2. Dr OshaghiRTL Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 8 Original Article Blood Meal Identification in Field-Captured Sand flies: Comparison of PCR-RFLP and ELISA Assays N Maleki-Ravasan1, *MA Oshaghi 2, E Javadian 2, Y Rassi 2, J Sadraei 1, F Mohtarami 2 1Department of Medical Parasitology and Entomology, College of Medical Sciences, Tarbiat Modares University, Tehran, Iran 2Department of Medical Entomology and Vector Control, School of Public Health, Tehran University of Medical Sciences, Tehran, Iran (Received 8 Jun 2009; accepted 28 July 2009) Abstract Background: We aimed to develop a PCR-RFLP assay based on available sequences of putative vertebrate hosts to identify blood meals ingested by field female sand fly in the northwest of Iran. In addition, the utility of PCR-RFLP was compared with ELISA as a standard method. Methods: This experimental study was performed in the Insect Molecular Biology Laboratory of School of Public Health, Tehran University of Medical Sciences, Iran in 2006-2007. For PCR-RFLP a set of conserved vertebrate primers were used to amplify a part of the host mitochondrial cytochrome b (cyt b) gene followed by digestion of the PCR products by Hae III enzyme. Results: The PCR-RFLP and ELISA assays revealed that 34% and 27% of field-collected sand flies had fed on hu- mans, respectively. Additionally, PCR-RFLP assays could reveal specific host DNA as well as the components of mixed blood meals. Results of PCR-RFLP assay showed that the sand flies had fed on cow (54%), human (10%), dog (4%), human and cow (21%), dog and cow (14%), and human and dog (3%). Conclusion: The results can provide a novel method for rapid diagnosis of blood meal taken by sandflies. The advantages and limitations of PCR and ELISA assays are discussed. Keywords: Leishmaniasis, sand flies, blood meal, PCR, ELISA, Iran Introduction Leishmaniasis is caused by single-celled parasites of the genus Leishmania and is spread to humans through the bite of the sand fly. There are three main forms of leishmaniasis: cutaneous, mucocutaneous and visceral, each form is caused by different species of Leishma- nia (WHO 1990). Leishmaniasis affects over 12 million people distributed in 88 countries (Sakthianandeswaren et al. 2009). Every year, new cases amount to more than 2 million (Sak- thianandeswaren et al. 2009). Nearly one tenth of the world population is at risk of infection. These figures have led to the WHO to consider leishmaniasis as one of the most serious dis- eases of the world. Visceral leishmaniasis (VL) or kala-azar is the most severe manifestation of the disease and results fatal when is not treated promptly. There are 500,000 new cases of kala- azar every year and it is endemic in Asia, Europe, and South America (WHO 2002). The parasite is transmitted by sand flies, particularly members of the genera Phleboto- mus spp. and Lutzomyia spp. which are found in a wide range of habitat, from desert to tropi- cal rain forest. Sand flies take blood meals from a wide variety of hosts, including human, live- stock, dogs and chickens (Lainson and Rangel 2005). The Human Blood Index (HBI, pro- portion of blood meals of a haemophagous in- sects population obtained from man) is relevant Corresponding author: Dr Mohammad Ali Oshaghi, E-mail: moshaghi@sina.tums.ac.ir Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 9 to epidemiological assessment and to the modi- fication of measures to interrupt any Vector Borne Diseases transmission since the vectorial capacity of the vector varies as the square of the HBI (Macdonald 1957, Garrett-Jones 1964). Detailed knowledge of the feeding behavior of sand flies on their various vertebrate hosts is considered to be a prerequisite for a successful sand flies and leishmaniasis control program. Identification of the blood meals of ha- ematophagous insects to date has largely de- pended on serological techniques such as the precipitin test, latex agglutination test and the enzyme-linked immunosorbent assay (ELISA) (Boorman et al. 1977, Washino and Tempelis 1983, Beier et al. 1988, Gomes et al. 2001, Mwangangi et al. 2003). Although these meth- ods have yielded important information on the identity of the vertebrate hosts of many blood- feeding arthropods, they are time-consuming and lack sensitivity. So, an alternative method may be desired in laboratories not set up to perform immunologic assays, or if samples are already in the form of extracted DNA. The PCR-based identification of arthropod blood meals provides a convenient alternative for labo- ratories using primarily DNA-based techniques, and may be necessary when the study design already requires the use of individual DNA ex- tractions for multiple purposes such as species confirmation, determination of infection status for various