• View in gallery

    Linkage groups for Anopheles albimanus. The original linkage map for chromosome 2 based on morphological and isoenzyme markers is shown on the left.5 The cytological position of the morphological markers used in the present study is given. For linkage groups 1 (chromosome 3) and X, no order or genetic distance is intended. Distances are in cM.

  • 1

    Arredondo-Jimenez JI, Brown DN, Rodriguez MH, Villareal C, Loyola EG, Frederickson CE, 1992. Tests for the existence of genetic determination or conditioning in host selection by Anopheles albimanus (Diptera: Culicidae). J Med Entomol 29 :894–897.

    • Search Google Scholar
    • Export Citation
  • 2

    Narang SK, Seawright JA, Suarez MF, 1991. Genetic structure of natural populations of Anopheles albimanus in Colombia. J Am Mosq Control Assoc 7 :437–445.

    • Search Google Scholar
    • Export Citation
  • 3

    De Merida AM, De Mata MP, Molina E, Porter CH, Black IV WC, 1995. Variation in ribosomal DNA intergenic spacers among populations of Anopheles albimanus in South and Central America. Am J Trop Med Hyg 53 :469–477.

    • Search Google Scholar
    • Export Citation
  • 4

    Molina-Cruz A, De Merida AMP, Mills K, Rodriguez F, Schoua C, Yurrita MM, Molina E, Palmieri M, Black IV WC, 2004. Gene flow among Anopheles albimanus populations in Central America, South America, and the Caribbean assessed by microsatellites and mitochondrial DNA. Am J Trop Med Hyg 71 :350–359.

    • Search Google Scholar
    • Export Citation
  • 5

    Narang SK, Seawright JA, 1989. Linkage map of the mosquito (Anopheles albimanus) (2N=6). Genetic Maps. Locus Maps of Complex Genomes, ed. SJ O’Brien, Book 3, 3.269–3.272. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

  • 6

    Fischer D, Bachmann K, 1998. Microsatellite enrichment in organisms with large genomes (Allium cepa L.). Biotechniques 24 :796–800.

  • 7

    Rozen S, Skaletsky H, 2000. Primer3 on the WWW for general users and for biologist programmers. Methods Mol Biol 132 :365–386.

  • 8

    Benedict MQ, 1997. Care and maintenance of anopheline mosquito colonies. Molecular Biology of Insect Disease Vectors: A Methods Manual, ed. JM Crampton, CB Beard, C Louis, 1:3–12. Dordrect, Netherlands: Chapman and Hall.

  • 9

    Savage KE, Lowe RE, 1971. A one-piece aluminum cage designed for adult mosquitoes. Mosq News 31 :111–112.

  • 10

    Brogdon WG, McAllister JC, Corwin AM, Cordon-Rosales C, 1999. Independent selection of multiple mechanisms for pyre-throid resistance in Guatemalan Anopheles albimanus (Diptera: Culicidae). J Econ Entomol 92 :298–302.

    • Search Google Scholar
    • Export Citation
  • 11

    Benedict MQ, Seawright JA, Anthony DW, Avery SW, 1979. Ebony, a semidominant lethal mutant in the mosquito Anopheles albimanus. Can J Genet Cytol 21 :193–200.

    • Search Google Scholar
    • Export Citation
  • 12

    Georghiou GP, 1972. Studies on resistance to carbamate and organophosphorous insecticides in Anopheles albimanus. Am J Trop Med Hyg 21 :797–806.

    • Search Google Scholar
    • Export Citation
  • 13

    Georghiou GP, Gidden FE, Cameron JW, 1967. A Stripe character in Anopheles albimanus (Diptera: Culicidae). Ann Entomol Soc Am 60 :323–328.

    • Search Google Scholar
    • Export Citation
  • 14

    Collins FH, Mendez MA, Rasmussen MO, Mehaffey PC, Besansky NJ, Finnerty V, 1987. A ribosomal RNA gene probe differentiates member species of the Anopheles gambiae complex. Am J Trop Med Hyg 37 :37–41.

    • Search Google Scholar
    • Export Citation
  • 15

    Lander ES, Green P, Abrahamson J, Barlow A, Daly MJ, Lincoln SE, Newberg LA, 1987. MAPMAKER: an interactive computer package for constructing primary genetic linkage maps of experimental and natural populations. Genomics 1 :174–181.

