Differential Plasmodium falciparum infection of Anopheles gambiae s.s. molecular and chromosomal forms in Mali
© Trout Fryxell et al.; licensee BioMed Central Ltd. 2012
Received: 25 January 2012
Accepted: 27 April 2012
Published: 27 April 2012
Anopheles gambiae sensu stricto (s.s.) is a primary vector of Plasmodium falciparum in sub-Saharan Africa. Although some physiological differences among molecular and chromosomal forms of this species have been demonstrated, the relative susceptibility to malaria parasite infection among them has not been unequivocally shown. The objective of this study was to investigate P. falciparum circumsporozoite protein infection (CSP) positivity among An. gambiae s.s. chromosomal and molecular forms.
Wild An. gambiae from two sites Kela (n = 464) and Sidarebougou (n = 266) in Mali were screened for the presence of P. falciparum CSP using an enzyme-linked immunosorbent assay (ELISA). Samples were then identified to molecular form using multiple PCR diagnostics (n = 713) and chromosomal form using chromosomal karyotyping (n = 419).
Of 730 An. gambiae sensu lato (s.l.) mosquitoes, 89 (12.2%) were CSP ELISA positive. The percentage of positive mosquitoes varied by site: 52 (11.2%) in Kela and 37 (13.9%) in Sidarebougou. Eighty-seven of the positive mosquitoes were identified to molecular form and they consisted of nine Anopheles arabiensis (21.4%), 46 S (10.9%), 31 M (12.8%), and one MS hybrid (14.3%). Sixty of the positive mosquitoes were identified to chromosomal form and they consisted of five An. arabiensis (20.0%), 21 Savanna (15.1%), 21 Mopti (30.4%), 11 Bamako (9.2%), and two hybrids (20.0%).
In this collection, the prevalence of P. falciparum infection in the M form was equivalent to infection in the S form (no molecular form differential infection). There was a significant differential infection by chromosomal form such that, P. falciparum infection was more prevalent in the Mopti chromosomal forms than in the Bamako or Savanna forms; the Mopti form was also the most underrepresented in the collection. Continued research on the differential P. falciparum infection of An. gambiae s.s. chromosomal and molecular forms may suggest that Plasmodium – An. gambiae interactions play a role in malaria transmission.
Several members of the Anopheles gambiae complex are vectors of human malaria parasites, including Plasmodium falciparum, the species of greatest public health importance in Sub-Saharan Africa. The An. gambiae complex consists of at least seven member species and subspecies [1–3]. Anopheles gambiae sensu stricto (s.s.) is further divided into chromosomal and molecular forms . Molecular studies of the ribosomal DNA region on the X chromosome revealed a fixed difference between populations of An. gambiae that is the basis of the sub-division into the M and S molecular forms . The M and S forms are assortatively mating discreet forms [5–7] that are hypothesized to be undergoing ecotypic speciation due to different larval habitat adaptation [8–10]. Anopheles gambiae are also divided into chromosomal forms based on the arrangements of 5 paracentric inversions on 2R (j, b,c, u and d) and one on 2 L (a), which define Mopti from Savanna from Bamako forms [3, 11]. In Mali, the M form generally associates with the Mopti chromosomal form and the S form with the Savanna and Bamako forms . In some locations in Africa, including Mali, the presence of significant deficiencies in certain inversion heterozygotes suggests there are barriers to gene flow among the chromosomal forms [11, 13–16].
Genetic polymorphisms within An. gambiae have been associated with adaptation to different local environments [17–21]. For example, a number of genetic polymorphisms are associated with tolerance of arid conditions . There is also significant interest in genotypic variation associated with malaria parasite infectivity. Plasmodium falciparum circumsporozoite protein (CSP) positivity in An. gambiae was significantly greater in mosquitoes with isozyme allotypes Mpi 130/130 and Acp 110/100 . In addition, 2La/a individuals were nearly twice as likely to be positive than 2La-/a-. Quantitative trait loci associated with resistance to parasite development have also been identified on chromosome 2 L . Chromosome 2 L includes APL1, and chromosome 3 L includes TEP1r both genes are thought to have anti-parasitic properties. Genome-wide transcriptome analyses of An. gambiae have identified transcript expression patterns that are significantly associated with malaria parasite and bacterial infection [28–31]. Further, some transcription patterns and infection-associated sequence polymorphisms have been associated with An. gambiae laboratory and field-collected M and S molecular forms [32, 33]. Three single nucleotide polymorphisms (SNPs) in immune signaling genes of Malian An. gambiae were significantly associated with natural P. falciparum infection . These SNPs are predicted to alter the structure and function of the encoded proteins and, therefore, alter refractoriness and susceptibility to P. falciparum infection . Additionally, population-specific SNPs associated with either the M or S molecular forms were associated with P. falciparum infection, indicating potential differential immune responses of the two molecular forms to parasite infection . Recently, a cryptic subgroup of An. gambiae was identified in the Sudan-Savanna zone as susceptible to P. falciparum.