pathogens, and vector population genetic studies. Furthermore, engorged speci- mens collected in the field may be preserved dry, stored for long periods of time, and tested at facilities that may be physically distant from the point of collection. PCR-based identification of vertebrate host blood meals is a potentially convenient alter- native, which has already been performed on several vectors including ticks (Pichon et al. 2003, Estrada-Peña et al. 2005), triatomine bugs (Bosseno et al. 2006, Pizarro et al. 2007) and mosquitoes. PCR based on primers de- signed from multiple alignments of the mitocho- ndrial cytochrome b gene have identified avian and mammalian hosts of various species of mosquito (Ngo and Kramer 2003, Kent and Norris 2005, Molaei et al. 2006, Kent et al. 2006). PCR-RFLP cytochrome b analysis was also used to identify the origin of blood meals in the tick Ixodes ricinus (Kirstein and Gray 1996), tsetse flies (Steuber et al. 2005) and the mosquito Anopheles stephensi (Oshaghi et al. 2007a, and b). In their study, the cyt b se- quences showed sufficient inter-specific poly- morphism to distinguish between human, cow, sheep, chicken, and guinea pig hosts. Until recently sand fly host identification by blood meal analysis had been limited to serological studies using ELISA (Gomez et al. 1998, Agrela et al. 2002, Bongiorno et al. 2003, Svobodová et al. 2003, Marassá et al. 2006, Rossi et al. 2008), counter immunoelectropho- resis (Morsy et al. 1993), agarose gel diffusion (Srinivasan and Panicker 1992), precipitin test (Tesh et al. 1971, Tesh et al. 1972, Javadian et al. 1977, Morrison et al. 1993, Nery et al. 2004, Afonso et al. 2005) and a more laborious his- tological technique (Guzman et al. 1994). The first PCR-based method using the prepronoci- ceptin gene has been recently described (Haouas et al. 2007). In Iran, most studies for identifi- cation of sand fly blood meal have been based on ELISA (Javadian et al. 1977, Azizi et al. 2006, Rassi et al. 1999, 2005, Maleki 2007). In the present study, we adapted a PCR protocol designed by Kent and Norris (2005) and developed this with sequence analysis of major vertebrate hosts in north of Iran in order to find putative restriction enzyme to develop host specific RFLP patterns for DNA sources in sand flies. Also the ELISA method was applied to compare the results of these two methods for blood meal identification. Materials and Methods Study area The study was conducted in Germi dis- trict, Ardabil Province, in northwestern Iran. This region is 1,490 meters above the sea Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 10 level. The total population of Germi was ap- proximately 123,000 in 2002. The weather is hot (up to 40 °C) in summer and cold (less than −20 °C) in winter. The warm season is short (mid-May to mid-September). Annual rainfall is approximately 114 mm. The main occupa- tions of the population are farming and raising animals. On the basis of available epidemiologic data obtained from the Ministry of Health (MOH), local health authorities, and medical centers in Germi district, villages with higher incidence of VL were selected for the study. Three primary villages (Kalansora, Shah-Tapeh- si, and Hamzeh-Khanloo) were selected and analyzed in an entomologic survey. Three sec- ondary villages (Hasi-Kandy, Sarv-Aghaji, and Ghasem-Kandy) were also study periodically. Sample collection Sticky traps were used to collect sand flies from human and animal dwellings, rodent and fox burrows, under bridges, and on the shores of rivers. Traps were set at dusk and flies were collected at dawn. A total of 150– 200 sticky traps were set each day in each village. Sample collection began in early July 2008 and continued until late September 2008 when sand fly activity was reduced sharply. Sampling was carried out every 3 d in the pri- mary villages and 1-3 times in the secondary villages. Trapped sand flies removed from stick papers with needles, washed with absolute ethanol, and transferred into micro tubes filled with 96% ethanol. Tubes were kept frozen (-20 °C) until species identification and DNA extraction. Of the many samples collected in the region, 400 blood-fed females were ran- domly selected for detection of blood meal. Identification of sand fly species In the laboratory, samples were washed with detergent and double-distilled water, and heads and terminal abdomens of females were removed and mounted with Pouri solution on glass slides for diagnosis. For males, only heads were removed and mounted with Pouri solu- tion on glass slides. Species were identified by using specific morphologic keys. Middle parts of female sand flies were placed in micro tubes and kept frozen (−20 °C) until DNA extraction. Analysis of blood meals in sand flies Four hundred blood fed