Profiling of proteolytic enzymes in the gut of the tick Ixodes ricinus reveals an evolutionarily conserved network of aspartic and cysteine peptidases
© Sojka et al; licensee BioMed Central Ltd. 2008
Received: 29 January 2008
Accepted: 18 March 2008
Published: 18 March 2008
Ticks are vectors for a variety of viral, bacterial and parasitic diseases in human and domestic animals. To survive and reproduce ticks feed on host blood, yet our understanding of the intestinal proteolytic machinery used to derive absorbable nutrients from the blood meal is poor. Intestinal digestive processes are limiting factors for pathogen transmission since the tick gut presents the primary site of infection. Moreover, digestive enzymes may find practical application as anti-tick vaccine targets.
Using the hard tick, Ixodes ricinus, we performed a functional activity scan of the peptidase complement in gut tissue extracts that demonstrated the presence of five types of peptidases of the cysteine and aspartic classes. We followed up with genetic screens of gut-derived cDNA to identify and clone genes encoding the cysteine peptidases cathepsins B, L and C, an asparaginyl endopeptidase (legumain), and the aspartic peptidase, cathepsin D. By RT-PCR, expression of asparaginyl endopeptidase and cathepsins B and D was restricted to gut tissue and to those developmental stages feeding on blood.
Overall, our results demonstrate the presence of a network of cysteine and aspartic peptidases that conceivably operates to digest host blood proteins in a concerted manner. Significantly, the peptidase components of this digestive network are orthologous to those described in other parasites, including nematodes and flatworms. Accordingly, the present data and those available for other tick species support the notion of an evolutionary conservation of a cysteine/aspartic peptidase system for digestion that includes ticks, but differs from that of insects relying on serine peptidases.
Ticks are important vectors of infectious agents causing diseases in human and domestic animals . The castor bean tick Ixodes ricinus transmits Lyme disease caused by Borrelia burgdorferi spirochetes and tick borne encephalitis caused by the tick-borne encephalitis virus .
Blood-feeding and -digestion are essential activities for ticks. Blood provides a rich source of proteins and nutrients for anabolic processes such as vitellogenesis and egg production . Unlike other blood-feeding arthropods, ticks are believed to digest blood intracellularly – in the endo/lysosomal vesicles of gut cells  at pH values well below the pH 6.3 – 6.5 of the gut contents [5, 6]. Digestive gut cells use both receptor-mediated and fluid-phase endocytoses to uptake the liquid blood meal from the gut lumen . Lara et al.  showed that the digestive cells of Boophilus microplus have separated endocytic pathways for two major proteins of host blood – serum albumin and hemoglobin. The requirement for receptor-mediated endocytosis might be directly linked to the detoxification of released heme groups during intracellular digestion of hemoglobin. Most of the toxic heme forms a unique type of heme aggregate ultimately accumulated inside specialized organelles called hemosomes . The virtual absence of extracellular digestive enzymes in ticks enables the gut lumen to serve as a major storage organ .
In spite of the above studies, our understanding of the molecular proteolytic machinery involved in digesting host proteins in the tick gut is still rather fragmented. Previous studies have tended to focus on individual enzymes in particular species; all however, have identified either cysteine or aspartic peptidases; e.g., a cysteine class cathepsin L in B. microplus , two forms of a cathepsin L in Haemaphysalis longicornis  and the aspartic peptidase, cathepsin D (termed longepsin) in H. longicornis . Also, cysteine-class asparaginyl endopeptidases (AE, legumains) have been characterized in I. ricinus  and H. longicornis .
The data thus far from different tick species raise the hypothesis that tick intestinal digestion relies on an evolutionarily conserved network of cysteine and aspartic peptidases characterized in other parasites, including platyhelminths [15, 16] and nematodes . It comprises mainly cysteine peptidases cathepsin B, L, C, asparaginyl endopeptidase/legumain and an aspartic peptidase cathepsin D. To address this hypothesis we focused on a defined feeding phase of a single tick species, namely partially engorged females of I. ricinus. Two-pronged profiling strategy involving biochemical assays and PCR-based cloning displayed a simultaneous expression and activity of the above listed peptidase types in the tick digestive tissue. An improved global insight increases the possibilities for practical interventions involving vaccines and offers a better understanding of vector-pathogen interactions at the primary interface, namely the tick gut.
