGTCCAGCAGTG-3′
OmEcRC1 Forward5′-GTACGGCAATAACTGCGACA-3′
OmEcRC1 Reverse5′-CATTCCAACGCTGAGGCAC-3′
OmEcRC2 Forward5′-GGCAATAACTGCGACATCGAC-3′
OmEcRD1 Forward5′-GGAGGACCTCATCAACAAGC-3′
OmEcRD1 Reverse5′-GGTCGGTCTTTGTCCTTCTG-3′
OmEcRE1 Forward5′-GGGATTCGACACGCTCTTAC-3′
OmEcRE1 Reverse5′-TCTCGGTGATGTGCT-3′
OmEcRE2 Forward5′-ACCCGGCAAGAACTACTTTG-3′
OmEcRE2 Reverse5′-CACATCTCGGCGTTCATGTT-3′
OmEcRE3 Reverse5′-TCACTGTTGGATGTCCCAT-3′
pGEM T-Easy vectorT75′-TAATACGACTCACTATAGGG-3′
SP65′-TATTTAGGTGACACTATAG-3′
Real-time PCREC forward5′-GCACTGACCTGTGAAGGGTGT-3′
EC labelled reverse5′-gtacggTCGATGTCGCAGTTATTGCCG5AC-3′
ES forward5′-CGAACCACCAGTACCGTTCC-3′
ES labelled reverse5′-caacgCGCAGGTACAGGAGGCCG5TG-3′
Actin forward5′-GGGAATGGAATCCTGCGGTA-3′
Actin labelled reverse5′-gacctgGGACAGTGTTGGCGTACAGG6C-3′

5′ RACE was performed using a First Choice RLM-RACE Kit (Applied Biosystems, Foster City, CA, U.S.A.). A RNA Adaptor oligonucleotide was ligated to the total RNA and cDNA synthesis was performed as described by the manufacturer. Nested PCR for 5′ RLM-RACE was performed with Ex Taq (TaKaRa Bio) using 5′ RACE Outer Primer as a sense primer and OmEcRE2 Reverse as an anti-sense primer for first PCR, and 5′ RACE inner primer as a sense primer and OmEcRE1 Reverse as an anti-sense primer for the second PCR. The first PCR was performed under the same conditions as the first PCR for 3′ RACE. The second PCR was performed for 3 min at 94 °C followed by 25 cycles at 94 °C for 30 s, 63 °C for 1 min, and 72 °C for 2 min.

Purified PCR products from both 3′ and 5′ RACE were sequenced with a 377 ABI automated DNA sequencer or Beckman CEQ 2000 DNA analysis system. Primers were OmEcRC2 Forward, OmEcRE1 Forward, OmEcRE2 Reverse, OmEcRE2 Forward, and NotI d(T)18 for 3′ end sequencing, and were Inner primer, OmEcRD1 Reverse and OmEcRE1 Reverse for 5′ end sequencing.

To determine the complete ORF of OmEcR, PCR was performed using Platinum Taq DNA Polymerase (Invitrogen, San Diego, CA, U.S.A.) with OmEcRA11 Forward as a sense primer and OmEcRE3 Reverse as an anti-sense primer under the same conditions as the first PCR for 3′ RACE. PCR products were subcloned into a pGEM®-T Easy Vector (Promega, Madison, WI, U.S.A.). The sequences of plasmids were confirmed by DNA sequencing using T7, SP6, OmEcRA11 Reverse, OmEcRA12 Forward, OmEcRB1 Forward, OmEcRC1 Reverse, OmEcRD1 Forward, OmEcRE1 Reverse and OmEcRE2 Forward primers.

Phylogenetic analysis and pairwise alignment

Homology of each EcR domain was compared with several EcR genes from arthropods with global pairwise alignment by the EMBOSS needle program. Multiple alignments were peformed by the CLUSTAL X program (Thompson et al., 1997). Phylogenetic trees were profiled by the neighbour-joining method (Saitou & Nei, 1987) using the sequences of the DBDs and LBDs. Bootstrap values were assessed with 1000 replicates. The amino acid sequences of EcR were obtained from GenBank. GenBank accession numbers of the sequences used are as follows: O. moubata EcR (AB191193), A. aegypti EcRs (EcRA: AY345989, EcRB1: U02021), A. americanum EcRs (EcRA1: AF020187, EcRA2: AF020188, EcRA3: AF020186), B. germanica (AM039690), Uca pugilator (AF034086), D. melanogaster EcR (M74078), L. migratoria EcR (AF049136), M. sexta EcR (EcRA: U49246, EcRB1: U19812), Pheidole megacephala EcRA (AB194765) and Tenebrio molitor EcR (Y11533). The liver X receptor (LXR) of vertebrates is a nuclear receptor and belongs to the same group as EcR, therefore, LXRα of Homo sapiens (U22662) was used as an outgroup.

