- Open Access
The midgut of Aedes albopictus females expresses active trypsin-like serine peptidases
© Saboia-Vahia et al.; licensee BioMed Central Ltd. 2014
- Received: 26 August 2013
- Accepted: 6 May 2014
- Published: 30 May 2014
Aedes albopictus is widely distributed across tropical and sub-tropical regions and is associated with the transmission of several arboviruses. Although this species is increasingly relevant to public health due its ability to successfully colonize both urban and rural habitats, favoring the dispersion of viral infections, little is known about its biochemical traits, with all assumptions made based on studies of A. aegypti. In previous studies we characterized the peptidase profile of pre-imaginal stages of A. albopictus and we reported the first proteomic analysis of the midgut from sugar-fed females of this insect species.
In the present work, we further analyzed the peptidase expression in the midgut of sugar-fed females using 1DE-substrate gel zymography, two-dimensional electrophoresis (2DE), mass spectrometry (MS), and protein identification based on similarity.
The combination of zymography, in solution assays using fluorescent substrates and 2DE-MS/MS allowed us to identify the active serine peptidase “fingerprint” in the midgut of A. albopictus females. Zymographic analysis revealed a proteolytic profile composed of at least 13 bands ranging from ~25 to 250 kDa, which were identified as trypsin-like serine peptidases by using specific inhibitors of this class of enzymes. Concomitant use of the fluorogenic substrate Z-Phe-Arg-AMC and trypsin-like serine protease inhibitors corroborated the zymographic findings. Our proteomic approach allowed the identification of two different trypsin-like serine peptidases and one chymotrypsin in protein spots of the alkaline region in 2DE map of the A. albopictus female midgut. Identification of these protein coding genes was achieved by similarity to the A. aegypti genome sequences using Mascot and OMSSA search engines.
These results allowed us to detect, identify and characterize the expression of active trypsin-like serine peptidases in the midgut of sugar-fed A. albopictus females. In addition, proteomic analysis allowed us to confidently assign the expression of two trypsin genes and one chymotrypsin gene to the midgut of this mosquito. These results contribute to the gene annotation in this species of unknown genome and represent a small but important step toward the protein-level functional and localization assignment of trypsin-like serine peptidase genes in the Aedes genus.
- Aedes albopictus
- Two-dimensional electrophoresis
- Mass spectrometry
Aedes (Stegomyia) albopictus (Skuse) has a wide geographic distribution, covering all tropical and subtropical regions of the world, and is a vector for the viruses responsible for yellow fever and dengue . The World Health Organization estimates that more than 50–100 million cases of these two diseases can occur per year throughout the world [2–4]. In Brazil, A. albopictus has been reported in 21 states and 1,502 municipalities . In recent years, the relevance of this species to public health has increased because it is able to successfully colonize both urban and rural habitats, favoring the dispersion and interchange of the virus from one area to another, and thereby enabling the emergence of new areas of disease in small and large cities [6, 7].
The hydrolysis of proteins to amino acid residues by proteolytic enzymes is an important step in food digestion, protein turnover and proteostasis in eukaryotes [8, 9]. Proteolytic enzymes are divided into endopeptidases and exopeptidases. Endopeptidases are relatively small molecules (~25-30 kDa) that can pass through peritrophic membrane pores and endoperitrophic spaces in insects, where they cleave large protein complexes. Exopeptidases are large enzymes (>100 kDa) that are usually linked to the plasma membrane of the midgut epithelium and hydrolyze the ends of small proteins and peptides (N-terminus or C-terminus) . Among endopeptidases, trypsin-like and chymotrypsin-like serine peptidases are the most important enzymes for most insects, except for some species of coleoptera and hemiptera [11–16].
