• View in gallery

    Distribution of the Culex pipiens complex and its sibling species based on maps of Dahl,35 Belkin,36 Mattingly and others,37 and available literature.12,38,39 Light gray = Cx. pipiens; black = Cx. quinquefasciatus; dark gray = overlapping ranges of Cx. pipiens and Cx. quinquefasciatus; region marked by dotted line = Cx. torrentium; region marked by solid line = Cx. australicus; region marked by dashed line = Cx. pipiens pallens; New Zealand marked by dotted and dashed line = Cx. pervigilans.

  • View in gallery

    Polymorphisms across taxa and the relative position of the unique primers. Each species is represented by the inclusive consensus (shaded regions) of all its cloned sequences (GenBank accession #AY497523-26). Position 1 corresponds to the 5′ end of exon 2 (position 1 in Figure 2A in Malcolm and others25). Position 203 corresponds to the first basepair at a Sca I-cute site.10 B = C/G/T; W = A/T; R = A/G; Y = C/T; K = G/T; S = C/G; M = A/C. Ace = acetylcholinesterase; pip = pipiens; pal = pallens; aus; australicus; torr = torrentium; quin = quinquefasciatus. fasciatus,

  • View in gallery

    Fragments amplified using the regional multiplexes of primers. A, lane 2= Culex pipiens; lane 3 = Cx. quinquefasciatus; lane 4 = Cx. pervigilans; B, lane 6 = Cx. pipiens; lane 7 = Cx. quinquefasciatus; lane 8 = Cx. australicus; C, lane 10 = Cx. pipiens, lane 11, Cx. torrentium; D, lane 13 = Cx. pipiens; lane 14 = Cx. quinquefasciatus; lane 15 = Cx. pipiens × Cx. quinquefasciatus hybrid; lane 16 = Cx. p. pallens or putative pallens × Cx. quinquefasciatus hybrid. Lanes 1, 5, 9, 12, and 17 are size standards (100-basepair ladder; New England Biolabs, Beverly, MA).

  • View in gallery

    Correlation between the frequency of Culex pipiens acetylcholinesterase (ACE) alleles and the average frequency of the “pipiens” signature (probability of ancestry from Cx. pipiens) based on a panel of eight microsatellite loci across 10 populations from the United States.

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RAPID ASSAYS FOR IDENTIFICATION OF MEMBERS OF THE CULEX (CULEX) PIPIENS COMPLEX, THEIR HYBRIDS, AND OTHER SIBLING SPECIES (DIPTERA: CULICIDAE)

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  • 1 Genetics Program, National Museum of Natural History, Smithsonian Institution, Washington, District of Columbia

Mosquitoes in the Culex (Culex) pipiens complex of species, known as vectors of periodic filariasis and deadly encephalitides, have recently emerged as important vectors of West Nile virus in the United States. Highly conserved morphology but marked differences in potential vectorial capacity require the development of polymerase chain reaction (PCR)-based tests that unambiguously distinguish among the different species. We introduce and describe a series of PCR-based assays that use polymorphisms in the second intron of the acetylcholinesterase-2 (ace-2) locus for the identification of members of the Cx. pipiens complex (Cx. pipiens, Cx. quinquefasciatus, Cx. p. pallens, Cx. australicus), two other species that are commonly mislabeled as Cx. pipiens (Cx. torrentium and Cx. pervigilans), as well as hybrids between Cx. pipiens and Cx. quinquefasciatus.

INTRODUCTION

Mosquitoes in the Culex pipiens complex are important disease vectors with global distribution, yet remain difficult to identify in the field. The rapid and accurate identification of these mosquitoes, which vector West Nile Virus and St. Louis encephalitis in the eastern United States,1–3 and periodic lymphatic filariasis (Wuchereria bancrofti),4 avian malaria,5 and other encephalitides across the world,6 is critical to control efforts. Identifying members of the Cx. pipiens complex by morphologic methods is difficult, time-consuming, and often limited to adult males.7 A rapid polymerase chain reaction (PCR) assay based on polymorphisms in ribosomal DNA was developed by Crabtree and others8 to separate Cx. pipiens complex species from other species with similar morphology in the United States (Cx. restuans and Cx. salinarius). However, this test fails to differentiate the main taxonomic units within the Cx. pipiens complex. To date, molecular techniques used to distinguish members of the complex have included allozyme analyses,9 restriction fragment length polymorphism analysis of PCR products,10 and a PCR assay developed from subtractive hybridization that distinguishes between Cx. pipiens and Cx. quinquefasciatus by the presence of an amplification product in the former and its absence in the latter.11 These techniques only distinguish between the two major taxa of the complex: Cx. pipiens and Cx. quinquefasciatus.