Expression analysis of EcR in O. moubata

For analysis of OmEcR gene expression in O. moubata, last instar nymphs and females were used. Last instar nymphs moult at nearly 10 days after engorgement and most become females. OmEcR expression in last instar nymphs was determined with unfed and 0–9 days after engorgement. OmEcR expression in newly emerged mated and virgin females were determined from soon after emergence (D0) to every day for 5 days after emergence, and on 10 and 15 days after emergence. To clarify OmEcR expression during vitellogenesis, unfed virgin and mated females were fed and OmEcR expression determined everyday from 0 to 20 days after engorgement. Total RNA was isolated with TRIzol reagent (Invitrogen) as described by the manufacturer. Total RNA (2 µg) was treated with DNase I Amp grade (Invitrogen) as described by the manufacturer and used for cDNA synthesis with the SuperScript III First Strand Synthesis System (Invitrogen). The reverse transcription reaction was performed at 50 °C for 50 min followed by heating at 85 °C for 5 min with Oligo d(T)20 primers according to the manufacturer and used as template for mRNA quantification.

Determination of total OmEcR and OmEcRA expression was performed by real-time PCR. The actin gene (Horigane et al., 2007) was used as an internal control for O. moubata EcR expression. Expression levels of total OmEcR and OmEcRA in the unfed ticks of last instar nymphs, and mated and virgin females were used as the standard for each stage. Real-time PCR was performed with PCR Super Mix-UDG (Invitrogen) as described by the manufacturer. All primers were LUX fluorogenic primers designed with the D-LUXTM program (Invitrogen). Primers (EC) were designed from sequences of the DNA binding domain to determine total OmEcR expression and primers (ES) from the isoform-specific domain (A/B domain) to determine OmEcRA-specific expression. All primers were made by D-LUXTM Designer (Invitrogen). The primers for EC and ES were labelled with FAM while the primers for the actin gene were labelled with JOE (Table 2). Final concentration of each primer was 0.5 µM/well for actin and 0.25 µM/well for EC and ES. PCR reactions were performed for 2 min at 50 °C, 2 min at 95 °C, and 45 cycles at 95 °C for 15 s and 60 °C for 30 s. Triplicate reactions were run in an ABI Prism 7900HT (Applied Biosystems). The amplification efficiencies of all primers were 95–103% so the comparative CT method was used to analyse the whole body samples. Raw data were analysed by SDS 2.0 (Applied Biosystems) and Excel (Microsoft) by the comparative CT method according to the manufacturer's instructions (Applied Biosystems). To compare absolute gene copy numbers of total OmEcR and OmEcRA, the absolute standard curve method was also performed for nymphs 5 days after engorgement and for fed virgin and mated females 1 day after engorgement. An absolute standard curve was made with a 10-fold dilution of a known number of plasmids that contained the EcR fragment. The plasmids were constructed by subcloning the PCR products into a pGEM®-T Easy Vector (Promega) and the sequences confirmed by DNA sequencing. Total OmEcR and OmEcRA expression was determined by absolute standard curve methods and values compared with a t-test.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