female sand flies were randomly selected for blood meal iden- tification. Sand flies were selected on the basis of location and capture sites to obtain a repre- sentative sample of sand flies in a region. Sam- ples were divided randomly into two groups of 200 specimens; each group was analyzed by PCR-RFLP or ELISA method. Extraction of DNA from blood meals in sand flies DNA extraction from blood-fed female, and male, unfed female sand flies and water (used as a negative controls), and human and cow (used as a positive controls) was con- ducted according to the procedure of Steiner et al. (1995). Samples were individually dis- rupted by mechanical homogenization in buffer containing 10 mM Tris-HCl, pH 8.0, 312.5 mM EDTA, 1% (w/v) sodium lauryl sarcosine, and 1% polyvinyl pyrolidone. Homogenates were heated to 90 °C for 20 min and chilled on ice for 5 min. Samples were centrifuged at 13,000 RPM for 5 min at room temperature. The supernatant was removed and diluted 20- fold in 10 mM Tris-HCl, pH 8.0, 1 mM EDTA. PCR amplification of the mtDNA cyt b gene Two regions of the mtDNA cyt b gene were amplified for host blood meal iden- tification of the blood-fed female specimens. For identification of human blood meals a por- tion (358 bp) of the cyt b gene was amplified and digested with XhoI enzyme as previously explained by Oshaghi et al (2006a). The se- quence of the primers used were 5΄-CCATCCA- ACATCTCAGCATGATGAAA-3 ́ (forward) and 5΄-CCCCTCAG AATGATATTTGTCCT- CA-3 ́ (reverse) (Kocher et al. 1989, Boakye et al. 1999). The PCR amplifications were per- Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 11 formed in 25 µ L of a solution containing 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl 2, 0.001% gelatin, 200 mM deoxynu- cleotide triphosphates, 10 pmol of each primer, 1 unit of Taq DNA polymerase (Cinagene, Tehran, Iran), and 2.5 µ l of DNA template solution. Samples were incubated at 95 °C for 3.5 min; followed by 36 cycles at 95 °C for 30 s, 58 °C for 50 s, and 72 °C for 40 s; and 72 °C for 5 min. To discriminate animal host blood meals, a second region of the mtDNA cytB gene was amplified by using the proto- col of Kent and Norris (2005). The sequences of forward and reverse primers were respec- tively 5΄-TGAGGACAAATATCATTCTGA- GG-3 ́ (UNFOR403) and 5 -́GGTTGTCCTCC- AATTCATGTTA-3 ́ (UNREV1025), respec- tively. Primers amplified a 623-basepair region of the cytB gene of vertebrate mtDNA. The PCR amplifications were performed in 25 µ L of a solution containing 10 mM Tris, pH 8.3, 50 mM KCl, 1.5 mM MgCl 2, 0.01% gelatin, 1.0 mM deoxynucleotide triphosphates, 0.5 units of Taq polymerase, 50 pmol of each primer, and 2.5 µ L of extracted DNA. Samples were incu- bated at 95 °C for 5 min; followed by 35 cycles at 95 °C for 1 min, 58 °C for 1 min, and 72 °C for 1 min; and 72 °C for 7 min. Products were visualized by electrophoresis on 2% agarose gels stained with ethidium bromide. Electrophoresis was conducted using a GeneRuler 100-basepair molecular mass marker (Cinagene). Sequence analysis for selection of restric- tion enzymes Available sequences of the 623 bp for human and probable vertebrate hosts (cow, goat, horse, ass, dog, and other Canidae) in the study area were obtained from Gene Bank and checked for species-specific restriction en- zyme sites for each host DNA using the Neb- cutter program (http:/tools.neb.com/nebcutter. Twenty eight sequence analyses showed that Hae III did not have a restriction site on hu- man PCR products but it has various specific sites in PCR products for other vertebrates. This enzyme was selected for discrimination of the blood meal sources within sand flies. Diges- tion of PCR products was performed in 25 µL of a solution containing 15 µL of PCR product mixed with 2.5 µL of enzyme buffers and 5 units of the restriction enzyme overlaid with two drops of mineral oil. The mixture was incubated at the temperature recommended by enzyme suppli- ers. An aliquot (14 µL) of the digestion product was mixed with 6 mL of loading buffer (0.25% bromophenol blue, 0.25% xylene cyanol, 30% glycerol), loaded onto a 2.5% agarose gel, and subjected to electrophoresis. Gels were stained with ethidium bromide (2 mg/mL) and the RFLP profiles were visualized under ultraviolet light. Serological analysis Analysis was performed by ELISA as de- scribed by Edrissian et al. (1985) the abdomen of blood fed sand flies was dissected, placed in the well of a micro-ELISA plate (Nunc, Ros- kilde, Denmark), squashed with a glass rod, and eluted with 50 µL of distilled water for 2 h at room temperature. Fifty micro liters of coating buffer (carbonate bicarbonate, pH 9.6) was then added to each well. Plates were washed three times with phosphate-buffered saline, Tween 20, pH 7.2. Fifty micro liters of diluted goat anti-human