Members of the Cx. pipiens complex include Cx. (Cx.) pipiens L. 1758, Cx. (Cx.) quinquefasciatus Say 1823, Cx. (Cx.) pipiens pallens Coquillett 1898, and Cx. (Cx.) australicus Dobrotworsky & Drummond 1953. While the male genitalia (phallosoma) can be used to distinguish Cx. pipiens from Cx. quinquefasciatus, their hybrids commonly have intermediate shapes,12 which are similar to the phallosomes of Cx. australicus and Cx. p. pallens. This has led some to hypothesize the latter are actually hybrid forms.13 Two closely related species, Cx. (Cx.) torrentium Martini 1925, and Cx. (Cx.) pervigilans Von Bergroth 1889, are morphologically very similar to members of the Cx. pipiens complex. As a testament to their highly cryptic morphologic differences, Cx. torrentium occurs in sympatry with Cx. pipiens throughout Europe and in some parts of Asia,13 but the species escaped notice until 1925. Indeed, it remained undetected in Great Britain until 1951, although it is now known to have been present there since at least 1900.14 Culex pervigilans, which occurs exclusively in New Zealand and neighboring islands,15 closely resembles Cx. quinquefasciatus morphologically and some hybridization between the two has been hypothesized.15 Culex pipiens and Cx. quinquefasciatus are invasive, ubiquitous species, with geographic distributions closely overlapping those of humans, who are responsible for their introduction into many areas (Figure 1). Hybrid zones between the two species are known to occur in North America, Argentina, and Madagascar.9,16–18 The remaining species and subspecies in the complex have localized distributions (Figure 1). Our series of rapid assays is tailored to these geographic differences, so that a single assay uses fewer primers, reducing both the cost of the reaction and the chance for non-specific amplification.

Because of putative and realized differences in vectorial capacity within the complex,13 an assay that unambiguously identifies each taxon and may be rapidly executed even in local vector control offices is highly desirable.8 Our objective was to design a rapid and cost-effective PCR-based method in which each species/sub-species produces an amplification product of distinctive size.

On the heels of studies in which introgression between Cx. pipiens and Cx. quinquefasciatus was assessed using microsatellite markers, we also examined the ability of our simple diagnostic assay to provide information on the presence of hybrids. Those studies analyzed north-south transects between Massachusetts and Florida on the East Coast, and between Oregon and Jalisco, Mexico on the West Coast (Fonseca DM and others, unpublished data), mapping the extent of the hybrid zone between the two species using multilocus signatures19 unique to each species.20 Microsatellites are extremely polymorphic repeats of simple nucleotides motifs found in the genome of most organisms.21 Primers that amplify microsatellite-containing DNA fragments have been developed both for Cx. quinquefasciatus22 and Cx. pipiens,23 and there is a subset that will amplify in both species (Keyghobadi N and others, unpublished data). To evaluate the ability of the rapid assay to identify hybridization, we assessed the extent of introgression between Cx. pipiens and Cx. quinquefasciatus in 10 populations, using both a panel of eight microsatellite loci and the rapid assay. Previous studies have detected hybrids morphologically by differences in the shape of phallosoma, quantified as the DV/D ratio. This is the ratio between the extent to which the ventral arm of the phallosoma protrudes from the dorsal arm (DV) and the distance between the two dorsal arms (D).12 The use of microsatellites instead of the classic DV/D ratio to identify hybrids circumvents a potential bias due to the known effect of rearing temperature on the shape of the genitalia.24

The presence of two nuclear genes that encode acetylcholinesterase (ACE) was first discovered in Cx. pipiens25,26 and has since been found in other mosquito species.27 The ace-1 gene can confer resistance to organophosphate insecticides and is therefore subject to selection pressure. The ace-2 gene is sex-linked, and its exact function and the selection pressures acting on it are not known. We used polymorphisms in the second intron of the ace-2 gene to design species-specific primers for PCR-based assays.