Functional profiling of multiple peptidase activities in the gut of I. ricinus
Thus, five significant endo- and exopeptidase activities of the cysteine and aspartic classes of peptidases were profiled in the gut tissue of I. ricinus. The identified activities include (i) the CA clan (papain-type) cysteine peptidases: cathepsins B, L, and C and a Clan CD asparaginyl endopeptidase (legumain), and (ii) a Clan AA aspartic peptidase activity: cathepsin D.
Genetic screening of gut tissue identifies cDNAs encoding one aspartic and four cysteine peptidases
Degenerate PCR primers used for identification of genes encoding gut-associated peptidases of I. ricinus
Sm* forward domain
Sm* reverse domain
Hybridization screening of the gut cDNA library with radio-labeled PCR amplicons had been previously used to obtain the full-length coding sequence for IrAE . Here the same approach succeeded in identifying full length cDNA sequences for cathepsins B, L and D. The full cDNA sequence of cathepsin C was generated by overlapping PCR, 5' and 3' RACE PCR fragments.
For the sake of consistency, we adopted a nomenclature for these enzymes previously used for schistosomal peptidases [15, 18] – a nomenclature already applied to the I. ricinus asparaginyl endopeptidase (IrAE) . Thus, we designated the novel peptidases as IrCB for cathepsin B (form 1), IrCL for cathepsin L, IrCC for cathepsin C and IrCD for cathepsin D.
I. ricinus cathepsin B (IrCB)
I. ricinus cathepsin L (IrCL)
I. ricinus cathepsin C (IrCC)
I. ricinus cathepsin D (IrCD)
Differential expression of peptidases during development and in tissues
Gene specific primers used for RT-PCR expression profiling of I. ricinus peptidases
The present functional and genetic profiling in the gut of the hard tick I. ricinus has identified a number of peptidase activities and genes. The overall goal was to have a better global understanding of the component peptidases for one important tick species in a particular feeding phase, namely partially engorged I. ricinus females, in contrast to the present fragmented picture regarding individual enzymes in a variety of different tick species. Also, this broad approach offers both the possibility to compare entire digestive systems with other hematophagous parasites as well as to investigate the potential of one or more component peptidases as molecular vaccines.
Initial activity profiling of I. ricinus gut extracts using a biologically relevant protein substrate (AMC hemoglobin) indicated that hemoglobinolysis is optimal at acid pH, a finding in accordance with data presented for other tick species [5, 23, 24]. This suggested that proteolysis is mediated by peptidases belonging to the aspartic and/or cysteine peptidase classes which are known to operate optimally at acid pH . This conclusion was further supported by the sensitivity of hemoglobinolysis to class-selective peptidase inhibitors. Accordingly, we functionally scanned gut extracts for individual peptidase activities with a battery of diagnostic, small molecule substrates and inhibitors. These studies revealed the presence of four cysteine peptidase activities, cathepsins B, C, L and AE, and an aspartic peptidase activity, cathepsin D.
To identify the peptidase genes putatively responsible for the activities measured in gut extracts, we next screened gut-derived cDNA with degenerate primers designed to amplify individual peptidases. Five cysteine and one aspartic peptidases were classified: IrAE , IrCB, IrCL, IrCC and IrCD. Interestingly, from approximately 10 sequenced amplicons of each peptidase cDNA, only IrCB presented as two different isoforms. The finding is comparable with the data for hematophagous flukes, Schistosoma mansoni  and Trichobilharzia regenti , both of which have more than one cathepsin B isoforms.