IgG conjugated to alkaline phos- phates were added onto each well, incubated for 2 h at 37 °C, washed three times with phos- phate-buffered saline, Tween 20, pH 7.2. One hundred microliters of substrate solution (1 mg/ mL of p–nitrophenyl phosphate [Sigma, St. Louis, MO] in 10% diethanolamine buffer, pH 9.8, containing 0.5 mmol MgCl 2 and 0.02% NaN 3) was added to each well, and incubated in a dark chamber for 30 min at room temperature. Two wells that did not contain blood were used as negative controls and two wells that con- tained human blood were used as positive con- trols. Results were visually assessed, and abso- rbance was measured with an ELISA reader at 405 nm approximately 30 min after addition of substrate solution. The test well result was con- sidered positive if it a yellow color was observed. Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 12 Results PCR-RFLP DNA isolated from the blood-fed sand flies, positive controls (blood from a human and a cow), and negative controls (water, un- fed female sand fly, and a male sand fly) were used as a template in a PCR. Most host DNAs were amplified, and negative controls yielded no PCR product. This result implied that only host, but not sand fly, DNA patterns were detected in amplified specimens. DNA sequence analysis showed that the two regions of the mtDNA cyt b gene digested with either Xho I or Hae III could distinguish human DNA in blood of blood-fed sand flies from the DNA of blood of other vertebrates. Xho I di- gested only the 358-basepair PCR product of human DNA and produced two bands (215 bp and 143 bp) whereas it had no restriction site in the DNA of blood of other vertebrates. This finding was confirmed by results of di- gestion with Hae III on the 623-basepair region of the cyt b gene. This enzyme did not digest the 623 bp fragment of the cyt b gene in DNA from human blood, but it digested the equivalent DNA fragment from other verte- brates and could distinguish DNAs from cow, ass, goat, horse, dog, and other Canidae from each other. For example, Hae III produced two fragments of 345 bp and 304 bp from cow DNA, two fragments of 552 bp and 70 bp from Canidae DNA, and two fragments of 170 bp and 453 bp from goat DNA (Fig. 1). Based on PCR-RFLP analysis rates of blood meal sources was found to be 54% for cows, 19% mix of humans and cows, 14% mix of dogs and cows, 8% for humans, 3% mix of humans and dogs, and 2% for dogs. On overall, 34% of the female sand flies had human blood or mix of human and an animal blood. Except for a few specimens (n=4) that contained P. (Adlerius) spp., all blood-fed specimens were P. perfiliewi. Details of PCR-RFLP analysis are shown in Table 1. ELISA Serologic analysis by ELISA on 200 blood-fed sand flies showed that 54 (27%) spe- cimens fed on humans (Table 2). Fifty one of 54 human blood-fed samples were of P. perfe- liewi. The three other seropositive samples in human blood were P. (Adlerius) spp. Approxi- mately 61% of these blood-fed sand flies were found in either human or animal shelters, which suggested that they were highly endophilic. Generally, based on PCR-RFLP and ELISA assays, it seems that sand flies, par- ticularly P. perfiliewi, although have close re- lationship with human dwellings in the study area, but are more zoophilic than anthropo- philic (73-66% versus 27-34%), and in a de- scending order, they prefer to feed on cows, humans, and dogs. Table 1. Host blood meals ingested by female sand flies in Germi, northwest of Iran detected by PCR-RFLP analy- sis of a 623 bp fragment of the mitochondrial DNA cytochrome b (Cyt b) gene. Collection site Sand fly species Blood sources n (%) P. perfiliewi Human and Cow 4 (2) Human dwelling P. (Adlerius) spp. Human 4 (2) Human 16 (8) Dog 4 (2) Cow 108 (54) Human and Cow 36 (19) Human and Dog 6 (3) Animal dwelling P. perfiliewi Dog and Cow 28 (14) Artificial P. perfiliewi Dog 4 (2) Total – – 200 (100) Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 13 Table 2. Results of ELISA assay against human antibody for blood meal identification of fed female sand flies in Germi, northwest of Iran. Capture site Sand fly species No. of blood fed (%) No. of Human blood (%) Human P. perfiliewi P. (Adlerius) spp. * 38 (14) 11 (4) 9 (23) 3 (28) Animal P. perfiliewi 46 (17) 21 (46) Artificial P. perfiliewi P. (Adlerius) spp. * P. papatasi P. kandelaki 70 (36) 24 (9) 3 (1) 8 (4) 21 (30) 0 0 0 Total - 200 (100) 54 (27) *Males of P. brevis, P. halepensis, and P. longiductus of the Adlerius subgenus are morphologically indistinguishable. Fig. 1. Electrophoresis of mitochondrial DNA cytochrome B gene fragments from blood meals of sand flies di- gested with HaeIII. Partial fragments (623 bp) of the cytochrome B gene from vertebrate hosts (human, cow, dog) of sand flies were amplified by a