Background

Over 350 million people are at risk of suffering from leishmaniasis and over 2 million new cases are reported each year, which makes it one of the most neglected diseases globally [1, 2]. The sand-fly Phlebotomus papatasi (Scopoli) is the vector of the flagellate protozoan Leishmania major that causes zoonotic cutaneous leishmaniasis [3–5] in Asia and Africa [1, 6, 7]. Various species of burrowing rodents are reservoir hosts of L. major and sand flies are closely associated with the rodent burrows making vector control by direct insecticidal application difficult to achieve [6, 8–10]. Prevention of Leishmania spp. transmission to humans is largely reliant on vector control to avoid insect bites by use of screens, insecticide impregnated bed nets or curtains, establishment of barrier zones by insecticide applications, or environmental modification [2, 9, 11, 12]. Use of synthetic sand fly pheromone as an attractant was reported to improve effectiveness of pesticide application for control of the new world sand fly Lutzomyia longipalpis in chicken sheds [13]. Attempts to control Phlebotomus papatasi by various types of insecticide application has had questionable effectiveness, particularly in the harsh environments reported during combat operations by the U.S. military in Iraq [14, 15], although there was no evidence of significant insecticide resistance.

Insecticide resistance is an increasing problem in control of insect vectors of disease and there have been reports of possible resistance in sand fly populations to various insecticides [2, 11, 16]. Pesticide effectiveness for knockdown of Phlebotomine sand flies in Morocco was found to be variable for different fly populations, reflective of past history of high pesticide use in some areas for malaria control, although all of the sand flies tested as susceptible to the pesticides by bioassay, suggesting possible genetic changes that might lead to development of resistance [17]. Preliminary evidence for an altered acetylcholinesterase (E.C. 3.1.1.7, AChE) was reported in the sand fly Phlebotomus argentipes from Sri Lanka [18], and from Phlebotomus papatasi in Khartoum State, Sudan [19], each possibly the result of widespread use of insecticides in the Anti-Malarial Campaign. Coutinho-Abreu et al. [20] reported that the AChE gene of Lutzomyia longipalpis (Lutz & Neiva) appears to be structurally similar to other insect species in which point mutations result in an altered enzyme insensitive to organophosphate or carbamate inhibitors. Organophosphate pesticides function as quasi-irreversible inhibitors of AChE, resulting in failure of the central nervous system and death of the insect. Point mutations within the sequence encoding AChE resulting in production of an altered, insensitive enzyme have been reported to be a major mechanism of organophosphate resistance in insects [21–26]. Determination of the nucleotide sequence encoding AChE of P. papatasi should enable rapid identification of mutations potentially associated with resistance, and the development of rapid molecular tests to screen P. papatasi populations for the presence of specific mutations. Confirmation of AChE mutations resulting in production of an insensitive AChE is expected to facilitate the development of rapid molecular tests for pesticide resistance in sand flies. This report describes the identification of complementary DNA for AChE1 in P. papatasi and the biochemical characterization of recombinant P. papatasi AChE1 produced using a baculovirus expression system.

Methods

Sand flies

Sand flies used in this study were from a P. papatasi colony maintained at the USDA-ARS, Knipling-Bushland U.S. Livestock Insects Research Laboratory in Kerrville, Texas. The colony was established using pupae from an Israeli strain of P. papatasi maintained for 30 years and never exposed to pesticides at the Division of Entomology, Walter Reed Army Institute of Research (Silver Spring, Maryland). The colony is therefore considered to be generally susceptible to insecticides and baseline data on pesticide susceptibility by bioassay of colony sand flies is being collected for publication (A. Li, personal communication). Larvae were fed with a sand fly larval diet, a mixture of fermented rabbit feces and rabbit food [27]. Both larvae and adult flies were maintained at 26±2°C and a relative humidity of 85±2%. Males were fed with 30% sucrose water and females were fed with defibrinated cattle blood using an in vitro membrane feeding system. Larvae and newly emerged unfed adult male flies were collected and quick-frozen by immersion in liquid nitrogen, and stored at −70°C until use.

RNA preparation

Total RNA was prepared from frozen whole larvae or unfed adults by grinding in a glass tissue homogenizer containing Tri-Reagent (Sigma Chemical, St. Louis, MO) or by use of a MagMAX™-96 RNA Isolation Kit (Life Technologies, Carlsbad, CA) according to the manufacturers’ instructions.

