DeMar Taylor, Graduate School of Life and Environmental Sciences, University of Tsukuba, Tennodai 1-1-1, Tsukuba, Ibaraki, 305-8572, Japan. Tel./fax: 81 29 853 4806; e-mail: email@example.com
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.
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).
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.
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.
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
% Identity (similarity) of each domain
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.
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.
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.
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.
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.
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.
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.
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.