polymerase chain reaction and the products were digested with HaeIII. Lane 1, mix- ture of human and dog blood; Lane 2, cow blood; Lane 3, mixture of cow and dog blood; Lane 4, human blood; Lane 5, dog blood; Lane 6, mixture of human and cow blood; lane M, molecular weight marker (100 bp Cinnagen, Iran) Discussion In this study first we have adapted a PCR method developed by Kent and Norris (2005) and then we developed the technique by restriction fragment length polymorphisms (PCR-RFLP) to identify blood meals from wild- collected sand flies in an endemic area of zoo- notic visceral leishmaniasis in northwest Iran where L. infantum is transmitted. We also evaluated utility of PCR-RFLP with ELISA as a gold standard method. In our study succes- sful amplification was obtained from genomic DNA of host blood meal of sand flies fed on human, cow, and dogs. The RFLP profiles for these vertebrates were specific and easily they could be distinguished from each other. Theo- retically, based on sequence data available in Gene bank, the HaeIII enzyme could provide diagnostic profiles for other vertebrate hosts such as ass, goat, horse, and other Canidae, however, we did not have DNA sources for these ones to confirm their diagnostic profiles. Results of ELISA were almost similar with PCR-RFLP (27% versus 34%). ELISA method has provided countless valuable data Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 14 over the years for blood meal identification (Gomez et al. 1998, Agrela et al. 2002, Bon- giorno et al. 2003, Svobodová et al. 2003, Marassá et al. 2006, Rossi et al. 2008). In sand flies with 0.5 µ g engorged meal, the ELISA technique enables us to identify only one patent feed (Lehane 2005). However, ELISA methods are time-consuming and lack sensitivity. In addition, one of the major con- straints of ELISA is that sand flies are diminu- tive insects, only able to ingest very small quan- tities of blood (Rogers et al. 2002). Due to low volume of engorged blood meal in sand flies it is impossible to check more than one antigen source. Another constrain is that sand flies have multiple feeding habituate that has major consequence on the epidemiology of Zoo- notic diseases such as zoonotic visceral leishma- niasis (ZVL) and zoonotic cutaneous leishma- niasis (ZCL). These constrains direct researchers to use more sensitive and accurate methods such as PCR-RFLP to identify sand fly hosts for the study of vector–vertebrate host associations, as well as to improve current control interven- tions targeting sand flies vectors. PCR-RFLP assay has more advantage than ELISA since it possesses the unique abil- ity to analyze components of a multi species DNA sample. In this study, 36% of blood- fed field specimens contained multiple blood meals. This host-feeding behavior can influence pathogen transmission through increased fre- quency of vector-human contact, or possibly re- duce vector-human contact if some blood meals are taken from alternative mammalian hosts. Since this method could identify multiple blood meals it will be extremely useful for entomo- logically based projects involving blood-feed- ing behavior and vectorial capacity of sand flies in endemic areas of ZVL and ZCL. mtDNA Cytochrome B gene has been used to resolve vertebrate evolutionary ques- tions as well as served as a target for molecu- lar diagnostics (Ngo 2003). CytB has a proven utility for identifying arthropod blood meals due to high copy number as a mitochondrial gene and sufficient genetic variation at the primary sequence level among vertebrate taxa for reli- able identification. In addition to mtDNA cytB gene, other markers that have been used to identify blood meals from arthropods include vertebrate 18S ribosomal DNA (Pichon 2003) the hypervariable region 2 of mitochondrial DNA (Lord 1998) and TC-11 and VWA (HUMVWFA31/A, a repeat polymorphism in the von Will brand factor gene) loci (Muka- bana 2002). PCR-based methods have been used for diagnosis of infectious diseases, including Leish- mania detection in human patients (Dweik et al. 2007, Foulet et al. 2007, Kumar et al. 2007), infected dogs (de Andrade et al. 2006, Gomes et al. 2007, Solano-Gallego et al. 2007) and phlebotomine sand flies (Cabrera et al. 2002, Paiva et al. 2006, Myskova et al. 2008, Ranasinghe et al. 2008, Oshaghi et al. 2009). DNA prepared from whole body of Phle- botomus sand flies not only can be used for blood meal identification but also can be used for parasite detection/identification by PCR (Sant'Anna et al. 2008). This simple method- ology could be very useful in epidemiological studies in endemic areas for leishmaniasis as specimens suspected to contain parasites. In this study, sand fly of P. perfiliewi was the most predominant species and has a 27-34% tendency to human blood in