Serine peptidases are divided into families and subfamilies. The subfamily S1 consists of trypsins, chymotrypsins and elastases, and some serine collagenases were also recently included. The catalytic triad of serine peptidases is typically characterized by serine, histidine and aspartic acid residues [17, 18]. This triad hydrolyzes peptide bonds at the carboxylic ends of basic amino acids, with a 2-10-fold preference for Arg over Lys [19–21]. The A. aegypti genome contains 369 genes coding for serine peptidases, among which 66 are putative trypsins , but only 5 (three trypsins and two chymotrypsins) are well characterized in the midgut of females of this insect [13–16, 23]. The expansion of trypsin-like serine peptidase genes in mosquitoes has been shown to coincide with the development of the hematophagous trait . Trypsin-like serine peptidases in these insects play pivotal roles in oogenesis, immunity, metamorphosis, modulation of embryonic development and nutrition [25, 26]. These enzymes are mostly located in the insect midgut so that they can provide energy and essential amino acids for development [20, 27]. Furthermore, secretion of trypsin-like serine peptidases into the lumen of the midgut is involved in defense against pathogens [28, 29]. However, in the insect vector Anopheles gambiae, trypsin-like enzymes are exploited by pathogens such as Plasmodium sp. to activate their own peptidases, thus allowing the parasite to cross the peritrophic membrane and continue its developmental life cycle .
The plasticity exhibited by trypsin-like serine peptidases enables these enzymes to modulate various biological processes in insect vectors. Because of this characteristic, serine peptidases have been proposed as potential targets for insect control approaches. The biochemical characterization of these enzymes may thus support the development of new control strategies, enabling their appropriate use as targets and suggesting ways to interfere with the production of these enzymes or with the metabolic pathways in which they participate [31, 32].
In previous studies, we characterized the peptidase profile of the pre-imaginal stages of A. albopictus and reported the first proteomic analysis of the midgut of sugar-fed females of this insect species . In the present work, we further analyze the peptidase expression in the midgut of sugar-fed females using two-dimensional electrophoresis (2DE), mass spectrometry, 1DE-substrate gel, and data mining. This multi-methodological approach allowed us to identify the active serine peptidase “fingerprint” in the midgut of A. albopictus females.
To prepare the phenylmethylsulfonylfluoride (PMSF) stock solution, 250 mM of the reagent was diluted in isopropanol. Nα-tosyl-L-lysine chloromethyl ketone hydrochloride (TLCK) and N-p-tosyl-L-phenylalanine chloromethyl ketone (TPCK) were dissolved in methanol, both at 100 mM. Stock solutions of 1,10-phenanthroline (200 mM) and pepstatin A (1 mg/ml) were prepared in ethanol, and trans-epoxysuccinyl L-leucylamido-(4-guanidino) butane (E-64, 10 mM) was prepared in water. Stock and working solutions were maintained at -20°C. All chemicals were purchased from Sigma Chem. Co. (USA), unless otherwise specified.
Insect rearing and gut dissection
Aedes albopictus specimens reared in a closed colony (Laboratório de Transmissores de Hematozoários, Instituto Oswaldo Cruz, FIOCRUZ, Rio de Janeiro) were kindly provided by Dr. Nildimar A. Honorio. Mosquitoes were maintained on a 10% sucrose diet at 25 ± 1°C, with a relative humidity of 60 ± 10% and a light:dark photoperiod of 14:10 h. For each experiment, 50 female adults (2–5 days old) were cold-anesthetized on ice and decapitated. Midgut dissection was performed as previously described .