This research was supported in part by Grants-in-Aid for Scientific Research (16580035 and 18580050 to DT) from the Ministry of Education, Sciences and Culture, Japan.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  • Bonneton, F., Zelus, D., Iwema, T., Robinson-Rechavi, M. and Laudet, V. (2003) Rapid divergence of the ecdysone receptor in Diptera and Lepidoptera suggests coevolution between ECR and USP-RXR. Mol Biol Evol 20: 541553.
  • Cakouros, D., Daish, T., Martin, D., Baehrecke, E.H. and Kumar, S. (2002) Ecdysone-induced expression of the caspase DRONC during hormone-dependent programmed cell death in Drosophila is regulated by Broad-Complex. J Cell Biol 157: 985995.
  • Chinzei, Y., Chino, H. and Takahashi, K. (1983) Purification and properties of vitellogenin and vitellin from a tick, Ornithodoros moubata. J Comp Physiol B 152: 1321.
  • Chinzei, Y., Taylor, D. and Ando, K. (1991) Effects of juvenile hormone and its analogues on vitellogenin synthesis and ovarian development in the soft tick, Ornithodoros moubata (Acari: Argasidae). J Med Entomol 28: 506513.
  • Cho, W.L., Kapitskaya, M.Z. and Raikhel, A.S. (1995) Mosquito ecdysteroid receptor: analysis of the cDNA and expression during vitellogenesis. Insect Biochem Mol Biol 25: 1927.
  • Chung, A.C., Durica, D.S., Clifton, S.W., Roe, B.A. and Hopkins, P.M. (1998) Cloning of crustacean ecdysteroid receptor and retinoid-X receptor gene homologs and elevation of retinoid-X receptor mRNA by retinoic acid. Mol Cell Endocrinol 139: 209227.
  • Cruz, J., Mane-Padros, D., Belles, X. and Martin, D. (2006) Functions of the ecdysone receptor isoform-A in the hemimetabolous insect Blattella germanica revealed by systemic RNAi in vivo. Dev Biol 297: 158171.
  • Diel, P.A., Germond, J.E. and Morici, M. (1982) Correlations between ecdysteroid titer and integument structure in nymphs of the tick, Amblyomma hebraeum Koch (Acarina: Ixodidae). Rev Suisse Zool 89: 859868.
  • Durica, D.S. and Hopkins, P.M. (1996) Expression of the genes encoding the ecdysteroid and retinoid receptors in regenerating limb tissues from the fiddler crab, Uca pugilator. Gene 171: 237241.
  • Durica, D.S., Wu, X., Anilkumar, G., Hopkins, P.M. and Chung, A.C. (2002) Characterization of crab EcR and RXR homologs and expression during limb regeneration and oocyte maturation. Mol Cell Endocrinol 189: 5976.
  • Fujiwara, H., Jindra, M., Newitt, R., Palli, S. R., Hiruma, K. and Riddiford, L.M. (1995) Cloning of an ecdysone receptor homolog from Manduca sexta and the developmental profile of its mRNA in wings. Insect Biochem Mol Biol 25: 845856.
  • Germond, J.E., Diehl, P.A. and Morici, M. (1982) Correlations between integument structure and ecdysteroid titers in fifth-stage nymphs of the tick, Ornithodoros moubata (Murray, 1877; sensu Walton, 1962). Gen Comp Endocrinol 46: 255266.
  • Guo, X., Harmon, M.A., Laudet, V., Mangelsdorf, D.J. and Palmer, M.J. (1997) Isolation of a functional ecdysteroid receptor homologue from the ixodid tick Amblyomma americanum (L.). Insect Biochem Mol Biol 27: 945962.
  • Guo, X., Xu, Q., Harmon, M.A., Jin, X., Laudet, V., Mangelsdorf, D.J., et al . (1998) Isolation of two functional retinoid X receptor subtypes from the Ixodid tick, Amblyomma americanum (L.). Mol Cell Endocrinol 139: 4560.
  • Hall, B.L. and Thummel, C.S. (1998) The RXR homolog ultraspiracle is an essential component of the Drosophila ecdysone receptor. Development 125: 47094717.
  • Henrich, V.C. and Brown, N.E. (1995) Insect nuclear receptors: a developmental and comparative perspective. Insect Biochem Mol Biol 25: 881897.
  • Hiruma, K., Bocking, D., Lafont, R. and Riddiford, L.M. (1997) Action of different ecdysteroids on the regulation of mRNAs for the ecdysone receptor, MHR3, dopa decarboxylase, and a larval cuticle protein in the larval epidermis of the tobacco hornworm, Manduca sexta. Gen Comp Endocrinol 107: 8497.
  • Horigane, M., Ogihara, K., Nakajima, Y., Honda, H. and Taylor, D. (2007) Identification and expression analysis of an actin gene from the soft tick, Ornithodoros moubata (Acari: Argasidae). Arch Insect Biochem Physiol 64: 186199.