Reports of genetic, proteomic, and genomic differences in An. gambiae s.s. suggest, by extension, that susceptibility to P. falciparum infection varies among different molecular forms and populations of An. gambiae[25, 27, 33, 35]. This study examined in more detail the hypothesis that P. falciparum infection prevalence of An. gambiae s.s. differs among molecular and chromosomal forms in Mali. Natural P. falciparum infection levels were compared among An. gambiae molecular and chromosomal forms in two villages in Mali where these forms occur in sympatry. Inversion-specific correlations between P. falciparum infection and standard, heterozygous, and homozygous chromosomal arrangements  were also investigated.
Adult An. gambiae mosquitoes were collected in October 2009 from the villages of Kela (11.88683 N, -8.44744 W), and Sidarebougou (11.4568 N,–5.7323 W) joined with Kolayerebougou (11.4563 N,–5.746 W) in Mali. Resting mosquitoes were collected in the morning via mouth aspirators from inside homes. Mosquitoes were held in cups until they had reached the half-gravid stage. Mosquitoes morphologically identified as An. gambiae s.l. were dissected and separated into head/thorax for P. falciparum CSP ELISA, abdomen/wings/legs for molecular form identification via polymerase chain reaction (PCR), and ovaries for chromosomal form identification via karyotyping. Head/thorax samples were stored in 100% ethanol, while abdomens/legs/wings were stored in 70% ethanol and half-gravid extracted ovaries were stored in modified Carnoy’s solution (3:1 ethanol to glacial acetic acid).
ELISA identification of P. falciparum infection
Lysates of head/thorax samples were assayed using a P. falciparum CSP ELISA [36, 37] according to protocols provided by the Centers for Disease Control and Prevention (Atlanta, Georgia, USA) to identify the sporozoite stage (not gametocyte) and that the P. falciparum protozoan had disseminated across the midgut. The head and thorax, stored in 100% ethanol, were dried prior to tissue lysis. For each ELISA plate, a minimum of two colony-reared An. gambiae mosquitoes (e.g., negative controls) and serial dilutions of P. falciparum monoclonal antibodies (i.e., sensitivity positive controls) were used. The positive control P. falciparum CSP was serially diluted (i.e., 100 pg to 1.5 pg of antigen per 50 μl of blocking buffer) to quantify CSP in field-collected mosquitoes. Samples with absorbance values greater than three times the standard deviations from the mean of the negative control samples on each ELISA plate were designated as “positive” for P. falciparum infection . CSP ELISAs were conducted instead of PCR for molecular detection of P. falciparum to ensure the protozoan had disseminated the midgut and that the protozoan was in the ‘infective’ sporozoite phase.
Identification of species and molecular forms
Abdomen, legs and wings stored in ethanol were ground using a TissueLyser (Qiagen, Valencia, CA, USA), after which DNA was extracted using the BioSprint 96 Bloodkit and automated workstation (Qiagen, Valencia CA, USA). Mosquitoes morphologically identified as An. gambiae s.l. were identified to species  and An. gambiae molecular form identifications were performed on each mosquito [40–42].
Polytene chromosome spreads were prepared from ovarian nurse cells , except that spreads were not stained with lacto-orcein prior to examination. Chromosome banding patterns were visualized using an Olympus BX-50 phase contrast microscope. Species identification and paracentric inversion scoring were accomplished using the polytene chromosome maps for An. gambiae complex and chromosomal forms [3, 11].
Summary statistics, relative abundance of forms, Fisher’s exact tests and two-tailed T-tests were performed in Excel 2007 to determine differences within populations . Where appropriate, we adjusted p-values for multiple comparisons using the Bonferroni correction for an α of 0.05. For the molecular form comparisons, five Chi-square comparisons were conducted (form, village, form x site) that generated a significant p-value less than 0.010. Six comparisons were performed with the chromosomal form data (form, village, form x village) and after the Bonferroni correction, Chi-square comparisons were considered significant if the p-value was less than 0.008. Eighteen comparisons were performed with the karyotype data (arrangement, village, arrangement x village) and, after the Bonferroni correction was applied, Chi-square comparisons were considered significant if the p-value was less than 0.003.