    • Search Google Scholar
    • Export Citation
  • 16

    Stam P, 1993. Construction of integrated genetic-linkage maps by means of a new computer package—Joinmap. Plant J 3 :739–744.

  • 17

    Penilla RP, Rodriguez AD, Hemingway J, Torres JL, Arredondo-Jimenez JI, Rodriguez MH, 1998. Resistance management strategies in malaria vector mosquito control. Baseline data for a large-scale field trial against Anopheles albimanus in Mexico. J Med Entomol 12 :217–233.

    • Search Google Scholar
    • Export Citation
  • 18

    Seawright JA, Benedict MQ, Narang S, 1984. Use of deficiencies for mapping four mutant loci on the salivary gland chromosomes of Anopheles albimanus. Mosq News 44 :568–572.

    • Search Google Scholar
    • Export Citation
  • 19

    Zheng L, Benedict MQ, Cornel AJ, Collins FH, Kafatos FC, 1996. An integrated genetic map of the African human malaria vector mosquito, Anopheles gambiae. Genetics 143 :941–952.

    • Search Google Scholar
    • Export Citation
  • 20

    Cornel AJ, Collins FH, 2000. Maintenance of chromosome arm integrity between two Anopheles mosquito subgenera. J Hered 91 :364–370.

  • 21

    Seawright JA, Benedict MQ, Narang S, 1985. Color mutants in Anopheles albimanus (Diptera: Culicidae). Ann Entomol Soc Am 78 :177–181.

 
 
 

 

 
 
 

 

 

 

 

 

 

Towards a Genetic Map for Anopheles albimanus: Identification of Microsatellite Markers and a Preliminary Linkage Map for Chromosome 2

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  • 1 Centro Regional de Investigación en Salud Pública/Instituto Nacional de Salud Pública, Tapachula, Chiapas, México; Liverpool School of Tropical Medicine, Liverpool, United Kingdom; Center for Health Studies, Universidad del Valle de Guatemala; Centers for Disease Control and Prevention, Atlanta, Georgia; Department of Microbiology, Immunology and Pathology, Colorado State University, Fort Collins, Colorado

Fifty microsatellite loci were identified in the malaria vector Anopheles albimanus. Markers segregating in F2 progeny of crosses between laboratory strains of An. albimanus were used to construct a preliminary genetic map. More than 300 progeny were genotyped, but the resolution of the map was limited by the lack of polymorphisms in the microsatellite alleles. A robust linkage map for chromosome 2 was established, and additional markers were assigned to the third and X chromosomes by linkage to morphological markers of known physical location. Additional non-informative microsatellite sequences are provided including some showing similarity to those of An. gambiae. This study significantly increases the number of genetic markers available for An. albimanus and provides useful tools for population genetics and genetic mapping studies in this important malaria vector.

INTRODUCTION

Anopheles albimanus is widely distributed throughout the tropics and subtropics of the Americas, extending from the southern United States to northern Peru and the Caribbean Islands. It is the primary coastal vector of malaria in these regions. Unlike many anophelines, which are members of species complexes, there is no evidence for cryptic species of this vector. 1,2 Nevertheless, An. albimanus populations differ in their host preference and vectorial capacity, and barriers to gene flow between Atlantic and Pacific populations have been detected. 3,4 The power of genetic analyses has been limited by a lack of neutral markers for An. albimanus. A linkage map consisting of biochemical and morphological markers has been reported,5 but this contained no microsatellite or SNP markers, and few of the markers are extant for developing the map further.

In this study we report the sequences of 50 new microsatellite markers that will facilitate detailed population genetic studies of this malaria vector. We were able to use a small number of the existing biochemical markers, whose cytological location had been determined,5 to anchor the microsatellite-based map to chromosome arms of An. albimanus. In addition, we have constructed a well-supported low-resolution map for chromosome 2 of An. albimanus. The development of this integrated genetic and cytogenetic map will facilitate molecular and population genetic studies in this species.

MATERIALS AND METHODS

Identification of microsatellite markers.