  • Jindra, M., Malone, F., Hiruma, K. and Riddiford, L.M. (1996) Developmental profiles and ecdysteroid regulation of the mRNAs for two ecdysone receptor isoforms in the epidermis and wings of the tobacco hornworm, Manduca sexta. Dev Biol 180: 258272.
  • Kapitskaya, M., Wang, S., Cress, D. E., Dhadialla, T.S. and Raikhel, A.S. (1996) The mosquito ultraspiracle homologue, a partner of ecdysteroid receptor heterodimer: cloning and characterization of isoforms expressed during vitellogenesis. Mol Cell Endocrinol 121: 119132.
  • Kim, H.-W., Chang, E.S. and Mykles, D.L. (2005) Three calpains and ecdysone receptor in the land crab Gecarcinus lateralis: sequences, expression and effects of elevated ecdysteroid induced by eyestalk ablation. J Exp Biol 208: 31773197.
  • King-Jones, K. and Thummel, C.S. (2005) Nuclear receptors-a perspective from Drosophila. Nat Rev Genet 6: 311323.
  • Koelle, M.R., Talbot, W.S., Segraves, W.A., Bender, M.T., Cherbas, P. and Hogness, D.S. (1991) The Drosophila EcR gene encodes an ecdysone receptor, a new member of the steroid receptor superfamily. Cell 67: 5977.
  • Kokoza, V.A., Martin, D., Mienaltowski, M.J., Ahmed, A., Morton, C.M. and Raikhel, A.S. (2001) Transcriptional regulation of the mosquito vitellogenin gene via a blood meal-triggered cascade. Gene 274: 4765.
  • Kozlova, T. and Thummel, C.S. (2002) Spatial patterns of ecdysteroid receptor activation during the onset of Drosophila metamorphosis. Development 129: 17391750.
  • Lee, C.Y., Simon, C.R., Woodard, C.T. and Baehrecke, E.H. (2002) Genetic mechanism for the stage-and tissue-specific regulation of steroid triggered programmed cell death in Drosophila. Dev Biol 252: 138148.
  • Mao, H. and Kaufman, W.R. (1998) DNA binding properties of the ecdysteroid receptor in the salivary gland of the female ixodid tick, Amblyomma hebraeum. Insect Biochem Mol Biol 28: 947957.
  • Margam, V.M., Gelman, D.B. and Palli, S.R. (2006) Ecdysteroid titers and developmental expression of ecdysteroid-regulated genes during metamorphosis of the yellow fever mosquito, Aedes aegypti (Diptera: Culicidae). J Insect Physiol 52: 558568.
  • Martin, D., Wang, S.F. and Raikhel, A.S. (2001) The vitellogenin gene of the mosquito Aedes aegypti is a direct target of ecdysteroid receptor. Mol Cell Endocrinol 173: 7586.
  • Neese, P.A., Sonenshine, D.E., Kallapur, V.L., Apperson, C.S. and Roe, R.M. (2000) Absence of insect juvenile hormones in the American dog tick, Dermacentor variabilis (Say) (Acari: Ixodidae), and in Ornithodoros parkeri Cooley (Acari: Argasidae). J Insect Physiol 46: 477490.
  • Ogihara, K., Horigane, M., Nakajima, Y., Moribayashi, A. and Taylor, D. (2007) Ecdysteroid hormone titer and its relationship to vitellogenesis in the soft tick, Ornithodoros moubata (Acari: Argasidae). Gen Comp Endocrinol 150: 371380.
  • Ogura, T., Minakuchi, C., Nakagawa, Y., Smagghe, G. and Miyagawa, H. (2005) Molecular cloning, expression analysis and functional confirmation of ecdysone receptor and ultraspiracle from the Colorado potato beetle Leptinotarsa decemlineata. FEBS J 272: 41144128.
  • Palmer, M.J., Warren, J.T., Jin, X., Guo, X. and Gilbert, L.I. (2002) Developmental profiles of ecdysteroids, ecdysteroid receptor mRNAs and DNA binding properties of ecdysteroid receptors in the Ixodid tick Amblyomma americanum (L.). Insect Biochem Mol Biol 32: 465476.
  • Raikhel, A.S. and Dhadialla, T.S. (1992) Accumulation of yolk proteins in insect oocytes. Annu Rev Entomol 37: 217251.
  • Raikhel, A.S., Kokoza, V.A., Zhu, J., Martin, D., Wang, S.F., Li, C., et al . (2002) Molecular biology of mosquito vitellogenesis: from basic studies to genetic engineering of antipathogen immunity. Insect Biochem Mol Biol 32: 12751286.
  • Rees, H.H. (2004) Hormonal control of tick development and reproduction. Parasitology 129: S127143.
  • Saitou, N. and Nei, M. (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406425.
  • Saleh, D.S., Zhang, J., Wyatt, G.R. and Walker, V.K. (1998) Cloning and characterization of an ecdysone receptor cDNA from Locusta migratoria. Mol Cell Endocrinol 143: 9199.