MATERIALS AND METHODS

Mosquitoes.

Origins and sources of mosquito samples are listed in Table 1. Genomic DNA was extracted from individual mosquitoes using a phenol-chloroform based protocol as previously described.5 In addition, some samples of Cx. torrentium were extracted using a salting out method.26 Eight hybrid populations of Cx. pipiens and Cx. quinquefasciatus were first pinpointed using microsatellite allelic sizes and frequencies (Fonseca DM and others, unpublished data). The microsatellite-containing fragments (loci CQ11, CQ26, qGT4, qGT6b, pGT4, pGT9, pGT12, and pGT46)22,23 were amplified as previously described22,23 and sized in a ABI3100 capillary automatic sequencer (Applied Biosystems, Foster City, CA). Non-hybrid populations of Cx. pipiens were from Boston, Massachusetts and non-hybrid populations of Cx. quinquefasciatus were from Archer, Florida and New Orleans, Louisiana. The eight hybrid populations tested with our rapid assay are listed in Table 1; two additional populations were used only for sequencing because of their limited sample sizes. We analyzed a minimum of eight specimens from each population (mean ± SD = 8.7 ± 0.15).

Primer design.

Sequences of sections of exons 2 and 3 and the entire intron II in the ace-2 gene (the ACE locus) were obtained using the oligonucleotide primers F1457 (5′-GAG-GAGATGTGGAATCCCAA-3′) and B1246 (5′-TGGAGC-CTCCTCTTCACGGC-3′) and the conditions described in Bourguet and others.10 Except for Cx. australicus and Cx. pervigilans, we had access to several populations of each species; therefore, we amplified and cloned (TOPO TA cloning kit; Invitrogen, Carlsbad, CA) the ACE locus from three specimens from several of those populations (Table 1). We sequenced two clones per specimen using standard cycle-sequencing conditions, and analyzed the resulting fragments by electrophoresis in a slab gel (ABI 377) automated sequencer (Applied Biosystems). We aligned the sequences in Sequencher version 4.1 (GeneCodes, Ann Arbor, MI) and manually added deletions/insertions. Selective forward-primers were designed for each species so that unique polymorphisms occurred within the first 2–4 basepairs (bp) of the 3′ end (Figure 2). Each of these primers was used in conjunction with the reverse primer B1246, which was modified by removing one nucleotide from the 3′ end to reduce primer dimers7 (Table 2). We used the online program Primer 328 to check for compatibility and self-annealing in the primers. Primers were first optimized individually, and then multiplexes (mixtures of two or more primer pairs) were created and optimized so that a single amplification reaction would distinguish any of the species present in a given geographic region (Figure 3 and Table 3).

Polymerase chain reaction assay.

The PCR assays were optimized for a 20-μL volume since the product is needed only for fragment size analysis by electrophoresis on a 1.5% agarose gel. Reactions contained 1× PCR buffer, 250 μM of each dNTP, 2 mM MgCl2, 0.15 mg/mL of bovine serum albumin, one unit of Taq polymerase (Applied Biosystems), and approximately 6 ng of genomic DNA. The concentration of primers varied with geographic combination (Table 3). The amplification program consisted of one cycle at 94°C for five minutes, followed by 35 cycles at 94°C for 30 seconds, 55°C for 30 seconds, 72°C for one minute, and one cycle at 72°C for five minutes. DNA from individuals previously identified to each relevant species was included in every assay both as a positive control and as a demonstration of the expected size of each taxon-specific fragment. A negative control to which no DNA was added was also included in every run.

The diagnostic abilities of the amplification assays were tested using a minimum of six specimens from populations across the world other than those from which specimens were sequenced (Table 1) in geographically appropriate mixtures (Table 3).