Clones were identified by two methods: A genomic plasmid library was created by digesting c-purified An. albimanus deoxyribonucleic acid (DNA) with Sau 3AI and ligating it into phosphatased Bam HI-digested pUC18 plasmids. Colonies were blotted onto nitrocellulose membranes and probed with digoxigenin-labeled microsatellite primers including various di-, tri-, and tetranucleotide repeats (GC10, TA10, ATG8, CTG8, AAC8, GAA8, TCAG6, TGAA6, CTGA6, CAGA6). Hybridization-positive clones were purified and sequenced using M13 forward and/or reverse primers. Additional loci were identified using a procedure to enrich the library for microsatellite-containing plasmids.6 Primers to amplify regions containing microsatellites were designed using the program Primer37 or by eye. DNA sequencing was performed on an Applied Biosystems ABI 377 sequencer (Life Technologies, Carlsbad, CA) using BigDye reagents and standard M13 forward and reverse primers.

Mosquito culture and genetic crosses.

Mosquitoes were cultured under conditions described by Benedict.8 Briefly, crosses were performed in aluminum and gauze cages9 or pint (approximately 1/2 L) paper cups. Two types of genetic crosses were conducted. In the first, a stock previously selected by insecticide exposure for high levels of oxidase and esterase (HIOX/HIEST) 10 was crossed with PREBST+. The latter resulted from genetic mapping crosses reported elsewhere, 11 and had high frequencies of the dominant alleles of propoxur resistance, 7,12 ebony,11 and Stripe.13 Individual females were separated after blood feeding for oviposition. The F1 families were reared separately and intercrossed to produce the F2 generation. L3 and L4 larvae were scored for the two morphological markers, ebony and Stripe, or were cultured to the pupal stage and scored for sex and, in some families, Stripe, before being frozen for later DNA extraction. In the second set of crosses, a stock originating from Santa Tecla, El Salvador (STECLA) was crossed with PREBST+. STECLA is pure-breeding for the recessive alleles of the PREBST+ markers described above. Individual F1 siblings were crossed to produce F2 families. These were raised to the pupal stage and scored for sex and, in some families, the presence of the Stripe phenotype, before being frozen for later DNA extraction.

Four F2 families were selected for genotyping, each originating from a distinct parental pair. For crosses resulting in multiple isofemale lines, the families with the largest number of progeny were selected. One of these, family AJ-1, originated from the HIOX/HIEST × PREBST+ cross, and the remaining three (families 108N, 121D and 129D) were from crosses between STECLA and PREBST+.

Analysis of Microsatellites.

Genomic DNA was extracted from the parents and progeny of the single pair crosses as described previously. 14 The microsatellite loci were amplified using fluorescently labeled primers with a reaction cycle of 95°C for 5 minutes followed by 35 cycles of 94°C for 30 seconds, 55°C for 30 seconds, and 72°C for 30 seconds. The polymerase chain reaction (PCR) products were resolved by capillary electrophoresis using either an ABI 377 or a CEQ8000 (Beckman Coulter Inc., Fullerton, CA) automatic sequencer. A size standard was included in each lane and allele sizes were determined using the Applied Biosystems GeneScan or CEQ8000 (Beckman Coulter Inc., Fullterton, CA) software. Initially the parental and F1 generations were genotyped at 50 loci to identify informative markers for each family. The F2 progeny were then scored for each of these informative markers and χ2 goodness of fit tests were performed to identify markers not segregating according to Hardy Weinberg equilibrium. MapMaker/Exp V3.0 15 and JoinMap V3.0 16 were used to identify linkage groups. The nomenclature used for the markers and reported in the GenBank entries is the pattern “Aalbi-CA- #” where C is the chromosome if assigned (X, Y, 2, 3) or U if unassigned, A is the arm if assigned or N if unknown, and # is an arbitrary identification number. Throughout this manuscript, we will refer to the markers only by their number. Eleven adult An. albimanus collected from cattle corrals from six villages from coastal Chiapas province, Mexico between May 1997 and June 1998 as part of a large scale insecticide resistance management trial 17 were also used to further assess the polymorphism of some of the microsatellite markers.

RESULTS

Identification of An. albimanus microsatellite markers.