Sequence comparison with human cathepsin B reveals that IrCB has the signature 'occluding loop' necessary for its exopeptidase (specifically, peptidyl dipeptidase) activity . Regarding murine cathepsin C, this enzyme was previously shown to process and activate granulocyte serine peptidases by the removal of N-terminal dipeptides . Thus, it would be of interest to test the potential competence of cathepsin C to processes cubulin-like serine peptidases inducing lysis of host blood cells . The primary structure of the IrCD precursor is homologous to longepsin from H. longicornis  with two conserved Asp-Thr-Gly (DTG) catalytic site motifs either side of the substrate binding groove, a structure not shared by the more evolutionary distinct yolk-processing tick cathepsin D  and tick heme-binding aspartic peptidase  in the eggs of B. microplus. Finally, a search through the available EST database (NCBI Blast with a limitation to tick ESTs) indicated the existence of several isoforms for cathepsins B, L, AE and cathepsin D, but only one form of cathepsin C in the whole body derived cDNA of the closely related tick species, Ixodes scapularis . Certainly, these preliminary data need to await final contig assembly and gene annotations and show the need for tick gut transcriptome projects.
With the exception of the eggs, host blood is taken up and processed to provide energy and nutrients for the transition from larva to nymph and finally, to adult male or female . By RT-PCR, enzymes under study are expressed in all the feeding developmental stages indicating their simultaneous action in digestive cells. Notably, however, IrCB and IrCD are not expressed in eggs, suggesting a function specifically associated with blood digestion. In support of this notion, the tissue-specific RT-PCR demonstrated that both peptidases are expressed solely in the gut. Likewise, IrAE is also restricted to the gut but, being also found in eggs, must have an additional function(s) not associated with the blood meal.
The combined biochemical and genetic analyses presented in this study demonstrate that I. ricinus expresses a suite of gut-associated cysteine and aspartic peptidases in order to catabolize ingested host proteins as a nutrient source. The data accord with previous results for enzyme activities in different tick species [5, 24]. The particular combination of cysteine and aspartic peptidases comprising AE, and cathepsins B, C, D and L, operating at acidic pH and localized to the gut, is remarkably similar to those found in phylogenetically distant nematodes [17, 34, 35] and platyhelminths [15, 16, 34]. Indeed, cysteine and aspartic peptidases also contribute to amino acid acquisition in protozoa such as Plasmodium [36, 37]. Therefore, and as noted by Delcroix et al.  for the platyhelminth S. mansoni, digestive systems based on cysteine and aspartic peptidases are widespread in invertebrates and stand in contrast to those systems utilizing serine peptidases (e.g., in insects and vertebrates). The present report extends this observation to include arthropods, specifically, ixodid ticks.
On an applied note, gut-associated peptidases may prove useful as vaccine targets. Other ixodid gut proteins, such as Bm86/Bm95 or Bm91, are suitable antigens for vaccination strategies (reviewed in de la Fuente and Kocan ). With this goal in mind, the detailed molecular and cellular characterization of the I. ricinus peptidases will be the subject of future reports.
I. ricinus ticks were collected by flagging in woodland localities around České Budějovice in the Czech Republic. Adult males and females were kept separately in glass vials in wet chambers with humidity of about 95% and temperature 26°C. If not stated otherwise, the females were allowed to feed naturally for 5 days on laboratory guinea pigs, carefully removed by forceps and referred to as partially engorged ticks in experiments described below. Laboratory animals were treated in accordance with the Animal Protection Law of the Czech Republic no. 246/1992 Sb.
The 7-amino-4-methylcoumarin (AMC)-conjugated bovine hemoglobin was prepared according to Partanen et al. . All peptidyl AMC-substrates were from Bachem, Abz-Lys-Pro-Ala-Glu-Phe-Nph-Ala-Leu (Abz, aminobenzoic acid; Nph, 4-nitrophenylalanine) substrate was prepared as described in Máša et al. . Peptidase inhibitors were from Bachem, Gly-Phe-diazomethyl ketone (DMK) was prepared as described in Green and Shaw . The aza-peptide Michael acceptor (CBz-Ala-Ala-(aza-Asn)-CH = CH-COOEt) further referred to as Aza-N-11a  was kindly donated by Dr. J.C. Powers of the School of Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia.