Oligonucleotide synthesis

Synthetic oligodeoxynucleotides were purchased from Sigma-Genosys (The Woodlands, TX).

cDNA synthesis

Complementary DNA (cDNA) was synthesized by reverse transcription from template RNA using the SuperScript™ Choice System for cDNA Synthesis (Life Technologies) according to the manufacturer’s instructions. Oligodeoxynucleotide primers were oligo(dT18V) or gene-specific primers based on known sequence.

Polymerase chain reaction (PCR)

We utilized automatic hot-start PCR using AmpliTaq Gold® DNA polymerase (Applied Biosystems, Foster City, CA) or Advantage® 2 HF DNA Polymerase (BD Biosciences Clontech, Mountain View, CA) according to the manufacturers’ instructions.

Agarose gel electrophoresis

DNA preparations generated from PCR, RT-PCR, or bacterial plasmids were analyzed by agarose gel electrophoresis in tris-borate-EDTA buffer by standard techniques [28]. DNA bands were visualized by staining with GelStar® Nucleic Acid Gel Stain (Cambrex, Rockland, MD) and photographed on a Kodak GelLogic 440 Imaging system (Eastman Kodak Co., Rochester, NY) with uv illumination.

Rapid Amplification of cDNA Ends (RACE)

5’- and 3’-RACE was performed using the SMARTer™ RACE cDNA Amplification Kit (BD Biosciences Clontech, Palo Alto, CA).

DNA cloning

Double-stranded cDNA was inserted into bacterial plasmid DNA using the TOPO TA Cloning® Kit for Sequencing (Invitrogen, Carlsbad, CA), transformed into One Shot® TOP10 chemically competent Escherichia coli (Invitrogen) for amplification, screened for appropriate inserts by PCR, and sequenced by normal sequencing procedures (see below).

DNA sequencing

For sequencing, PCR amplification reactions were scaled up to 50 μl and purified using ExoSAP-IT® (Affymetrix, Santa Clara, CA). The sequencing reaction using BigDye® terminator (Applied Biosystems) and precipitation of the products with ethanol were performed according to the manufacturer’s instructions. Sequencing products were analyzed on a PRISM 3130xl Genetic Analyzer (Applied Biosystems).

Analysis of amino acid or nucleotide sequences

Sequence analysis to design oligonucleotide primers used for PCR or sequencing utilized OLIGO® Primer Analysis Software, Version 5.0 (1996, National Biosciences Inc, Plymouth, MN). Sequence chromatograms and sequence assembly utilized ChromasPro ver. 1.32 (Technelysium Pty Ltd., available at http://technelysium.com.au/?page_id=27). Clustal W (http://www.ebi.ac.uk/Tools/msa/clustalw2/) was used to perform multiple sequence alignments [29]. BLAST search of GenBank (http://blast.ncbi.nlm.nih.gov/Blast.cgi), [30] and on-line protein sequence analyses were done using ExPASy Proteomics tools.

PpAChE expression construct

Double stranded cDNA containing the complete PpAChE1 coding sequence [GenBank: JQ922267] was obtained by PCR amplification of cDNA using PpAce18F-273U23 and PpAce18F-2561 L20 (Table 1) using an annealing temperature of 62.5°C and extension time of 3 min. at 72°C. The cDNA was cloned into pBlueBac4.5/V5-His TOPO® (Applied Biosystems/Life Technologies) and sequenced to verify proper construction.

Table 1 Oligodeoxynucleotide primers for AChE of P. papatasi
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Baculovirus expression of recombinant PpAChE

PpAChE1 clones were constructed, sequenced, and expressed as baculovirus expression clones in Sf21 insect cells in sf900 III serum-free medium (Life Technologies/Gibco) as previously described [31, 32]. Serum-free cell culture baculovirus lysates were collected and centrifuged to remove cell debris and supernatants were used for biochemical characterization of recombinant PpAChE1.

Biochemical characterization of rPpAChE1

AChE activity of baculovirus culture supernatant was measured by a modified Ellman’s method as described previously [33] except that the reaction was monitored every minute for 30 min to establish initial velocity. For inhibition studies, rPpAChE was preincubated with an appropriate concentration of the inhibitor at 23°C for 10 min in 100 μl in the microtiter plate without substrate or DTNB [(5’.5’-dithiobis-(2-nitrobenzoic acid)] and AChE activity was monitored for 10 min after initiation of the reaction by addition of 100 μl substrate and DTNB. Cholinesterase inhibitors included the AChE-specific inhibitors eserine and BW284c51, butyrylcholinesterase-specific inhibitors ethopropazine and iso-OMPA, and oxidized organophosphates, paraoxon and malaoxon. All chemicals were of reagent grade obtained from Sigma Chemical Co. Data were analyzed and plotted using GraphPad Prism ver. 5.0 (GraphPad Prism, Inc., La Jolla, CA).