Ninety-six sequences were identified by screening a plasmid library for repetitive DNA. The inserts were sequenced and 12 were discarded either because they contained less than four repeat units or because the repeat was at the end of the clone, and hence flanking primers could not be designed. Primers were designed to amplify the microsatellite repeats in the remaining 84 clones. Some plasmids contained more than one repeat, and in these cases, primers were designed to independently amplify each microsatellite repeat. Each primer pair was tested for amplification using genomic DNA extracted from three laboratory strains of An. albimanus. Products of the expected size were obtained for 75 loci. Fifty markers were selected for the present study. Table 1 summarizes the procedure for the selection of microsatellite markers used in this study.

Markers were tested for polymorphism using laboratory strains and field collected mosquitoes from six sites in Chiapas, Mexico. More than one allele was detected for 36 of the microsatellites. The primer sequences, expected allele size, and presence or absence of polymorphism for each of the 50 markers are shown in Table 2.

Linkage analysis.

Four F2 families were selected for genotyping. Informative markers were identified by scoring the F0 parents of the crosses and the results confirmed by genotyping of the F1 generation. A total of 303 F2 progeny were analyzed from these four families (94 from family AJ-1, 90 from 108N, 80 from 121D, and 39 from 129D). The number of informative markers ranged from 14 (family 129D) to 23 (family AJ-1).

Sex-linked markers.

All markers were analyzed for Hardy Weinberg equilibrium assuming a segregation ratio of 1:2:1 (homozygous parental: heterozygous: homozygous parental) in the F2 progeny. Those markers that significantly deviated from the expected ratios were re-analyzed for sex linkage. Only females are informative for X-linked markers and therefore, for those families where the sex of the progeny was known, the males were excluded from this analysis. Two candidate X linked markers were identified. Marker 0086, which was informative in three families, segregated in a 1:1 ratio of heterozygous: homozygous for the maternal genotype in the female progeny (P = 0.48) suggesting that this marker is located on the X chromosome. The white locus, which has previously been localized to the X chromosome in Anopheles albimanus, also segregated in the ratio expected for a sex-linked marker. This marker was only informative for family AJ-1 whose sex was not determined. Assuming this family has an equal number of males and females in the F2 generation, a ratio of 2:1:1 (homozygous maternal: heterozygous: homozygous paternal) would be expected. The observed ratio for the white locus in this family matched this expected distribution (P = 0.184). Unfortunately, as none of the families analyzed were informative for both 0086 and white, it was not possible to test for linkage between these putative X-linked markers.

Autosomal markers.

Of the remaining 23 informative loci, five loci showed segregation disorders in one or more families (Table 3). One of these (0056) never segregated as expected and was therefore removed from the subsequent analysis. In all other cases markers showing segregation disorders were removed from selected families.

Initially each family was analyzed separately, using Map-Maker/Exp with a minimum logarithm of the odds (LOD) score of 3.0, to identify putative linkage groups. Three distinct linkage groups (linkage group 1–3) were identified within each of three of the four families. However, in the largest family, AJ-1, a single linkage group, consisting of 15 markers was obtained. The AJ-1 linkage group contained markers from the initial linkage groups 2 and 3. We therefore used Joinmap to integrate the maps from the different families. This confirmed the linkage between groups 2 and 3. This large linkage group, which will hereafter be referred to as linkage group 2, consisted of 14 markers with defined order spanning 124 cM (Figure 1).

Linkage group 1, consisting solely of markers 0143 and 0057, was unaffected by the pooling of the data. This linkage group was not detected in family AJ-1 as marker 0143 was non-informative in this family.

Unassigned markers.

Seven markers were unlinked in the pooled analysis. Three of these, 0020, 0022, and 0087, were associated with linkage group 2 in the single family analysis but were only loosely associated with linkage group 2 in the integrated map. The remaining four markers (0078, 0101, 0129, and 0077) consistently could not be assigned.

Anchoring of linkage groups to An. albimanus chromosomes.

One family, AJ-1, was informative for the semi-dominant morphological marker, ebony, that had been mapped using deficiencies to division 20A on the left arm of chromosome 2. 18 The F1 parents were both heterozygous at this locus, and the F2 progeny segregated in the expected 1:2:1 ratio. The ebony locus was genetically mapped between markers 0108 and 0014, thereby anchoring linkage group 2 to the second chromosome.