Preparation of the tick gut tissue extract
For an experiment, tissues were dissected from 10 partially engorged I. ricinus females. The gut contents were carefully removed with a special care not to disrupt the epithelium. Cleaned guts were washed in phosphate-buffered saline solution (PBS) and pooled. The gut tissue extract (150 μg protein/ml) was prepared by homogenization of the gut tissue (without contents) in 1 ml of 0.1 M Na-acetate, pH 4.5, 1% CHAPS, 2.5 mM DTT using teflon-glass homogenizer on ice. The homogenate was centrifuged for 10 min at 10000 × g, the supernatant was filtered with Micropure-0.22 Separator (Millipore) and stored at -80°C.
Quantification of hemoglobin degradation
Hemoglobinolytic activity was assayed using AMC-hemoglobin as a fluorogenic substrate . Digestion of fluorogenic AMC-hemoglobin (0.5 μg supplemented with 2 μg of bovine hemoglobin) was performed at 35°C with the gut tissue extract (20-fold diluted stock solution) in 0.1 M Na-citrate-phosphate, pH 2.5–8.0 including 2.5 mM DTT, 25 mM NaCl and 0.05% Tween 20 in a reaction mixture of 100 μl. The proteolytic fragmentation of AMC-hemoglobin results in an increase of the fluorescence intensity that was continuously monitored to determine the relative reaction rate. The fluorescence signal was measured using a GENios Plus reader (TECAN) at 360 nm excitation and 465 nm emission wavelengths. For the hemoglobinolytic assay in the presence of a peptidase inhibitor, an aliquot of the extract was preincubated (15 min at 35°C) in the same buffer pH 4.0 with 10 μM E64, 10 μM pepstatin, 1 mM Pefabloc or 1 mM EDTA.
Profiling component gut peptidases with substrates and inhibitors
Peptidase activities were identified and characterized by hydrolysis of the following fluorogenic substrates: 25 μM Z-Arg-Arg-AMC for cathepsin B , 25 μM Z-Phe-Arg-AMC for cathepsin L , 30 μM Gly-Arg-AMC for cathepsin C , 40 μM Abz-Lys-Pro-Ala-Glu-Phe-Nph-Ala-Leu for cathepsin D , and 30 μM Z-Ala-Ala-Asn-AMC for asparaginyl endopeptidase  as described previously [40, 47]. The activity measurement was performed at 35°C using an aliquot of the gut tissue extract (20 to 200-fold diluted stock solution) in 0.1 M Na-acetate, pH 4.0 including 2.5 mM DTT (for cysteine peptidases) and 25 mM NaCl (for cathepsin C). For the activity assay in the presence of peptidase inhibitors, an aliquot of the extract was pre-incubated (15 min at 35°C) in the same buffer with the inhibitor: 10 μM CA-074 for cathepsin B , 10 μM Z-Phe-Phe-DMK for cathepsin L , 1 μM Gly-Phe-DMK for cathepsin C , 10 μM pepstatin for cathepsin D  or 1 μM Aza-N-11a for asparaginyl endopeptidase . Hydrolytic activity was continuously measured after addition of substrate in a fluorescence reader GENios Plus at 320 nm excitation and 420 nm emission wavelengths (for Abz-containing substrate) or at 360 nm excitation and 465 nm emission wavelengths (for AMC-containing substrates). Assays of asparaginyl endopeptidase and cathepsin L were measured in the presence of 10 μM CA-074 to prevent confounding hydrolysis by cathepsin B [16, 47].