In total, 730 An. gambiae s.l. were analysed, of which 42 (5.8%) were identified as An. arabiensis[3, 39]. Data from Kolayerebougou and Sidarebougou were combined (hereafter Sidarebougou) because they are located within 1.5 km of one another, have similar habitats, and likely represent a single Mendelian population. In Kela, 21.9% (7/25) of Anopheles arabiensis were CSP ELISA positive and 20% (2/10) were CSP ELISA positive in Sidarebougou. Seventeen of the mosquitoes were not identified to molecular form or karyotyped, and two were positive, both from Kela. The remaining 671 mosquitoes were identified as An. gambiae s.s., of which 78 (11.6%) were P. falciparum CSP ELISA positive. Infection prevalence differed for the two collection sites, from 13.9% (35/251) in Sidarebougou and 10.2% (43/420) in Kela, but they were not significantly different (X 2 = 2.101, df = 1, P = 0.1472).
Molecular form and P. Falciparum infection
The M molecular form of Anopheles gambiae s.s. was associated with greater P. falciparum CSP positivity than were other molecular forms in October 2009
No. Pos. (% Pos.)
Total in Mali
Chromosomal form and P. Falciparum infection
The Mopti chromosomal form of Anopheles gambiae s.s. was associated with greater P. falciparum CSP positivity than were other chromosomal forms or hybrids in October 2009
No. Pos. (% Pos.)
Bamako x Savanna
Mopti or Savanna
Bamako x Savanna
Mopti or Savanna
Bamako x Savanna
Mopti or Savanna
Based on site, there were no significant differences between the number of CSP positive and negative Bamako (X 2 = 0.050; df = 1; P = 0.651), Savanna (X 2 = 4.35; df = 1; P = 0.037), or Mopti chromosomal forms (X 2 = 0.029; df = 1; P = 0.864). Within Kela, Savanna and Mopti chromosomal forms were significantly more likely to be positive than the Bamako form (X 2 = 13.4; df = 2; P = 0.001: Bonferroni adjusted P value = 0.0085; Table 2). There were no significant differences in infection prevalence based on chromosomal forms within Sidarebougou (X 2 = 5.47; df = 2; P = 0.065; Table 2). The trend that the most prevalent form from each village (Bamako in Kela and Savanna in Sidarebougou) was least likely to be infected, was noted.
Chromosomal inversions and P. Falciparum infection
Chromosomal inversions in Anopheles gambiae s.s. were not significantly associated with P. falciparum CSP positivity in October 2009 after the Bonferroni correction ( P < 0.003)
No. Positive / No. Screened (% Positive)
32 / 215 (14.9%)
27 / 179 (15.1%)
59 / 335 (15.0%)
0 / 0 (0%)
0 / 0 (0%)
0 / 0 (0%)
1 / 2 (50%)
1 / 14 (7.1%)
2/ 16 (12.5%)
31 / 213 (14.6%
26 / 165 (15.8%)
57 / 378 (15.1%)
X 2 = 0.080; df = 1;
P = 0.777
8 / 83 (9.6%)
1 / 9 (11.1%)
9 / 92 (9.8%)
14 / 90 (15.6%)
7 / 47 (14.9%)
21 / 137 (15.3%)
10 / 42 (23.8%)
19 / 123 (15.4%)
29 / 165 (17.6%)
X 2 = 4.48; df = 2;
P = 0.107
X 2 = 0.125; df = 2;
P = 0.939
X 2 = 2.84; df = 2;
P = 0.242
11 / 46 (23.9%)
19 / 139 (13.7%)
30 / 185 (16.2%)
9 / 46 (19.6%)
5 / 30 (16.7%)
14 / 76 (18.4%)
12 / 123 (9.8%)
3 / 10 (30.0%)
15 / 133 (11.3%)
X 2 = 6.31; df = 2;
P = 0.043
X 2 = 2.01; df = 2;
P = 0.366
X 2 = 2.36; df = 2;
P = 0.307
31 / 210 (14.8%)
24 / 166 (14.5%)
55 / 376 (14.6%)
1 / 5 (20.0%)
3 / 12 (25.0%)
4 / 17 (23.5%)
0 / 0 (0%)
0 / 1 (0.0%)
0 / 1 (0.0%)
X 2 = 0.967; df = 1;
P = 0.326
X 2 = 1.01; df = 1;
P = 0.315
20 / 93(21.5%)
27 / 176 (15.3%)
47 / 269 (17.5%)
0 / 0 (0.0%)
0 / 1 (0%)
0 / 1 (0.0%)
12 / 122 (9.8%)
0 / 2 (0%)
12 / 124 (9.7%)
X 2 = 5.67; df = 1;
P = 0.017
X 2 = 4.04; df = 1;
P = 0.044
9 / 40 (22.5%)
24 / 153 (15.7%)
33 / 193 (17.1%)
12 / 52 (23.1%)
3 / 24 (12.5%)
15 / 76 (19.7%)
11 / 123 (8.9%)
0 / 2 (0%)
11 / 125 (8.8%)
X 2 = 8.01; df = 2;
P = 0.018
X 2 = 0.163; df = 1;
P = 0.686
X 2 = 5.78; df = 2;
P = 0.056
Molecular and chromosomal forms and P. Falciparum infection
P. falciparum CSP positivity of Anopheles gambiae s.s. identified to both molecular and chromosomal form. Status as M molecular form and Mopti chromosomal form was significantly associated with CSP positivity in October 2009 ( P = 0.001)
No. CSP Pos. / No. Screened (% Pos.)