Stripe is a polymorphic gene whose various alleles are common in wild mosquitoes. It has been located by deletion analysis to chromosome 3R in division 33B. 18 Two families, AJ-1 and 108N, were informative for this second morphological marker. The female parent of both families was homozygous for the dominant allele whereas the male carried the recessive allele. As heterozygotes could not be distinguished from homozygotes, the expected ratio of Stripe:wild type is 3:1. This ratio was observed in family AJ-1 but not in family 108N (P = 0.024). When the Stripe locus was included as a genetic marker, weak linkage was found to linkage group 1 in family 108N (LOD 2.0) possibly indicating that this linkage group is found on chromosome 3. No linkage was observed in family AJ-1, but this is perhaps expected as this family was non-informative for the markers putatively located on chromosome 3.

DISCUSSION

The present manuscript catalogues a novel set of microsatellite markers for An. albimanus and presents a preliminary genetic map for chromosome 2. Two markers were also assigned to each of the additional chromosomes (X and 3) by linkage to physically mapped morphological markers: 32 microsatellite markers could not be assigned to any chromosome largely due to lack of polymorphism in the parents of the crosses. The bias towards informative markers on chromosome 2 was unexpected, but a similar dichotomy in the distribution of randomly identified microsatellite markers between the autosomal chromosomes was also observed in An. gambiae.19 We are unable to offer an explanation for why this is so. Similarly perhaps, during the period in which fairly intense screens for morphological mutations were performed in An. albimanus, morphological markers were found on all arms except 3L though numerous enzyme polymorphisms were found on that arm.5 We feel certain that further surveys for both would detect markers on chromosome 3 and that this is probably simply a sample size effect which is exaggerated by the need for several markers to be clearly linked.

We compared the size and orientation of the chromosome 2 linkage group as determined here with that estimated previously.5 Because no chromosome rearrangements exist in any of the stocks used in the mapping crosses (nor have any been observed in spite of extensive examination of natural populations), and recombination rates between males and females are the same, we determined that the minimum length of chromosome 2 is 124 cM. The previously published map estimated a length of 168 cM for linkage group 2.5 Although the calculations of lengths may not be directly comparable, the map that we present appears to contain most of the linkage group.

The use of the marker ebony that is common to both our microsatellite and previous linkage maps allows us to give a probable orientation for the chromosome 2 map as shown in Figure 1 and which we reason as follows: The total distance from ebony to the end of 2L was estimated to be approximately 40 cM by its distance from the sub-telomeric marker brown larva (bw).20 The most distant markers from ebony in our study are 0025 (28 cM distant) and 0109 (96 cM distant). Unless the estimate of the location of bw was grossly underestimated relative to the telomere, there is insufficient recombination distance for a marker to be located on 2L at a distance as great as that of 0109. In contrast, approximately 109 cM exists between ebony and the end of 2R. Therefore, the marker-deficient area of our chromosome 2 map likely lies primarily at the telomeric end of 2R.

In situ hybridizations have previously demonstrated that within arms, the gene compositions of An. gambiae and An. albimanus are largely conserved. 20 However, the arms are arranged in different chromosomes: 2R of An. gambiae and An. albimanus are syntenic, however 3R of A. gambiae is syntenic with 2L of An. albimanus. This conclusion is supported by the occurrence of a BLAST hit of the An. albimanus microsatellite 0113, located on chromosome 2R in An. albimanus, on An gambiae division 12E, chromosome 2R. Furthermore, 0128, located on chromosome 2L in An. albimanus has a BLAST hit on division 29C, chromosome 3R of An. gambiae. Further confirmation of synteny can be found by comparing the location of isozymes mapped previously 21 with their map location in An. gambiae. Propoxur resistance (insensitive AChE), 6-phosphogluconate-dehydrogenase-2, phosphoglucomutase, aconitase, and hexokinase all map to the chromosomes previously identified as syntenic.

Because the genome of An. gambiae has been sequenced, it provides a useful source of information with which to expand our conclusions. Recently, further sequencing has identified over 100 additional sequences containing repeat elements (Table S1, Genbank numbers are within the file). The sequence and synteny of chromosome arms has allowed us to identify 68 untested microsatellites from this new sequence data set that have similarity to An. gambiae sequences (by BLAST similarity), and their probable location in the genome of An. albimanus will provide a useful starting point for those wishing to add loci to the existing map. In addition, single nucleotide polymorphisms segregating in F2 progeny from the same crosses are being genotyped in the Centro Regional de Investigación en Salud Pública en Chiapas leading to improved resolution of the existing map.