  • Schubiger, M., Carre, C., Antoniewski, C. and Truman, J.W. (2005) Ligand-dependent de-repression via EcR/USP acts as a gate to coordinate the differentiation of sensory neurons in the Drosophila wing. Development 132: 52395248.
  • Stauffer A. and Conneat, J.-L. (1990) Anteroposterior gradient during nyphal-adult moulting cycle of the tropical bond tick, Amblyomma variegatum (Acarina: Ixodidae). Roux's Arch Dev Biol 198: 309321.
  • Sullivan, A.A. and Thummel, C.S. (2003) Temporal profiles of nuclear receptor gene expression reveal coordinate transcriptional responses during Drosophila development. Mol Endocrinol 17: 21252137.
  • Swevers, L., Drevet, J.R., Lunke, M.D. and Iatrou, K. (1995) The silkmoth homolog of the Drosophila ecdysone receptor (B1 isoform): cloning and analysis of expression during follicular cell differentiation. Insect Biochem Mol Biol 25: 857866.
  • Talbot, W.S., Swyryd, E.A. and Hogness, D.S. (1993) Drosophila tissues with different metamorphic responses to ecdysone express different ecdysone receptor isoforms. Cell 73: 13231237.
  • Taylor, D. and Chinzei, Y. (2002) Vitellogenesis in Ticks, Chapter 6. In Reproductive Biology of Invertebrates Volume XII – Recent Progress in Vitellogenesis (Raikhel, A.S. and Sappington, T.W., eds), pp. 175199. Science Publishers Inc., Enfield, NH.
  • Taylor, D., Chinzei, Y. and Ando, K. (1991) Vitellogenin synthesis, processing and hormonal regulation in the tick, Ornithodoros parkeri (Acari: Argasidae). Insect Biochem 21: 723733.
  • Taylor, D., Moribayashi, A., Agui, N., Shono, T. and Chinzei, Y. (1997) Hormonal regulation of vitellogenesis in the soft tick, Ornithodoros moubata. In Advances in Comparative Endocrinology (Kawashima, S. and Kikuyama, S., eds), pp. 213220. Monduzzi Editore, Bologna, Italy.
  • Taylor, D., Nakajima, Y. and Chinzei, Y. (2000) Ecdysteroids and vitellogenesis in the soft tick Ornithodoros moubata (Acari: Argasidae). In Proceedings of the 3rd International Conference on Ticks and Tick-borne Pathogens: Into the 21st Century (Kazimirova, M., Labuda, M. and Nuttall, P.A., eds), pp. 223227. Institute of Zoology, Slovak Academy of Sciences, Bratislava, Slovakia.
  • Thomas, H.E., Stunnenberg, H.G. and Stewart, A.F. (1993) Heterodimerization of the Drosophila ecdysone receptor with retinoid X receptor and ultraspiracle. Nature 362: 471475.
  • Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F. and Higgins, D.G. (1997) The ClustalX windows interface: flexible strategies for multiple sequence and alignment aided by quality analysis tools. Nucleic Acids Res 24: 48764882.
  • Truman, J.W., Talbot, W.S., Fahrbach, S.E. and Hogness, D.S. (1994) Ecdysone receptor expression in the CNS correlates with stage-specific responses to ecdysteroids during Drosophila and Manduca development. Development 120: 219234.
  • Wang, S.F., Ayer, S., Segraves, W.A., Williams, D.R. and Raikhel, A.S. (2000a) Molecular determinants of differential ligand sensitivities of insect ecdysteroid receptors. Mol Cell Biol 20: 38703879.
  • Wang, S.F., Li, C., Zhu, J., Miura, K., Miksicek, R.J. and Raikhel, A.S. (2000b) Differential expression and regulation by 20-hydroxyecdysone of mosquito ultraspiracle isoforms. Dev Biol 218: 99113.
  • Wang, S.F., Li, C., Sun, G., Zhu, J. and Raikhel, A.S. (2002) Differential expression and regulation by 20-hydroxyecdysone of mosquito ecdysteroid receptor isoforms A and B. Mol Cell Endocrinol 196: 2942.
  • Wu, X., Hopkins, P.M., Palli, S.R. and Durica, D.S. (2004) Crustacean retinoid-X receptor isoforms: distinctive DNA binding and receptor–receptor interaction with a cognate ecdysteroid receptor. Mol Cell Endocrinol 218: 2138.
  • Yao, T.P., Segraves, W.A., Oro, A.E., McKeown, M. and Evans, R.M. (1992) Drosophila ultraspiracle modulates ecdysone receptor function via heterodimer formation. Cell 71: 6372.
  • Yao, T.P., Forman, B.M., Jiang, Z., Cherbas, L., Chen, J.D., McKeown, M., et al . (1993) Functional ecdysone receptor is the product of EcR and Ultraspiracle genes. Nature 366: 476479.
  • Yin, V.P. and Thummel, C.S. (2004) A balance between the diap1 death inhibitor and reaper and hid death inducers controls steroid-triggered cell death in Drosophila. Proc Natl Acad Sci USA 101: 80228027.