RESULTS

Because ACE is a nuclear locus, the amplified PCR product had to be cloned to recover both alleles for sequencing. We cloned three individuals per population and obtained a minimum of six and a maximum of 49 (mean ± SE = 18.7 ± 7.7) sequences per species. The size of the amplified product was 634–636 bp for Cx. pipiens (n [number of cloned sequences] = 36), 626–634 bp for Cx. quinquefasciatus (n = 49), 636–641 bp for Cx. p. pallens (n = 6), 627–633 bp for Cx. australicus (n = 6), 691–706 bp for Cx. pervigilans (n = 9), and 512–513 bp for Cx. torrentium (n = 6). Intra-specifically and within the Cx. pipiens complex, the variation in the size of fragments is almost exclusively in single base pair microsatellites (runs of As or Ts). In contrast, relative to the Cx. pipiens complex, Cx. pervigilans and Cx. torrentium have large insertions (66 bp) or deletions (209 bp), respectively, in the intron region. Specimens from the Cx. pipiens × Cx. quinquefasciatus hybrid populations often produced clones that matched both those from Cx. pipiens and those from Cx. quinquefasciatus. Similarly, specimens identified as Cx. p. pallens through male genitalia analysis produced clones that matched Cx. quinquefasciatus and clones that were unique to Cx. p. pallens.

Primers were successfully designed for the identification of Cx. pipiens, Cx. quinquefasciatus, Cx. p. pallens, Cx. australicus, and Cx. torrentium. We did not design a unique primer for Cx. pervigilans since it can be amplified using the unique primer for Cx. pipiens, producing a fragment approximately 60 bp larger than in Cx. pipiens. Unfortunately, there is only a single polymorphism (A/T) that makes ACEpip unique for Cx. pipiens (Figure 2). It is therefore important to use very stringent conditions to avoid the amplification of that fragment in other species within the Cx. pipiens complex. Since the other species have their own unique fragments, this is not a major problem except during the evaluation of putative hybrids. It is important to note, however, that in hybrids the bands unique for Cx. pipiens and Cx. quinquefasciatus both amplify strongly (Figure 3), while the occasional non-specific amplification of the Cx. pipiens fragment generates a weak band (see Cx. australicus in Figure 3).

A multiplex of primers was optimized for each of five geographic regions based on which species are present there (Table 3 and Figure 1). The first regional multiplex, designed for use in Africa and the Americas, will produce a 610-bp fragment in Cx. pipiens, a 274-bp fragment in Cx. quinque-and may produce one or both fragments in hybrids of the two species (Figure 3). These are approximate sizes based on single individuals since, as mentioned, a small number of insertions and/or deletions occur intra-specifically. These fragment sizes are diagnostic for Cx. pipiens and Cx. quinquefasciatus regardless of which multiplex is used. Diagnostic primers for these two species are included in all multiplexes except Eurasia where only Cx. pipiens occurs. Besides the diagnostic fragments for Cx. pipiens and Cx. quinquefasciatus, 1) the Australian multiplex amplifies a 437-bp fragment in Cx. australicus; 2) although as mentioned earlier we did not design a primer specific for Cx. pervigilans, the New Zealand multiplex produces a 668-bp fragment in Cx. pervigilans specimens in contrast to the 610-bp fragment produced by Cx. pipiens; 3) the East Asian multiplex produces a 478-bp fragment in Cx. p. pallens and very commonly also the 274-bp fragment characteristic of Cx. quinquefasciatus in specimens with the male genitalia of Cx. p. pallens; and 4) the Eurasian multiplex produces a unique 416-bp fragment in Cx. torrentium.

Our rapid assay was able to detect hybrids between Cx. pipiens and Cx. quinquefasciatus in North America. We found that there is a strong correlation (r2 = 0.92) between the population frequency of the ACE allele unique to one of the species (in this case we used Cx. pipiens) and the average probability of ancestry from Cx. pipiens in that population based on a multilocus genotype analysis using the panel of eight microsatellite loci (Figure 4).

DISCUSSION

We found a high degree of intra-specific polymorphism in the intron region (Figure 2) of the ACE locus, especially if one considers the low levels of mitochondrial polymorphism commonly found across some of the species in the complex.5,29 The fixed polymorphisms at this nuclear intron between species in the Cx. pipiens complex and its sibling species might reveal more of the true phylogeny of the group than the mitochondrial DNA that has probably been highly modified by selective sweeps linked to Wolbachia pipientis.30

Although the primers that identify Cx. australicus, Cx. torrentium, and Cx. p. pallens generate fragments of very similar sizes, their distributions are not known to coincide. Therefore, we decided it was more important to separate each from their respective local species than from other species with which they do not co-exist. From Figure 2, it is apparent that other primers can be designed specifically to separate them if needed. It must be noted, however, that adding more primers to a reaction may require extensive optimization.

