High genetic differentiation between the M and S molecular forms of Anopheles gambiae in Africa.
ABSTRACT Anopheles gambiae, a major vector of malaria, is widely distributed throughout sub-Saharan Africa. In an attempt to eliminate infective mosquitoes, researchers are trying to develop transgenic strains that are refractory to the Plasmodium parasite. Before any release of transgenic mosquitoes can be envisaged, we need an accurate picture of the differentiation between the two molecular forms of An. gambiae, termed M and S, which are of uncertain taxonomic status.
Insertion patterns of three transposable elements (TEs) were determined in populations from Benin, Burkina Faso, Cameroon, Ghana, Ivory Coast, Madagascar, Mali, Mozambique, Niger, and Tanzania, using Transposon Display, a TE-anchored strategy based on Amplified Fragment Length Polymorphism. The results reveal a clear differentiation between the M and S forms, whatever their geographical origin, suggesting an incipient speciation process.
Any attempt to control the transmission of malaria by An. gambiae using either conventional or novel technologies must take the M/S genetic differentiation into account. In addition, we localized three TE insertion sites that were present either in every individual or at a high frequency in the M molecular form. These sites were found to be located outside the chromosomal regions that are suspected of involvement in the speciation event between the two forms. This suggests that these chromosomal regions are either larger than previously thought, or there are additional differentiated genomic regions interspersed with undifferentiated regions.
Article: Current distribution of a pyrethroid resistance gene (kdr) in Anopheles gambiae complex from west Africa and further evidence for reproductive isolation of the Mopti form.[show abstract] [hide abstract]
ABSTRACT: In the field, the kdr mutation, involved in pyrethroid resistance, has been found widely distributed in the Savanna form of Anopheles gambiae s.s., but never in wild populations of the Mopti form or An. arabiensis, even in areas where both occur in sympatry with resistant Savanna populations. Under laboratory conditions, Mopti and Savanna forms were fully able to interbreed and the kdr mutation was transmissible from one form to the other. Both forms appeared to be exposed to pyrethroid selection pressure in the field. The absence of the kdr mutation in the Mopti form and the total lack of Mopti-Savanna heterozygotes in field populations provides further evidence of a pre-copulatory barrier to gene flow between these two forms. Molecular markers, including kdr, are powerful tools for studying population genetics and circulation of resistance genes, and should be used through an integrated approach for a better understanding of the speciation process.Parassitologia 10/1999; 41(1-3):319-22.
[show abstract] [hide abstract]
ABSTRACT: No longer a major public health concern in developed countries, malaria kills 1-3 million people annually, mostly children under the age of five in sub-Saharan Africa. In 1998, the WHO launched the Roll Back Malaria (RBM) drive to halve malaria mortality by 2010. This article contrasts the problems confronting RBM with the successful Italian drive to eradicate malaria between the late 19th and mid 20th centuries. The Italians employed education and applied socio-political will; however, ecological and socio-economic conditions in sub-Saharan Africa are more hospitable to the disease. RBM strategies should consider the Italian experience while awaiting a major scientific breakthrough necessary to achieve success.Health & Place 04/2005; 11(1):67-73. · 2.67 Impact Factor
[show abstract] [hide abstract]
ABSTRACT: Restrictions to gene flow among molecular forms of the mosquito Anopheles gambiae sensu stricto reveal an ongoing speciation process affecting the epidemiology of malaria in sub-Saharan Africa.Science 11/2002; 298(5591):115-7. · 31.20 Impact Factor