Among mosquitoes identified as Mopti chromosomal and M molecular forms, 15.8% (6/38) and 26.3% (5/19) were CSP positive in Kela and Sidarebougou, respectively. Among mosquitoes identified as Bamako chromosomal and S molecular forms in Kela, 9.4% (11/106) were CSP positive, whereas none (0/2) with this chromosomal and molecular form combination were CSP positive in Sidarebougou. Among mosquitoes identified as Savanna chromosomal and S molecular forms in Kela, 50.0% (5/10) were CSP positive, while 13.9% (15/108) of this form combination were CSP positive in Sidarebougou. These differences were significant in Kela (X 2 = 13.4; df = 2; P < 0.001), but not in Sidarebougou (X 2 = 1.88; df = 1; P = 0.170). On average across villages, 19.3% (11/57) of Mopti M forms were CSP positive, 16.9% (20/118) of Savanna S forms were CSP positive, and 9.2% (11/119) Bamako S forms were positive and these patterns were not significant (X2 =4.32, df = 2, P = 0.115).
An average of 12.2% of An. gambiae s.s. resting indoors were P. falciparum CSP positive from southern Mali in October 2009. Both molecular forms were CSP ELISA positive and there was no differential infection rate among molecular forms. Significantly more Mopti chromosomal forms (30.4%) were positive than were the Savanna (15.1%) and Bamako (9.2%) chromosomal forms. As the Mopti chromosomal form corresponds to M molecular form in Mali in most cases , finding that both M molecular and Mopti chromosomal forms were significantly associated with P. falciparum infection is not surprising. Site-specific differences in the number of CSP positive chromosomal form infection between Kela and Sidarebougou were also observed. In particular, the most common chromosomal form in each village, Bamako in Kela and Savanna in Sidarebougou, was least likely to be CSP positive.
The insignificant infection prevalence in the M molecular form in southern Mali corroborates with other studies from Cameroon and Senegal that reported no differences in P. falciparum infection between M and S molecular forms [6, 46]. A recent P. falciparum susceptibility assay among An. gambiae s.s. molecular forms from Senegal found significantly higher numbers of P. falciparum oocysts and sporozoites in the S molecular form than in the M form . This study analysed field-collected specimens of an unknown-age structure that were naturally infected with P. falciparum, whereas the Senegal study  collected eggs from the field and allowed the surviving adults to feed directly on a membrane with P. falciparum to standardize age and potential for infection. These conflicting findings may result from the origin of field collected samples (Mali vs. Senegal), to different techniques (natural vs. artificial infection), or to a differences in the age structure of the samples. Field studies from multiple sites and over multiple sampling periods are necessary to confirm the observed patterns.
Cryptic genetic differences in An. gambiae s.s. among sample sites can also limit comparisons among the previous studies [6, 46] and the present study. Genetic subdivisions beyond the M and S form designations have been reported and it is possible that differences among these subdivisions include genes associated with differential response to parasite infection. For example, recent studies in Cameroon demonstrated a subdivision in the M molecular form into discrete Forest-M, characterized as M molecular form and Forest chromosomal form (fixed for standard gene arrangement) and Mopti-M populations with typical Mopti karyotypes [12, 21]. In analyses using SNPs from immune signaling genes, three genetically distinct An. gambiae s.s. populations were observed in Mali: the M molecular form, the S molecular form (S1), and a subdivided S Pimperena form (S2) . Further, SNPs associated with P. falciparum infection were differentially distributed among M, S1, and S2 populations . Of interest, data presented here is similar to that reported in Riehle et al where a cryptic subgroup of An. gambiae, indistinguishable in molecular form but distinguishable via microsatellites amplified from chromosome 3, was susceptible to P. falciparum.