the re- gion. Previous studies on P. perfiliewi, have suggested nearly identical tendency to human blood (Rassi 1999). It seems that this sand fly is not highly anthropophilic and preferred cows or other animals. In rural environment where large domestic mammalian species occur in abu- ndance, this may reduce P. perfiliewi vecto- rial capacity. In conclusion, blood meal identification in field-caught sand flies can confirm a strong association between sand flies and reservoir hosts such as dogs in rural areas and help to improve understanding the role of domestic animals in transmission of L. infantum in en- demic foci. The greater sensitivity of the PCR- Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 15 RFLP method in comparison with ELISA re- ported here means that information can be ob- tained from specimens that have ingested re- latively small amounts of blood of one or mul- tiple hosts which could be used in vector in- crimination and reservoir determination of vec- tor borne diseases such as ZVL and ZCL. Acknowledgments We thank Abolhassani, Eskandari, and Hosseni for their perfect technical assistance. This project was founded by Tehran Univer- sity of Medical Sciences (Grant number: 86- 03-27-6268) to MA.Oshaghi. The authors de- clare that they have no conflicts of interest. References Afonso MM, Gomes AC, Meneses CR, Rangel EF (2005) Studies on the feed- ing habits of Lutzomyia (N.) interme- dia (Diptera, Psychodidae), vector of cutaneous leishmaniasis in Brazil, Cad Saude Publica. 21: 1816–1820. Agrela I, Sanchez E, Gomez B, Feliciangeli MD (2002) Feeding behavior of Lutzo- myia pseudolongipalpis (Diptera: Psy- chodidae), a putative vector of visceral leishmaniasis in Venezuela. J Med En- tomol. 39: 440–445. Andrade HM, Reis AB, Santos SL, Volpini AC, Marques MJ, Romanha AJ (2006) Use of PCR-RFLP to identify Leishma- nia species in naturally-infected dogs. Vet Parasitol. 140: 231–238. Azizi K, Rassi Y, Javadian E, Motazedian MH, Rafizadeh S, Yaghoobi Ershadi MR, Mo- hebali M (2006) Phlebtomus (Paraphleb- tomus) alexandri a Probable Vector of Leishmania infantum in Iran. J Ann Trop Med Parasitol. 100(1): 63–68. Beier JC, Perkins PV, Wirtz RA, Koros J, Diggs D, Gargan TP, Koech DK (1988) Blood meal identification by direct enzy- me-linked immunosorbent assay (ELISA), tested on Anopheles (Diptera: Culicidae) in Kenya. J Med Entomol. 25: 9–16. Boakye DA, Tang J, Truc P, Merriweather A, Unnasch TR (1999) Identification of bloodmeals in haematophagous Diptera by cytochrome B heteroduplex analysis. Med Vet Entomol. 13: 282–287. Bongiorno G, Habluetzel A, Khoury C, Maroli M (2003) Host preferences of phlebo- tomine sand flies at a hypoendemic focus of canine leishmaniasis in central Italy. Acta Trop. 88: 109–116. Boorman J, Mellor PS, Boreham PFL, Hewett RS (1977) A latex agglutination test for the identification of blood-meals of Cu- licoides (Diptera: Ceratopogonidae). Bull Entomol Res. 67: 305–311. Bosseno MF, García LS, Baunaure F, Gastelúm EM, Gutierrez MS, Kasten FL, Dumonteil E, Brenière SF (2006) Identification in triatomine vectors of feeding sources and Trypanosoma cruzi variants by hetero- duplex assay and a multiplex miniexon polymerase chain reaction. Am J Trop Med Hyg. 74: 303–305. Cabrera OL, Munsterman LE, Cárdenas R, Gutiérrez R, Ferro C (2002) Definition of appropriate temperature and storage conditions in the detection of Leishmania DNA with PCR in phlebotomine flies. Biomedica. 22: 296–302. Dweik A, Schönian G, Karanis P (2007) Evaluation of PCR-RFLP (based on ITS-1 and Hae III) for the detection of Leishmania species, using Greek canine isolates and Jordanian clinical material. Ann Trop Med Parasitol. 101: 399–407. Edrissian Gh, Manochehri AV, Hafizi A (1985) Application of an Enzyme-Linked Immu- nosorbent Assay (ELISA) for determina- tion of the human blood index in ano- pheline mosquitoes collected in Iran. J Am Mosq Cont Assoc. 3:349–352. Estrada-Peña A, Osácar JJ, Pichon B, Gray JS (2005) Hosts and pathogen detection for immature stages of Ixodes ricinus Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 16 (Acari: Ixodidae) in north-central Spain. Exp Appl Acarol. 37: 257–268. Foulet F, Botterel F, Buffet P, Morizot G, Rivollet D, Deniau M, Pratlong F, Costa JM, Bretagne S (2007) Detection and identification of Leishmania species from clinical specimens using real-time PCR assay and sequencing of the cytochrome b gene. J Clin Microbiol. 45: 2110–2115. Garrett-Jones C (1964) Prognosis for interrup- tion of malaria transmission through as- sessment of the mosquito's vectorial ca- pacity. Nature. 4964: 1173–1175. Gomes AH, Ferreira IM, Lima ML, Cunha EA, Garcia AS, Araújo MF, Pereira- Chioccola VL (2007) PCR identification of Leishmania in diagnosis and control of canine leishmaniasis. Vet Parasitol. 