High Genetic Differentiation between the M and S
Molecular Forms of Anopheles gambiae in Africa
Caroline Esnault1, Matthieu Boulesteix1, Jean Bernard Duchemin2, Alphonsine A. Koffi3, Fabrice
Chandre4, Roch Dabire ´5, Vincent Robert6,7, Fre ´de ´ric Simard8,9, Fre ´de ´ric Tripet10, Martin J. Donnelly11,
Didier Fontenille9, Christian Bie ´mont1*
1Laboratoire de Biome ´trie et Biologie Evolutive (UMR 5558), CNRS, Universite ´ de Lyon, Universite ´ Lyon1, Villeurbanne, France, 2Centre de Recherche Me ´dicale et
Sanitaire (CERMES), Re ´seau International de l’Institut Pasteur, Niamey, Niger, 3Institut Pierre Richet, Institut National de Sante ´ Publique, Abidjan, Co ˆte d’Ivoire, 4Unite ´ de
Recherche 016, Institut de Recherche pour le De ´veloppement (IRD), CREC, Cotonou, Be ´nin, 5Institut de Recherche en Sciences de la Sante ´ (IRSS), Bobo Dioulasso, Burkina
Faso, 6Unite ´ de Recherche 77, Institut de Recherche pour le De ´veloppement (IRD), Unite ´ Scientifique du Muse ´um 504, Muse ´um National d’Histoire Naturelle, Paris,
France, 7Institut Pasteur, Antananarivo, Madagascar, 8Laboratoire de Recherche sur le Paludisme, Organisation de Coordination pour la lutte contre les Ende ´mies en
Afrique Centrale (OCEAC), Yaounde ´, Cameroun, 9Unite ´ de Recherche 016, Institut de Recherche pour le De ´veloppement (IRD), Montpellier, France, 10Centre for Applied
Entomology and Parasitology, School of Life Sciences, Keele University, Staffordshire, United Kingdom, 11Vector Group, Liverpool School of Tropical Medicine, Pembroke
Place, Liverpool, United Kingdom
Background: Anopheles gambiae, a major vector of malaria, is widely distributed throughout sub-Saharan Africa. In an
attempt to eliminate infective mosquitoes, researchers are trying to develop transgenic strains that are refractory to the
Plasmodium parasite. Before any release of transgenic mosquitoes can be envisaged, we need an accurate picture of the
differentiation between the two molecular forms of An. gambiae, termed M and S, which are of uncertain taxonomic status.
Methodology/Principal Findings: Insertion patterns of three transposable elements (TEs) were determined in populations
from Benin, Burkina Faso, Cameroon, Ghana, Ivory Coast, Madagascar, Mali, Mozambique, Niger, and Tanzania, using
Transposon Display, a TE-anchored strategy based on Amplified Fragment Length Polymorphism. The results reveal a clear
differentiation between the M and S forms, whatever their geographical origin, suggesting an incipient speciation process.
Conclusions/Significance: Any attempt to control the transmission of malaria by An. gambiae using either conventional or
novel technologies must take the M/S genetic differentiation into account. In addition, we localized three TE insertion sites
that were present either in every individual or at a high frequency in the M molecular form. These sites were found to be
located outside the chromosomal regions that are suspected of involvement in the speciation event between the two
forms. This suggests that these chromosomal regions are either larger than previously thought, or there are additional
differentiated genomic regions interspersed with undifferentiated regions.
Citation: Esnault C, Boulesteix M, Duchemin JB, Koffi AA, Chandre F, et al. (2008) High Genetic Differentiation between the M and S Molecular Forms of Anopheles
gambiae in Africa. PLoS ONE 3(4): e1968. doi:10.1371/journal.pone.0001968
Editor: Philip Awadalla, University of Montreal, Canada
Received December 21, 2007; Accepted March 7, 2008; Published April 16, 2008
Copyright: ? 2008 Esnault et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the PAL+ program of the french Ministry of Research and CNRS.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: firstname.lastname@example.org
Malaria causes the deaths of more than one million people each
year, mostly in Africa (WHO/UNICEF World Malaria Report
2005). This disease and the relevant mortality are due to one of
four Plasmodium species, which are transmitted by mosquitoes.