Tissues (gut, salivary glands, ovaries and Malpighian tubules) were dissected from partially engorged females in a wax filled Petri dish with phosphate-buffered saline (PBS) under a binocular dissection microscope. The whole body homogenates from different developmental stages were prepared by crushing the appropriate number of eggs, larvae, nymphs, males, unfed females and females removed from guinea pigs one day after attachment using mortar and pestle and repeated freezing under liquid nitrogen. For total RNA isolation, the samples were further homogenized in a micro-tube with a plastic pestle in the TRI Reagent® solution (Sigma) at 1 ml per 50–100 mg of wet tissue and processed according to the instructions provided with the TRI-reagent kit (Sigma). Isolated total RNA was stored at -80°C and further used for preparing single stranded cDNA templates using Superscript II (Invitrogen) and oligo(dT) primers, following the instructions provided by the manufacturer or for RT-PCR experiments described below.
Designing PCR oligonuclotide primers
Degenerate primers were designed from conserved domains of Schistosoma mansoni, S. japonicum, mosquito, rat and human peptidases. Protein and nucleotide sequences downloaded from the NCBI GenBank web site were used for multiple ClustalW alignments in DNAstar MegAlign software (Lasergene). The oligonucleotides are listed in Table 1.
PCR and rapid amplification of cDNA ends (5'- and 3-' RACE)
Mastercycler gradient (Eppendorf) was used to optimize PCR amplifications. Amplicons were purified, ligated into vector plasmid pCR 4-TOPO using the TOPO TA® Cloning Kit (Invitrogen) and transformed into E. coli TOP 10 cells (Invitrogen). Clones containing ligated PCR products were sequenced using an automated sequencer model ABI Prism 3130 XL and the BigDye® Terminator sequencing kit (Applied Biosystems) with appropriate sequencing primers. Sequence data were compared by blastn  against the NCBI GenBank database records. To obtain complete cDNA sequences, 3' RACE PCR was performed using a modified protocol for SMART™ cDNA Library Construction Kit (Clontech, BD Biosciences) described previously . The N-terminal sequences including signal peptides and the 5' untranslated regions were determined using the Invitrogen 5'RACE system and instructions provided by the manufacturer.
cDNA library construction and screening
The protocol for construction of the I. ricinus gut-derived cDNA library with SMART™ cDNA Library Construction Kit (Clontech, BD Biosciences) and the Gigapack® III Gold packaging extract (Stratagene) as well as the method of cDNA library screening by [P32]dATP radio-labeled gene-specific probes have been described previously .
The primary sequences used for phylogenetic analysis comprised the conserved domains spanning across the mature enzyme sequences without pro-domains. Sequences were obtained from the MEROPS database  and aligned in the program ClustalX 1.81 . The alignment was manually checked using the BioEdit program . Tree reconstruction employed the Neighbor Joining (NJ) method  in the program MEGA 2.1 . Nodal supports were calculated with 1000 replications.
Gene specific PCR primer pairs (listed in Table 2) were designed for each peptidase type with DNAstar PrimerSelect software (Lasergene). Two-step RT-PCR was performed using total RNA templates (prepared as described above; 50 ng/μl final concentration) and the Enhanced Avian HS RT-PCR Kit (Sigma) according to the protocol provided by the manufacturer. Amplification of the ferritin mRNA, previously shown to be presented in all tick tissues and expressed independently of feeding , was used as a loading control.
expressed sequence tag
National Center for Biotechnology Information
polymerase chain reaction
rapid amplification of cDNA ends
reverse transcription polymerase chain reaction.
This work was supported by the grant No. 206/06/0865 to P.K. and M.M., PhD program No. 524/03/H133 to D.S. and Z.F. from the Grant Agency of the Czech Republic and Research Centre No. LC06009. M.H. was supported by the grant KJB400550516 from the Grant Agency of the Academy of Sciences of the Czech Republic and C.R.C. by the Sandler Family Supporting Foundation. Research at the Institute of Parasitology, BC ASCR, Institute of Organic Chemistry and Biochemistry ASCR and Faculty of Science, USB is covered by research plans Nos. Z60220518, Z40550506 and MSMT 6007665801, respectively. We thank James C. Powers, School of Chemistry and Biochemistry, Georgia Institute of Technology, GA, for the gift of Aza-N-11a.
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