Zymography and peptidase inhibition assays
For proteolytic assessment, midguts were washed twice with PBS pH 7.2 and lysed as previously described . Briefly, midguts were lysed with a cell disruption motor drive and pestle in a tube containing 10% glycerol, 0.6% Triton X-100, 100 mM Tris–HCl pH 6.8 and 150 mM NaCl . The protein concentration of the resulting extracts was determined using the Pierce 660 nm Protein assay (Thermo Scientific). For protein separation, 30 μg of protein were loaded in 12% polyacrylamide gels copolymerized with 0.1% porcine gelatin as the substrate. Electrophoreses were performed at 4°C with a constant voltage of 110 V. Peptidase activity was detected as previously reported  with few modifications. The gels were incubated at 37°C for 2, 4, 6, 12 or 24 h in reaction buffer containing 100 mM sodium acetate (at pH 3.5 or 5.5) or 100 mM Tris–HCl (pH 7.5 or 10.0). Substrate degradation was visualized as clear bands after staining the gels with 0.2% Coomassie blue R-250 in methanol/acetic acid (40:10) and destaining in 10% acetic acid. The relative molecular masses of the activity bands were estimated by comparison with the mobility of a commercial molecular mass standard (PageRuler™ Protein Ladder, Fermentas). To determine the classes of peptidases detected by zymography, peptidase inhibition assays were conducted. Midgut homogenates were pre-incubated (before electrophoresis) for 30 min at 4°C with one of the following peptidase inhibitors: 20 μM E-64, 5 mM PMSF, 100 μM TLCK, 100 μM TPCK, 10 μM pepstatin-A or 10 mM 1,10-phenanthroline. After electrophoresis, inhibitors were added to the reaction buffer at the same concentration, and the peptidases were resolved as described above. The results were derived from three independent experiments carried out in triplicate.
In-solution enzymatic assays
The effects of pH and peptidase inhibitors on the proteolytic activities of midgut homogenates were also evaluated by in-solution assays using the fluorogenic substrate Z-Phe-Arg-AMC. For both assays, 100 μM of substrate was used. The reactions were initiated by diluting 10 μg of protein from the midgut in 100 mM sodium acetate (at pH 3.5 or 5.5) or 100 mM Tris–HCl (pH 7.5 or 10.0) for pH evaluation or 100 mM Tris–HCl pH 7.5 with or without 100 μM TLCK, 100 μM TPCK, 20 μM E-64 or 5 mM PMSF. The fluorescence intensity was measured by spectrophotofluorometry every 5 min for a 60 min period (SpectraMax Gemini XPS, Molecular Devices, CA) using excitation and emission wavelengths of 380 and 460 nm, respectively. As blank, the substrate (100 μM) was diluted in the reaction buffer [100 mM sodium acetate (at pH 3.5 or 5.5) or 100 mM Tris–HCl (pH 7.5 or 10.0)]. The value of the blank was automatically discounted by the fluorometer software (SoftMax®Pro, Molecular Devices, CA) when the data were acquired. All assays were performed at 37°C. The results were derived from three independent experiments performed in triplicate.
2DE electrophoresis and protein identification
Protein extraction, separation and identification were performed as previously described . Briefly, 50 pooled midguts were mechanically disrupted with a pestle and a motor drive in a tube containing lysis buffer (9 M urea, 4% CHAPS, 65 mM dithiothreitol, DTT, and 1% ampholytes, pH 3–10, with 5 mM PMSF and a protease inhibitor cocktail). Proteins were precipitated and resuspended in 9 M urea, 4% CHAPS, 65 mM DTT and 1% ampholytes, pH 3–10 NL. The protein concentration was determined using the 2D Quant Kit (GE Healthcare), and 100 μg were subjected to isoelectric focusing over a nonlinear pH gradient of 3–10 on a 7 cm strip (GE Healthcare) on an Ettan IPGphor 3 instrument (GE Healthcare). The focusing parameters were set as previously described . After reduction and alkylation, proteins were separated vertically across 12% SDS-PAGE gels using standard Tris/glycine/SDS buffer. Gels were stained with colloidal Coomassie Brilliant Blue G-250, documented using a GS-800™ calibrated imaging densitometer (Bio-Rad) and analyzed using PDQuest™ software (Bio-Rad). Experimental pI and M r were calibrated using a select set of reliable identification landmarks distributed throughout the entire gel.