Anopheles gambiae is the major vector in sub-Saharan Africa, which
has the greatest disease burden. Various methods have been
developed to control mosquitoes. However, the failure of
traditional measures together with the spread of insecticide-
resistance in natural vector populations [1,2], have spurred on
attempts to find alternative, unconventional approaches. One of
the most innovative strategies sets out to replace the entire wild
populations of An. gambiae with genetically modified, Plasmodium-
resistant individuals. This idea seems more plausible following the
successful genetic transformation of some anopheline species,
including An. gambiae, and the identification of putative target
genes and gene drive mechanisms . To ensure that the
transgene spreads throughout the entire wild populations,
however, we need to understand the population structure and
level of gene flow of mosquito populations. This makes it very
important to know whether the genetic differentiation between the
two ‘molecular forms’ of An. gambiae, termed M and S, which are
suspected of currently undergoing speciation [4–6], is a general
phenomenon affecting all African populations. The distinction
between the two forms was primarily based on sequence
polymorphism in ribosomal DNA loci , which was subsequently
confirmed by microsatellite data in Cameroon  and Mali , by
the insertion patterns of various transposable elements (TEs) in
Cameroon  and of the short interspersed nuclear elements
(SINEs) Maque and SINE200 in Burkina Faso, Central African
Republic, Mali, and Kenya [11,12]. The kdr allele, which confers
PLoS ONE | www.plosone.org1April 2008 | Volume 3 | Issue 4 | e1968
knock-down resistance to pyrethroid insecticides and dichlorodi-
phenyltrichloroethane (DDT), was mainly present in S individuals,
and so this too was segregated between the two molecular forms
[see 4, 12 for reviews]. Studies of the gene flow within and between
the two molecular forms revealed, however, complex patterns of
differentiation. Some analyses revealed greater differences be-
tween ecological zones , and between allopatric populations of
a given molecular form than between the M and S populations
, suggesting that the M and S speciation is not yet complete.
Some data suggest that islands of speciation are present within the
genomes of these two forms, mostly in the region near the
centromeres of the X and 2L chromosomes and in a region of the
2R chromosome, whereas genetic differentiation remains weak in
other regions of the genome [15–18]. This could explain why the
estimates of genetic differentiation between the M and S forms
vary depending on the type of markers used, and the location of
the markers in the genome [8,9,13,19]. To find out whether the
genetic differentiation between the M and S molecular forms is
found throughout the geographical range of An. gambiae, we
studied the insertion polymorphism of three TEs. Because the
insertion sites of these TEs were scattered throughout the An.
gambiae genome, this study provides an overview of a large portion
of the mosquito genome. The insertion patterns of these TEs
reveal clear differentiation between the M and S forms, whatever
the geographical origin of the populations.
Results and Discussion
Twenty-one An. gambiae populations from ten African countries
were studied: two populations from Benin, two from Burkina Faso,
two from Cameroon, three from Ghana, five from Ivory Coast,
one from Madagascar, one from Mali, one from Mozambique,
three from Niger, and one from Tanzania (see Fig. 1, which also
indicates the number of mosquitoes of the M and S molecular
forms in each population). The M and S forms were distinguished
on the basis of their rDNA sequence polymorphism. The non-
Long Terminal Repeat (LTR) retrotransposon Aara8, the LTR
retrotransposon Ozymandias, and the DNA transposon Crusoe were
taken into consideration . Individual TE insertion profiles
were obtained by the Transposon Display method [20,21]. This
technique is very similar to the Sequence-Specific Amplification
Polymorphism, except that the PCR amplifies a DNA sequence
defined by one primer anchoring to a conserved region of the TE,
and another primer anchoring to an adaptor attached to flanking
sites generated by enzymatic restriction digestion. The presence
and absence of TE insertions can thus be scored in individuals
(Fig. 2). We therefore compared the TE insertion profiles of
individuals from all the populations by estimating the inter-
population differentiation indices, Wst. This parameter is analo-
gous to Fst, and can be used to analyze the presence/absence data
 obtained by Transposon Display. The mean Wst values
between the M and S populations (0.5760.07 for Aara8,
0.1960.04 for Ozymandias, 0.2360.05 for Crusoe), are higher than
the Wst between M populations (0.1260.06, 0.0760.04, and
0.1160.07, for Aara8, Ozymandias, and Crusoe, respectively) or S
populations (0.1060.06, 0.1260.09, and 0.1260.07, for Aara8,
Ozymandias, and Crusoe, respectively) (see Table S1 for the between-
population Wst and the associated P-values). A graphical
representation using a Principal Coordinate Analysis (PCoA)
(Fig. 3–5) clearly distinguishes between individuals of these two
forms, whatever the TE and the population considered, and shows
that individuals of a given molecular form cluster together. This
similarity of the results for all three TEs is reinforced by significant
Pearson correlation coefficient values between the Wst values
obtained for the three TEs (r=0.72 for Aara8 vs Ozymandias;
r=0.70 for Aara8 vs Crusoe; r=0.77 for Ozymandias vs Crusoe; P-
values,161024). These data thus clearly reveal a high degree of
differentiation between the M and S molecular forms in all 21
populations studied, with some ‘‘specific’’ TE insertion sites being
present at high frequency in one or other form (Table 1). Among
20 such sites, 6 were found in all individuals (4 on M and 2 on S),
whereas 14 were present at high frequency in one form or the
other (see Fig 6). To check for possible differentiation between
populations of the M or S forms, we did Principal Coordinate
Analysis (PCoA) on either the M or the S populations. No
structuration between populations of either the M or S form was
detected, which suggests the absence of specific insertion sites for
groups of populations apart from the M and S forms. This
indicates that most insertion sites were widespread and highly
polymorphic between populations.