Genetic maps are limited by the number of progeny and the number of markers. In this case, the limited number of informative markers clearly restricts the resolution of the map. In future studies, genetic crosses of populations at the extremes of the geographical distribution of An. albimanus should increase the number of polymorphic markers and lead to a more densely populated map. Indeed, a further six markers that were non-informative in the genetic crosses were polymorphic in a small number of field-collected mosquitoes analyzed from Chiapas, Mexico.

Table 1

Steps for identifying microsatellite markers in Anopheles albimanus

Table 1
Table 2

Anopheles albimanus microsatellite markers

Table 2
Table 3

Markers showing segregation disorders* in Anopheles albimanus progeny

Table 3
Figure 1.
Figure 1.

Linkage groups for Anopheles albimanus. The original linkage map for chromosome 2 based on morphological and isoenzyme markers is shown on the left.5 The cytological position of the morphological markers used in the present study is given. For linkage groups 1 (chromosome 3) and X, no order or genetic distance is intended. Distances are in cM.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 81, 6; 10.4269/ajtmh.2009.08-0607

*

Address correspondence to Mark Q. Benedict, PO Box 105603, No. 6662, Atlanta, GA 30348. E-mail: MQBenedict@yahoo.com

Note: Supplemental material is available online at www.ajtmh.org.

Financial support: This work was supported by a Wellcome Trust Project Grant 065849/2/01/Z to Patricia Penilla and by a Gorgas Memorial Institute Research Award to Norma Padilla.

Authors’ addresses: R. Patricia Penilla and Américo D. Rodríguez, Centro Regional de Investigación en Salud Pública/Instituto Nacional de Salud Pública, Tapachula, Chiapas, México, E-mails: penilla@insp.mx and americo@insp.mx. Hilary Ranson, John C. Morgan, Keith Steen, Patricia Pignatelli, and Janet Hemmingway, Liverpool School of Tropical Medicine Vector Group, Liverpool School of Tropical Medicine, Pembroke Place, Liverpool, UK, E-mails: hranson@liverpool.ac.uk, jcmorgan@liv.ac.uk, steenk@liv.ac.uk, triciap@liv.ac.uk, and Hemingway@liverpool.ac.uk. Norma Padilla, Center for Health Studies, Universidad del Valle de Guatemala, Vista Hermosa III Guatemala, Guatemala, E-mail: npadilla@gt.cdc.gov. William G. Brogdon, Centers for Disease Control and Prevention, Chamblee, GA, E-mail: WGB1@CDC.gov. William C. Black IV, Department of Microbiology, Immunology and Pathology, Colorado State University, Fort Collins, CO, E-mail: William.Black@ColoState.EDU. Mark Q. Benedict, PO Box 105603, No. 6662, Atlanta, GA 30348, E-mail: MQBenedict@yahoo.com.

REFERENCES

  • 1

    Arredondo-Jimenez JI, Brown DN, Rodriguez MH, Villareal C, Loyola EG, Frederickson CE, 1992. Tests for the existence of genetic determination or conditioning in host selection by Anopheles albimanus (Diptera: Culicidae). J Med Entomol 29 :894–897.

    • Search Google Scholar
    • Export Citation
  • 2

    Narang SK, Seawright JA, Suarez MF, 1991. Genetic structure of natural populations of Anopheles albimanus in Colombia. J Am Mosq Control Assoc 7 :437–445.

    • Search Google Scholar
    • Export Citation
  • 3

    De Merida AM, De Mata MP, Molina E, Porter CH, Black IV WC, 1995. Variation in ribosomal DNA intergenic spacers among populations of Anopheles albimanus in South and Central America. Am J Trop Med Hyg 53 :469–477.

    • Search Google Scholar
    • Export Citation
  • 4

    Molina-Cruz A, De Merida AMP, Mills K, Rodriguez F, Schoua C, Yurrita MM, Molina E, Palmieri M, Black IV WC, 2004. Gene flow among Anopheles albimanus populations in Central America, South America, and the Caribbean assessed by microsatellites and mitochondrial DNA. Am J Trop Med Hyg 71 :350–359.