Protein digestion, peptide extraction and analysis by mass spectrometry were performed as previously described . Raw MS files were converted to MGF format using Mass Matrix MS Data File converter V. 3.9 http://www.massmatrix.net/mm-cgi/downloads.py. To maximize search sensitivity and peptidase identification, the data were searched using OMSSA  within the Proteomatic platform 1.2.1 http://www.proteomatic.org/download.html and the Mascot MS/MS ion search engine (http://www.matrixscience.com/search_form_select.html, Matrix Science, Oxford, UK, free online version). As the genome sequences of A. albopictus are not available, mass spectra were searched in OMSSA against an A. aegypti database downloaded (May 2013) from UniRef100 http://www.uniprot.org/, and in MASCOT against the non-redundant database of the National Center for Biotechnology (NCBI). Searches were performed with one missed cleavage, with carbamidomethylation of cysteine residues as a fixed modification, methionine oxidation as a variable modification and mass tolerances of 2.0 and 0.8 Da in OMSSA and 10 ppm and 0.4 Da in MASCOT for precursor and fragment ions, respectively. The Vectorbase database (http://www.vectorbase.org) was used to search for sequence information about the identified peptidases.
The A. albopictus female midgut exhibits a complex profile of active peptidases
Influence of the pH on the proteolytic profile of the A. albopictus midgut
Proteolytic activities in the A. albopictus female midgut mainly arise from trypsin-like serine peptidases
Trypsin-like serine peptidases and chymotrypsin were identified in the 2DE map of the A. albopictus female midgut
Aedes albopictus midgut peptidases automatically identified using the Mascot software
NCBI Accesion No.
VectorBase DB No.
Matching pep./ Pep. identified by MS/MS
Error ± ppm
Chymotrypsin, putative [Aedes aegypti]
Trypsin-alpha, putative [Aedes aegypti]
Trypsin [Aedes aegypti]
Although the A. aegypti genome has been reported to code for 380 trypsin-like serine peptidases, constituting one of the largest gene families in mosquitoes , the interrogation of the Vectorbase database using the words “trypsin” or “chymotrypsin” reveals only 80 coding genes for trypsin and 5 coding genes for chymotrypsin in the A. aegypti genome, among which are the genes identified here. As these peptidases are members of large gene families, it is difficult to ascertain which protein is expressed and active in a specific life cycle stage, in a specific tissue, under a specific condition. Therefore, we cannot rule out the possibility that other proteases, such as chymotrypsin, could be expressed in their active form in the midgut of blood feeding females. Our study shows the potential value of proteomic approaches combined with zymographic analysis for the identification and localization assignment of specific gene products.
The results obtained in this work allowed us to detect, identify and characterize the expression of active trypsin-like serine peptidases in the midgut of sugar-fed A. albopictus females. In addition, proteomic analysis allowed us to confidently assign the expression of two trypsin genes and one chymotrypsin gene to the midgut of this mosquito. These results contribute to the gene annotation in this species of an unknown genome and represent a small but important step toward the protein-level functional assignment of trypsin-like serine peptidase genes in the Aedes genus. As peptidases exert crucial roles during host-pathogen interactions and the midgut is the main setting for these interactions in blood feeding vector mosquitoes, the mapping and identification of the constitutively expressed peptidase profile in this tissue may allow for the comparison of the regulation of such enzymes in infected insects and/or mosquitoes fed on blood. Such approaches may produce valuable information on the roles of peptidases during host-pathogen interactions.
This work was supported by FAPEMIG (J.B.J Edital Universal Process No. APQ-01070-12), CNPq (J.B.J. PQ Process No. 308679/2012-1; L.S.V. Process No. 142964/2009-3), FAPERJ (C.B. Process E-26/110.594/2012 and CNE E-26/102.775/2012), FIOCRUIZ-IOC and CAPES. We thank Prof. Dr. Nildimar Honorio (Laboratório de Transmissores de Hematozoários of the Instituto Oswaldo Cruz) for kindly providing the insects.
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