Differences in the local environment may likewise affect associations between An. gambiae forms and P. falciparum infection. For example, Dolo et al demonstrated that irrigated zones of Mali allowed for constant CSP positivity across seasons along with low human blood feeding and sporozoites indices, whereas in the non-irrigated zones, CSP positivity fluctuated seasonally, being high in the wet season and low in the dry season. Dolo et al hypothesized that malaria prevalence in villages adjacent to irrigated rice fields is consistently low in this environment because adult density is inversely related to blood feeding due to high mosquito densities driving villagers to protect themselves with repellants and bed nets [49, 50]. The Bamako chromosomal and S molecular forms were dominant in Kela (~3 km to a river), whereas the Savanna chromosomal form and S molecular form predominated in Sidarebougou (~10 km to agriculture fields). Kela is located close to a river that has the ability to flood and create additional oviposition sites not in the dry season likely increasing mosquito densities (11.2% prevalence), whereas in Sidarebougou mosquitoes densities are likely dependent on the wet season (13.9% prevalence). These habitat differences could contribute to the genetic variation (and potentially phenotypic variation) observed at different locations and, as Dolo et al hypothesized, habitat may play a role in the vector ecology of An. gambiae. Collectively, these studies highlight the importance of both genetic and environmental determinants of susceptibility to infection.
There were statistically significant P. falciparum differential infection rates among chromosomal forms and trends among chromosome inversions. Mopti had the highest CSP positivity (30.4%), followed by Savanna (15.1%) and Bamako (9.2%) forms. The data presented here indicated that Mopti chromosomal forms (2Rbc/u) were more likely to be positive for P. falciparum CSP. A previous study in Kenya identified a significant association between the total number of inversions and a decreased likelihood for P. falciparum infection . Data presented here did not show this specific association to be statistically significant, but standard chromosomal arrangements tended to increase the likelihood of being CSP positive. In particular, in Kela where the Bamako form was dominant and least likely to be positive, mosquitoes with standard or heterozygous arrangements were more likely to be CSP positive than mosquitoes homozygous for 2Rjcu and 2Rjbcu (Bamako form) (Table 3).
A number of studies have examined associations of chromosomal polymorphisms with malaria infection. A small region of chromosome 2 L has been associated with infection susceptibility, regardless of P. falciparum genotype . Genes within this region encode for melanization or parasite encapsulation . Within the 2La region, the APL1 gene, which encodes for natural resistance to P. falciparum, exhibited extremely low genetic diversity within the M molecular form, but high diversity in the S molecular form that may have arisen from larval infection [26, 35]. Alternatively, higher diversity at the APL1 locus in the S molecular form may be associated with a more diverse array of responses to P. falciparum and reduced susceptibility to infection. Similar results were identified within the anti-parasite and anti-bacterial gene TEP1r. Specifically, TEP1r was diverged between the M and S molecular forms and one variant showed a strong association with resistance to infection with a rodent malaria parasite and with P. falciparum. Additional studies comparing An. gambiae molecular and chromosomal forms with P. falciparum infection that incorporate An. gambiae speciation, genetic diversity at immune loci (e.g., TEP1r, APL1, Toll5B) as well as larger temporal and spatial scales may help to extend findings reported in this study.
Whilst the correlation between M and Mopti forms was expected, higher infection prevalences in the Mopti chromosomal form have not been demonstrated previously. In general, significant differences in P. falciparum infection prevalence at geographic locations with multiple molecular and chromosomal forms are likely due to discrepancies in the relative abundance of forms, genetic diversity in immune signaling genes within different forms, age structure of field collections, and local environmental variations that influence infection and transmission success.
We are grateful to the field collectors and the Mali Malaria Research and Training Center (MRTC-Bamako) for support and assistance during field collections. We would like to thank Drs. Clare Marsden and Michelle Sanford (Dept. Pathology, Microbiology, and Immunology, School of Veterinary Medicine, University of California Davis) for edits in the initial drafts of the manuscript. We would like to acknowledge Sarah Han, Saman Mahmood, and Allison Weakley, (Dept. Pathology, Microbiology, and Immunology, School of Veterinary Medicine, University of California Davis) for helping with additional molecular form identifications. This research was funded by NIH 1R01 AI078183 to SL and GCL. RTF is supported by the NIH T32 AI074550 Training Grant.
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