The finding that specimens identified as Cx. p. pallens through genitalia analysis will generate fragments unique to Cx. quinquefasciatus adds to the debate over the taxonomic status of Cx. p. pallens.13 In these specimens, we cloned the unique Cx. p. pallens allele as well as alleles characteristic of Cx. quinquefasciatus, leading us to hypothesize that extensive hybridization may be occurring between the two forms. However, examination of the ACE locus sequences in a phylogenetic context (Fonseca DM and others, unpublished data) does not support the hypothesis that Cx. p. pallens represents simply an extensive hybridization between Cx. pipiens and Cx. quinquefasciatus, as has been proposed.13,16 We are currently performing additional analyses which include populations of Cx. p. pallens from China and northernmost Japan. In contrast, we found that Cx. australicus is genetically well differentiated from both Cx. pipiens and Cx. quinquefasciatus, a result that is supported by further analyses of mitochondrial and microsatellite loci (Fonseca DM and others, unpublished data). Also, we did not find evidence of introgression between Cx. quinquefasciatus and Cx. pervigilans. This last observation is based on the analysis of a single population, so further study will be necessary. The assay we developed will allow a more extensive examination of this hypothesis.

The strong agreement on the degree of introgression between Cx. pipiens and Cx. quinquefasciatus across populations, obtained through an extensive microsatellite analysis and the simple PCR assay we describe, was encouraging. While the power31 to detect low levels of introgression increases with the number of independent loci being assessed, areas of moderate hybridization can be easily pinpointed using just the quickly implemented ACE PCR assay. The much higher cost of a full microsatellite analysis can therefore be kept for questions that require a high level of precision or the need to know the likely ancestry of individual specimens.20

This rapid assay does have limitations when used with pooled samples in areas of hybridization, such as the United States. It will not be possible to determine if the pooled sample contains a mixture of pure Cx. pipiens and pure Cx. quinquefasciatus, and/or their hybrids. Also, in the United States, Cx. restuans, Cx. salinarius, and Cx. pipiens complex specimens are commonly found in pooled samples of potential West Nile virus vectors.32–34 In those instances, we recommend that the assay in Crabtree and others8 be used in conjunction with our assay.

In conclusion, we have developed assays that identify the members of the Cx. pipiens complex and other sibling species across several geographic regions worldwide, and that also detect introgression between Cx. pipiens and Cx. quinquefasciatus. The assays involve a single PCR per specimen, and our extensive population level examination for most of the species shows they consistently generate unique fragments easily resolved by electrophoresis on agarose gels. This methodology will be helpful to researchers and will aid vector control programs by facilitating the rapid and reliable identification of local mosquitoes.