144: 234–241. Gomes LA, Duarte R, Lima DC, Diniz BS, Serrão ML, Labarthe N (2001) Compari- son between precipitin and ELISA tests in the bloodmeal detection of Aedes ae- gypti (Linnaeus) and Aedes fluviatilis (Lutz) mosquitoes experimentally fed on feline, canine and human hosts. Mem Inst Oswaldo Cruz. 96: 693–695. Guzman H, Walters LL, Tesh RB (1994) Histologic detection of multiple blood meals in Phlebotomus duboscqi (Diptera: Psychodidae). J Med Entomol. 31: 890– 897. Haouas N, Pesson B, Boudabous R, Dedet JP, Babba H, Ravel (2007) Development of a molecular tool for the identification of Leishmania reservoir hosts by blood meal analysis in the insect vectors. Am J Trop Med Hyg. 77: 1054–1059. Javadian E, Tesh R, Saidi S, Nadim A (1977) Studies on the epidemiology of sandfly fever in Iran. III. Host-feeding patterns of Phlebotomus papatasi in an endemic area of the disease. Am J Trop Med Hyg. 26: 294-298. Kent RJ, Norris DE (2005) Identification of mammalian blood meals in mosquitoes by a multiplexed polymerase chain reac- tion targeting cytochrome b. Am J Trop Med Hyg. 73: 336–342. Kent RJ, Coetzee M, Mharakurwa S, Norris DE (2006) Feeding and indoor resting behaviour of the mosquito Anopheles lon- gipalpis in an area of hyperendemic ma- laria transmission in southern Zambia. Med Vet Entomol. 20: 459–463. Kirstein F, Gray JS (1996) A molecular marker for the identification of the zoonotic res- ervoirs of Lyme borreliosis by analysis of the blood meal in its European vector Ixodes ricinus. Appl Environ Microbiol. 62: 4060-4065. Kocher TD, Thomas WK, Meyer A, Edwards SV, Paabo S, Villablanca FX, Wilson AC (1989) Dynamics of mitochondrial DNA evolution in animals: amplification and se- quencing with conserved primers. Proc Natl Acad Sci USA. 86: 6196-6200. Kumar R, Bumb RA, Ansari NA, Mehta RD, Salotra P (2007) Cutaneous leishmani- asis caused by Leishmania tropica in Bikaner, India: parasite identification and characterization using molecular and im- munologic tools. Am J Trop Med Hyg. 76: 896–901. Lainson R, Rangel EF (2005) Lutzomyia longipalpis and the eco-epidemiology of American visceral leishmaniasis, with particular reference to Brazil: a review. Mem Inst Oswaldo Cruz. 100: 811–827. Lehane MJ (2005) The biology of blood sucking insects. Cambridge University Press, London. Lord WD, DiZinno JA, Wilson MR, Budowle B, Tapalin D, Meinking TL (1998) Iso- lation, amplification, and sequencing of hu- man mitochondrial DNA obtained from human crab louse, Pthirus pubis (L.), blood meals. J Forensic Sci. 43:1097–1100. Maleki-Ravasan N (2007) The Study of Sand flies Infection to Leishmania infantum Using Molecular Methods (PCR-RFLP) in Ardebil Province 2006-2007. [MSc dis- Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 17 sertation]. Department of Medical Para- sitology and Entomology, Colledge of Medical Sciences, Tarbiat Modaress Uni- versity, Iran. Marassá AM, Consales CA, Galati EA, Nunes VL (2006) Blood meal identification of Lutzomyia (Lutzomyia) longipalpis (Lutz and Neiva, 1912) and Lutzomyia (Lut- zomyia) almerioi (Galati and Nunes, 1999) by enzyme-linked immunosorbent assay biotin-avidin. Rev Soc Bras Med Trop. 39: 183–186. Molaei G, Oliver J, Andreadis TG, Armstrong PM, Howard JJ (2006) Molecular iden- tification of blood-meal sources in Cu- liseta melanura and Culiseta morsitans from an endemic focus of eastern equine encephalitis virus in New York. Am J Trop Med Hyg. 75: 1140–1147. Morrison AC, Ferro C, Tesh RB (1993) Host preferences of the sand fly Lutzomyia longipalpis at an endemic focus of Ameri- can visceral leishmaniasis in Colombia. Am J Trop Med Hyg. 49: 68–75. Morsy TA, Aboul Ela RG, Abdelmawla MM, Gozamy BM (1993) Counter immunoe- lectrophoresis, a modified technique for the identification of blood meals of sand- flies collected from Qualyobia Governo- rate. J Egypt Soc Parasitol. 23: 109–132. Mukabana WR, Takken W, Seda P, Killeen GF, Hawley WA, Knols BGJ (2002) Extent of digestion affects the success of amplifying human DNA from blood meals of Anopheles gambiae (Diptera: Culici- dae). Bull Entomol Res. 92: 233–239. Mwangangi JM, Mbogo CM, Nzovu JG, Githure JI, Yan G, Beier JC (2003) Blood-meal analysis for anopheline mos- quitoes sampled along the Kenyan coast. J Am Mosq Control Assoc. 19: 371–375. Myskova J, Votypka J, Volf P (2008) Leish- mania in sand flies: comparison of quanti- tative polymerase chain reaction with other techniques to determine the intensity of in- fection. J Med Entomol. 45: 133–138. Nery LC, Lorosa NE, Franco AM (2004) Feeding preference of the sand flies Lutzomyia umbratilis and L. spathotri- chia (Diptera: Psychodidae, Phlebotomi- nae) in an urban forest patch in the city of Manaus, Amazonas, Brazil. Mem Inst Oswaldo Cruz. 99: 571–574. Ngo KA, Kramer LD (2003) Identification of mosquito bloodmeals using polymerase chain reaction (PCR) with order-specific primers. J Med Entomol. 