Results

Beginning with a presumptive acetylcholinesterase partial sequence (111 nucleotides) of Phlebotomus perniciosus [GenBank: AJ865843], we constructed oligodeoxynucleotide primers for 5’-, and 3’-RACE to obtain the complete PpAChE cDNA sequence [GenBank: JQ922267] (Additional file 1). Both strands were sequenced for the entire presumptive coding sequence of the cDNA at least three times. Oligodeoxynucleotide primers used for sequencing or 3’-/5’-RACE are listed in Table 1. The sequence obtained for the presumptive PpAChE1 cDNA of P.  papatasi was comprised of 2309 nucleotides organized as a 71-nucleotide 5’-untranslated region, an open reading frame (ORF) of 2130 nucleotides, and a 105-nucleotide 3’-untranslated region. BLAST homology search [30] of GenBank (11,299,630 sequences) found significant nucleotide identity (73-83%) to the AChE mRNA sequences of a mosquito, Culex tritaeniorhynchus [GenBank: AB122152] and for the new world sand fly L. longipalpis (partial codes, 1188 nucleotides, [GenBank: DQ898276]) respectively, as well as to other arthropod AChEs.

The P. papatasi AChE cDNA ORF encodes a 710-amino acid protein [GenBank: AFP20868]; 79451 MW, 5.77 pI, http://web.expasy.org/compute_pi/) exhibiting 85% amino acid identity with AChEs of mosquitoes, Culex pipiens [GenBank: Q86GC8], Aedes aegypti [GenBank: XP_001656977], and 92% amino acid identity with the new world sand fly, L. longipalpis AChE (partial codes, 396 amino acids, [GenBank: ABI74669]. Comparison of the amino acid sequence of PpAChE1 with the AChE protein of L. longipalpis (partial codes) and the AChEs of Cx. pipiens and Ae. aegypti by Clustal W multiple sequence alignment is shown in Figure 1. As seen in Figure 1, there was very high amino acid identity throughout the partial sequence of L. longipalpis AChE as well as substantial identity with the two mosquito AChEs. Presumptive disulfide bonds, members of the catalytic triad, and residues lining the catalytic gorge [34, 35] are indicated in the Clustal W multiple sequence alignment with AChEs of Torpedo californica [GenBank: 1EA5_A] and Drosophila melanogaster [GenBank: 1QO9_A] (Additional file 2).

Figure 1
figure 1

Clustal W 2.1 multiple sequence alignment of AChE protein sequences for Phlebotomus papatasi [GenBank: JQ922267], Lutzomyia longipalpis [GenBank: ABI74669], Culex pipiens AChE [GenBank: Q86GC8], and Aedes aegypti [GenBank: XP_001656977]. The consensus line below the aligned sequences indicates positions of conserved amino acid identity (*) or similarity (: or .). Positions of the 3 disulfide bond linkages are indicated by numbers above participating cysteine pairs. The members of the catalytic triad (S, E, H) which make up the catalytic site are indicated by (‡) above the participating amino acid.

Full size image

Recombinant PpAChE1 was expressed in the baculovirus system and used for biochemical characterization. Biochemical properties (substrate Km and relative sensitivities to inhibitors) obtained for rPpAChE1 are presented in Table 2. The enzyme was strongly inhibited by eserine, paraoxon, malaoxon, and the acetylcholinesterase inhibitor Bw284c51 and exhibited much lower sensitivity to the butyrylcholinesterase inhibitors iso-OMPA and ethopropazine. In addition, rPpAChE1 hydrolyzed 0.1 mM acetylthiocholine (AcSCh) 17-fold faster than 0.1 mM butyrylthiocholine (BuSCh), demonstrating substrate preference for AcSCh over BuSCh and was inhibited at substrate concentrations above 5 mM.

Table 2 Biochemical properties of rPpAChE
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Discussion

Biochemical properties of rPpAChE1 presented in this report are consistent with classification as an insect acetylcholinesterase as defined by Toutant [36]. There is substrate preference for acetylthiocholine over butyrylthiocholine, significant inhibition by the AChE-specific inhibitors eserine and BW284c51 with relative insensitivity to inhibitors (ethopropazine and iso-OMPA) classified as specific for butyrylcholinesterase. The enzyme is strongly inhibited by paraoxon and malaoxon (oxidized forms of parathion and malathion) and exhibits a Km for acetylthiocholine (37.9 μM) similar to values reported for recombinant acetylcholinesterases of Ae. aegypti (13.79 μM) [37], Stomoxys calcitrans (63.9 μM) [33], Haematobia irritans (31.3 μM) [38], or Musca domestica (150 μM) [39].