Table 1

Populations of Culex species used in this study and their source

SpeciesOriginSource*
*Dr. Michael Service, Liverpool School of Tropical Medicine, Liverpool, United Kingdom; Dr. Harry Savage, Centers for Disease Control and Prevention, Fort Collins, CO; Dr. Chris Curtis, London School of Hygiene and Tropical Medicine, London, United Kingdom; Dr. Francis Schaffner, Adege, Entente Interdépartmentale pour la Démoustication Méditerranée, Montpellier, France; Mike Reddy, Harvard School of Public Health, Boston, MA; Dr. Laura Kramer, Wadsworth Center, Albany, NY: Dr. Sheryl Yamamoto, Sacramento-Yolo Mosquito and Vector Control, Elk Grove, CA; Dr. Motoyoshi Mogi, Saga Medical School, Saga, Japan; Dr. Mike Carroll, New Orleans Mosquito Control, New Orleans, LA: Jo Kent and Craig Williams, University of South Australia, Adelaide, Australia; Dr. George O’Meara, Florida Medical Entomology Laboratory, Vero Beach, FL; Lloyd Shimoda, Hawaii Vector Control, Hilo, MI; Dr. Shirley Luckhart, Virginia Tech, Blacksburg, VA; Dr. Jan Conn, Wadsworth Center, Albany, NY; Dr. Ichiro Miyagi, University of the Ryukyus, Okinawa, Japan; Felipe Noguera, Estacion Biologica de Chamela, Jalisco, Mexico; Mark Bullians, AgriQuality New Zealand, Auckland, New Zealand; Dr. Bruce Harrison, Dept. of Environment and Natural Resources, Winston-Salem, NC; Dr. Chris Evans, Department of Health and Environmental Control, Columbia, SC; Dr. Michael Womack, Macon State College, Macon, GA; Dr. Min Lee Cheng, West Valley Mosquito and Vector Control District, Chino, CA; Dr. Anton Cornel, University of California, Davis, CA; Dr. Colin Malcolm, University of London, London, United Kingdom.
†Populations from which the acetylcholinesterase locus was sequenced (from 3 specimens). A genitalia analysis12 was performed prior to DNA extraction to confirm species identification so only male specimens were sequenced. The other populations, which included males and females, were used to examine the diagnostic ability of the various assays.
Cx. pipiensLiverpool, England†M.W. Service/H. Savage
Wedmore, EnglandC. Curtis
Loughborough, EnglandC. Curtis
Alsace, FranceF. Schaffner
Boston, MAM. Reddy
Albany, NYL. Kramer
Portland, ORS. Yamamoto
Saga, Japan†M. Mogi
Amman, Jordan†M. Carroll
Point Willunga, Australia†J. Kent/C. Williams
Pryca, Spain†M. Mogi
Cx. quinquefasciatusNew Orleans, LA†M. Carroll
Archer, FLG. O’Meara
Maui, HI†D. Fonseca/L. Shimoda
Kisumu, KenyaS. Luikart/D. Fonseca
Makurdi, Nigeria†J. Conn
Kupang, West Timor†M. Mogi
Okinawa, Japan†I. Miyagi
Jalisco, MexicoF. Noguera
Auckland, New Zealand†M. Bullians
Adelaide, Australia†J. Kent/C. Williams
Cx. pipiens-Cx. quinquefasciatus hybridsSuitland, MD†D. Fonseca
Walkertown, NCB. Harrison
Greenville, SCC. Evans
Blairsville, GAM. Womack
Valdosta, GAM. Womack
Hickman, CAS. Yamamoto
Marysville, CAS. Yamamoto
Buttonwillow, CAS. Yamamoto
Chino, CAM.L. Cheng
Shashta, CA†A. Cornell
Cx. pipiens pallensChoushouji, Saga, Japan†M. Mogi
Tenman, Saga, Japan†M. Mogi
Cx. torrentiumLiverpool, England†M.W. Service/H. Savage
Middleton, ScotlandC. Malcolm
Manor Farm, ScotlandC. Malcolm
Glenochil, ScotlandC. Malcolm
Alsace, FranceF. Schaffner
Cx. australicusEmbarka, Australia†C. Williams
Cx. pervigilansAuckland, New Zealand†M. Bullians
Table 2

Diagnostic primers for species in the Culex pipiens complex and its sibling species*

PrimerPrimer sequence
*The first five primers are all forward primers and need to be run with B1246s as the reverse primer.
ACEaus5′-CTTGTGGTGATTTAGTTGTTCGG-3′
ACEquin5′-CCTTCTTGAATGGCTGTGGCA-3′
ACEpall5′-ATGGTGGAGACGCATGACG-3′
ACEpip5′-GGAAACAACGACGTATGTACT-3′
ACEtorr5′-TGCCTGTGCTACCAGTGATGTT-3′
B1246s5′-TGGAGCCTCCTCTTCACGG-3′
Table 3

Summary of the main geographic species groups and the appropriate primer multiplexes*