40: 215-222. Oshaghi MA, Chavshin AR, Vatandoost H (2006a) Analysis of mosquito blood meals using RFLP Markers. Exp Parasitol. 114: 259–264. Oshaghi MA, Chavshin AR, Vatandoost H, Yaaghoobi F, Mohtarami F, Noorjah N (2006b) Effects of post-ingestion and phy- sical conditions on PCR amplification of host blood meal DNA in mosquitoes. Exp Parasitol. 112: 232–236. Oshaghi MA, Maleki-Ravasan N, Javadian E, Mohebali M, Hajjaran H, Zare Z, Mohtarami F, Rassi Y (2009) Vector In- crimination of Sand Flies in the Most Im- portant Visceral Leishmaniasis Focus in Iran. Am J Trop Med Hyg. (in press). Paiva BR, Secundino NF, Nascimento JC, Pimenta PF, Galati EA, Junior HF, Malafronte RS (2006) Detection and iden- tification of Leishmania species in field- captured phlebotomine sandflies based on mini-exon gene PCR. Acta Trop. 99: 252–259. Pichon B, Egan D, Rogers M, Gray J (2003) Detection and identification of pathogens and host DNA in unfed host-seeking Ixodes ricinus L. (Acari: Ixodidae). J Med Entomol. 40: 723–731. Pizarro JC, Lucero D, Stevens L (2007) A method for the identification of guinea pig blood meal in the Chagas disease vector, Triatoma infestans. Kinetoplastid Biol Dis. 6: 1. Ranasinghe S, Rogers ME, Hamilton JGC, Bates PA, Maingon RDC (2008) A real Iranian J Arthropod-Borne Dis, (2009), 3(1): 8-18 N Maleki-Ravasan et al: Blood Meal Identification… 18 time PCR assay to estimate Leishmania chagasi load in its natural sand fly vec- tor Lutzomyia longipalpis. Trans R Soc Trop Med Hyg. (in press). Rassi Y, Javadian E (1999) Study on Phleb- tomus perfiliewi the probable vector of visceral leishmaniasis in northwest of Iran (1995-1998). 3rd International sympo- sium on Phlebtominae sand flies, 1999 August 23–27, Montpollier, France, p 1. Rassi Y, Javadian E, Nadim A, ZAhraii A (2005) Phlebtomus (Larrossius) kande- lakii the Principle and proven Vector of Visceral Leishmaniasis in Northwest of Iran. Pak J Bio Sc. 8(12): 1802–1806. Rogers ME, Chance ML, Bates PA (2002) The role of promastigote secretory gel in the origin and transmission of the infective stage of Leishmania mexicana by the sandfly Lutzomyia longipalpis. Parasitol. 124: 495–507. Rossi E, Bongiorno G, Ciolli E, Muccio T, Scalone A, Gramiccia M, Gradoni L, Maroli M (2008) Seasonal phenology, host-blood feeding preferences and natu- ral Leishmania infection of Phleboto- mus perniciosus (Diptera, Psychodidae) in a high-endemic focus of canine leishma- niasis in Rome province, Italy. Acta Trop. 105: 158-165 Sakthianandeswaren A, Foote SJ, Handman E (2009) The role of host genetics in leish- maniasis. Trends Parasitol. 25(8): 383-391. Sant'Anna MR, Jones NG, Hindley JA, Men- des-Sousa AF, Dillon RJ, Cavalcante RR, Alexander B, Bates PA (2008) Blood meal identification and parasite de- tection in laboratory-fed and field-cap- tured Lutzomyia longipalpis by PCR us- ing FTA databasing paper. Acta Trop. 107(3): 230–237. Solano-Gallego L, Rodriguez-Cortes A, Trotta M, Zampieron C, Razia L, Furlanello T, Caldin M, Roura X, Alberola J (2007) Detection of Leishmania infantum DNA by fret-based real-time PCR in urine from dogs with natural clinical leish- maniosis. Vet Parasitol. 147: 315–319. Srinivasan R, Panicker KN (1992) Identifica- tion of bloodmeals of phlebotomine sand- flies using the agarose gel diffusion me- thod. Southeast Asian J Trop Med Publ Health. 23: 486–488. Steiner JJ, Poklimba CJ, Fjellestrom RG, El- liot LF (1995) A rapid one tube ge- nomic DNA extraction process for PCR and RAPD analysis. Nucl Acid Res. 23: 2569-2570. Steuber S, Abdel-Rady A, Clausen PH (2005) PCR-RFLP analysis: a promising tech- nique for host species identification of blood meals from tsetse flies (Diptera: Glossinidae). Parasitol Res. 97: 247–254. Svobodová M, Sádlová J, Chang KP, Volf P (2003) Distribution and feeding prefer- ence of the sand flies Phlebotomus ser- genti and P. papatasi in a cutaneous leishmaniasis focus in Sanliurfa, Tur- key. Am J Trop Med Hyg. 68: 6–9. Tesh RB, Chaniotis BN, Aronson MD, John- son KM (1971) Natural host preferences of Panamanian phlebotomine sandflies as determined by precipitin test. Am J Trop Med Hyg. 20: 150-156. Tesh RB, Chaniotis BN, Carrera BR, John- son KM (1972) Further studies on the natural host preferences of Panamanian phlebotomine sandflies. Am J Epidemiol. 95: 88–93. Washino RK, Tempelis CH (1983) Mosquito host blood meal identification: methodol- ogy and data analysis. Annu Rev Ento- mol. 28: 179-201. WHO (1990) Expert committee: control of the leishmaniasis. WHO Techn Ser. 793. WHO (2002) Annex 3 burden of disease in DALYs by cause, sex and mortality stra- tum in WHO regions, estimates for 2001: 192–197. World Health Organization, Geneva.