    • Search Google Scholar
    • Export Citation
  • 5

    Narang SK, Seawright JA, 1989. Linkage map of the mosquito (Anopheles albimanus) (2N=6). Genetic Maps. Locus Maps of Complex Genomes, ed. SJ O’Brien, Book 3, 3.269–3.272. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

  • 6

    Fischer D, Bachmann K, 1998. Microsatellite enrichment in organisms with large genomes (Allium cepa L.). Biotechniques 24 :796–800.

  • 7

    Rozen S, Skaletsky H, 2000. Primer3 on the WWW for general users and for biologist programmers. Methods Mol Biol 132 :365–386.

  • 8

    Benedict MQ, 1997. Care and maintenance of anopheline mosquito colonies. Molecular Biology of Insect Disease Vectors: A Methods Manual, ed. JM Crampton, CB Beard, C Louis, 1:3–12. Dordrect, Netherlands: Chapman and Hall.

  • 9

    Savage KE, Lowe RE, 1971. A one-piece aluminum cage designed for adult mosquitoes. Mosq News 31 :111–112.

  • 10

    Brogdon WG, McAllister JC, Corwin AM, Cordon-Rosales C, 1999. Independent selection of multiple mechanisms for pyre-throid resistance in Guatemalan Anopheles albimanus (Diptera: Culicidae). J Econ Entomol 92 :298–302.

    • Search Google Scholar
    • Export Citation
  • 11

    Benedict MQ, Seawright JA, Anthony DW, Avery SW, 1979. Ebony, a semidominant lethal mutant in the mosquito Anopheles albimanus. Can J Genet Cytol 21 :193–200.

    • Search Google Scholar
    • Export Citation
  • 12

    Georghiou GP, 1972. Studies on resistance to carbamate and organophosphorous insecticides in Anopheles albimanus. Am J Trop Med Hyg 21 :797–806.

    • Search Google Scholar
    • Export Citation
  • 13

    Georghiou GP, Gidden FE, Cameron JW, 1967. A Stripe character in Anopheles albimanus (Diptera: Culicidae). Ann Entomol Soc Am 60 :323–328.

    • Search Google Scholar
    • Export Citation
  • 14

    Collins FH, Mendez MA, Rasmussen MO, Mehaffey PC, Besansky NJ, Finnerty V, 1987. A ribosomal RNA gene probe differentiates member species of the Anopheles gambiae complex. Am J Trop Med Hyg 37 :37–41.

    • Search Google Scholar
    • Export Citation
  • 15

    Lander ES, Green P, Abrahamson J, Barlow A, Daly MJ, Lincoln SE, Newberg LA, 1987. MAPMAKER: an interactive computer package for constructing primary genetic linkage maps of experimental and natural populations. Genomics 1 :174–181.

    • Search Google Scholar
    • Export Citation
  • 16

    Stam P, 1993. Construction of integrated genetic-linkage maps by means of a new computer package—Joinmap. Plant J 3 :739–744.

  • 17

    Penilla RP, Rodriguez AD, Hemingway J, Torres JL, Arredondo-Jimenez JI, Rodriguez MH, 1998. Resistance management strategies in malaria vector mosquito control. Baseline data for a large-scale field trial against Anopheles albimanus in Mexico. J Med Entomol 12 :217–233.

    • Search Google Scholar
    • Export Citation
  • 18

    Seawright JA, Benedict MQ, Narang S, 1984. Use of deficiencies for mapping four mutant loci on the salivary gland chromosomes of Anopheles albimanus. Mosq News 44 :568–572.

    • Search Google Scholar
    • Export Citation
  • 19

    Zheng L, Benedict MQ, Cornel AJ, Collins FH, Kafatos FC, 1996. An integrated genetic map of the African human malaria vector mosquito, Anopheles gambiae. Genetics 143 :941–952.

    • Search Google Scholar
    • Export Citation
  • 20

    Cornel AJ, Collins FH, 2000. Maintenance of chromosome arm integrity between two Anopheles mosquito subgenera. J Hered 91 :364–370.

  • 21

    Seawright JA, Benedict MQ, Narang S, 1985. Color mutants in Anopheles albimanus (Diptera: Culicidae). Ann Entomol Soc Am 78 :177–181.

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