RegionSpecies present and approximate size of diagnostic fragmentsPrimers to usePrimer concentration (μM)
*bp = basepairs.
Africa, AmericasCulex (Culex) pipiens (610 bp)ACEpip0.2
Cx. (Cx.) quinquefasciatus (274 bp)ACEquin0.4
Hybrids (610-bp and 274 bp)B1246s0.4
EurasiaCx (Cx.) pipiens (610 bp)ACEpip0.1
Cx. (Cx.) torrentium (416 bp)ACEtorr0.1
B1246s0.2
East AsiaCx. (Cx.) pipiens (610 bp)ACEpip0.1
Cx. (Cx.) quinquefasciatus (274 bp)ACEquin0.1
Cx. (Cx.) p. pallens (478 bp)ACEpall0.1
Putative hybrids (478 bp and 274 bp)B1246s0.3
AustraliaCx. (Cx.) pipiens (610 bp)ACEpip0.2
Cx. (Cx.) quinquefasciatus (274 bp)ACEquin0.2
Cx. (Cx.) australicus (437 bp)ACEaus0.2
B1246s0.6
New ZealandCx. (Cx.) quinquefasciatus (274 bp)ACEpip0.1
Cx. (Cx.) pervigilans (668 bp)ACEquin0.1
B1246s0.2
Figure 1.
Figure 1.

Distribution of the Culex pipiens complex and its sibling species based on maps of Dahl,35 Belkin,36 Mattingly and others,37 and available literature.12,38,39 Light gray = Cx. pipiens; black = Cx. quinquefasciatus; dark gray = overlapping ranges of Cx. pipiens and Cx. quinquefasciatus; region marked by dotted line = Cx. torrentium; region marked by solid line = Cx. australicus; region marked by dashed line = Cx. pipiens pallens; New Zealand marked by dotted and dashed line = Cx. pervigilans.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 4; 10.4269/ajtmh.2004.70.339

Figure 2.
Figure 2.

Polymorphisms across taxa and the relative position of the unique primers. Each species is represented by the inclusive consensus (shaded regions) of all its cloned sequences (GenBank accession #AY497523-26). Position 1 corresponds to the 5′ end of exon 2 (position 1 in Figure 2A in Malcolm and others25). Position 203 corresponds to the first basepair at a Sca I-cute site.10 B = C/G/T; W = A/T; R = A/G; Y = C/T; K = G/T; S = C/G; M = A/C. Ace = acetylcholinesterase; pip = pipiens; pal = pallens; aus; australicus; torr = torrentium; quin = quinquefasciatus. fasciatus,

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 4; 10.4269/ajtmh.2004.70.339

Figure 3.
Figure 3.

Fragments amplified using the regional multiplexes of primers. A, lane 2= Culex pipiens; lane 3 = Cx. quinquefasciatus; lane 4 = Cx. pervigilans; B, lane 6 = Cx. pipiens; lane 7 = Cx. quinquefasciatus; lane 8 = Cx. australicus; C, lane 10 = Cx. pipiens, lane 11, Cx. torrentium; D, lane 13 = Cx. pipiens; lane 14 = Cx. quinquefasciatus; lane 15 = Cx. pipiens × Cx. quinquefasciatus hybrid; lane 16 = Cx. p. pallens or putative pallens × Cx. quinquefasciatus hybrid. Lanes 1, 5, 9, 12, and 17 are size standards (100-basepair ladder; New England Biolabs, Beverly, MA).

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 4; 10.4269/ajtmh.2004.70.339

Figure 4.
Figure 4.

Correlation between the frequency of Culex pipiens acetylcholinesterase (ACE) alleles and the average frequency of the “pipiens” signature (probability of ancestry from Cx. pipiens) based on a panel of eight microsatellite loci across 10 populations from the United States.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 70, 4; 10.4269/ajtmh.2004.70.339

Authors’ addresses: Julie L. Smith, Academy of Natural Sciences, 1900 Benjamin Franklin Parkway, Philadelphia, PA 19103, E-mail: jsmith@acnatsci. Dina M. Fonseca, Academy of Natural Sciences, 1900 Benjamin Franklin Parkway, Philadelphia, PA 19103, E-mail: fonseca@acnatsci.org.

Acknowledgments: We are indebted to all the collaborators for providing specimens. We thank Katherine Unger and Nusha Keyghobadi for their editorial comments and Tapan Ganguly and the DNA Sequencing Facility, University of Pennsylvania (Philadelphia, PA) for technical assistance.

Financial support: This research was partially funded by a National Research Council Associateship through the Walter Reed Army Institute of Research to Dina M. Fonseca, by Centers for Disease Control and Prevention/National Institutes of Health (NIH) grant U50/ CCU220532, and by NIH grant 1R01GM063258.

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