P. papatasi is a significant public health problem in the Middle East, Asia and Africa where it transmits L. major that causes zoonotic cutaneous leishmaniasis in humans [40]. Additionally, zoonotic cutaneous leishmaniasis can seriously affect the operational readiness of United States military personnel deployed in Iraq and Afghanistan [41]. Attempts by the U.S. military to control sand flies through the use of chemical pesticides were not highly successful, possibly due to extreme environmental conditions [14, 15], and pesticide bioassays did not reveal evidence of insecticide resistance. Chemical pesticides have been used in several ways for insect control, including area-wide applications for malaria control in the same areas inhabited by P. papatasi. Recent reports of pesticide resistance in sand flies, possibly resulting from insensitive AChE [18, 19], highlight the urgent need to characterize the mechanisms responsible for insecticide resistance and the relevance of efforts to develop rapid tests to identify resistant pest populations.

Point mutations in pest AChEs have been shown to generate altered enzymes with decreased inhibition by organophosphate [16, 21, 42–45]. The present report demonstrating very high amino acid sequence identity of the P. papatasi AChE to those of Cx. pipiens and Ae. aegypti strongly suggests that this PpAChE1 is the target site for organophosphate and will facilitate identification of PpAChE1 mutations that produce insensitivity to organophosphate or carbamate pesticides.

In mosquito vectors, very high resistance to OPs and carbamates results from a single amino acid substitution of Serine for Glycine (codon GGC, PpAChE1 pos 837, Gly→AGC, Ser) [46–50]. Ae. aegypti is reported to have the GGA codon at this position (as does our PpAChE1), and when converted to AGC, the A. aegypti enzyme has the identical high level resistance as Anopheles[49]. It has been suggested that GGA at this position is extremely unlikely to evolve to AGC, and that in nature, only Gly or Ser are found at this position, which indicates that no other amino acids are allowed in AChE1 at that position [49]. Our results suggest significant OP-insensitivity in AChE of P. papatasi would be unlikely to involve PpAChE1, unless nucleotide pos 839 exists in a polymorphic state (i.e., GGA/C) in nature. Alternatively, it is hypothesized that, if present, other AChEs in P. papatasi could provide the mechanism to develop target insensitivity as documented in Ae. aegypti where a high level of AChE insensitivity was due to substitutions at positions F455W (Tc pos Phe331) and Ile697 of AChE2 [51, 52].

Conclusions

Use of the recombinant P. papatasi AChE1 will facilitate rapid in vitro screening to identify novel PpAChE inhibitors, and comparative studies on biochemical kinetics of inhibition. Information on the identification and characterization of PpAChE1 presented here will facilitate the development of a rapid molecular assay for the GGC codon (pos. 839) in P. papatasi populations, and other molecular tools to screen for mutations giving rise to an organophosphate-insensitive PpAChE1. Molecular data on PpAChE1 could also be used in modeling studies to predict in vivo insecticidal activity for novel inhibitors as described by Naik et al. [53]. Availability of the recombinant PpAChE1 will enable the creation of mechanism-based screens to discover more effective inhibitors that may be developed to innovate safer vector control technologies. Novel synthetic carbamates have already been identified by screening using the recombinant PpAChE that present essentially equivalent inhibition of the target AChE with substantially improved target specificity resulting in significantly enhanced safety [Swale DR, Tong F, Temeyer KB, Li AY, Totrov MM, Carlier PR, unpublished].

Endnotes

1This article reports the results of research only. Mention of a proprietary product does not constitute an endorsement by the USDA for its use.

2In conducting research described in this report, investigators adhered to the “Guide for the Care and Use of Laboratory Animals,” as promulgated by the Committee on Care and Use of Laboratory Animals of the Institute of Laboratory Animal Resources, National Research Council. A protocol describing routing procedures for animal care and use is on file with the Animal Care Committee at the research location.

3USDA is an equal opportunity provider and employer.

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5Disclaimer: "The views expressed in this article are those of the authors and do not necessarily reflect the official policy or position of the Department of the Navy, Department of Defense, nor the U.S Government.”