To localize the specific TE insertions on the chromosome arms
of An. gambiae, we extracted the corresponding33P labeled bands
from polyacrylamide gels, and sequenced the DNA to make sure
that the bands corresponded to the expected TEs and to obtain the
sequences flanking the TEs. Among the 20 specific insertion sites
that were attempted to be sequenced (13 on M and 7 on S), 7 were
not isolated, 7 were isolated but were found to be integrated within
repeated sequences or transposable elements and could not be
localized, 3 were located in the ‘‘unannotated’’ chromosome. This
suggests that some of these insertions were embedded within the
heterochromatin or were inserted within nests of TEs, which could
themselves be heterochromatic. The localization of some of the
TEs specific to one of the molecular forms within the
heterochromatin, raises the important possibility that drastic
differences in the composition of heterochromatin may exist
between populations, and the question of the influence of
heterochromatin on genetic differentiation and speciation pro-
cesses, once again highlighting the need for more intensive
research on this particular genomic region . Three of the
specific insertions were however unambiguously localized on
chromosomes. They consisted of two Crusoe (Crusoe-1, Crusoe-2) and
one Ozymandias (Ozym-1) insertions specific to the populations of
the M molecular form. These insertions were localized in the
division 21 of the 2L chromosome (outside the known inversions),
and division 16 of the 2R chromosome (outside the 2Rd inversion,
at 800 kb from the inversion breakpoint) for Crusoe, and division 33
of the 3R chromosome for Ozymandias. These locations are outside
the genomic regions previously identified as being genetically
differentiated in the M and S forms [4,8,9,16,17,19] (see Fig. 7),
suggesting either that the chromosomal regions involved in this
differentiation are more extensive than expected, or that there are
additional differentiated regions interspersed with undifferentiated
regions. More detailed analyses of these regions are necessary. It
has been shown that differential population adaptation can be
determined from a subset of genes while gene flow still exists
between the species under speciation [24,25]. The ‘‘islands of
speciation’’ that define the M and S forms may thus be
extending gradually, reducing gene flow and fixing some TE
insertions close to the selected islands. Because the three
localized insertions were outside the known inversions and not
in the ‘‘islands of speciation’’, these sites could result simply
from genetic drift that has occurred after the separation of the
M and S forms. If so, the fixed sites and the sites at high
frequency would correspond to the sites of high occupancy
frequency in the original founders, and the polymorphic
insertion sites (sites with low occupancy frequency) would
correspond to more recent transposition events, as observed in
colonizing species . This kind of insertion site frequency
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org2 April 2008 | Volume 3 | Issue 4 | e1968
pattern is compatible with the idea that An. gambiae has speciated
or differentiated relatively recently . According to the
hypothesis of founder effects, the presence of fixed TE insertion
sites in each molecular form could suggest that gene flow is more
restricted than it has usually been thought to be, which would be
consistent with the virtual absence of hybrids in nature [8,
although an unusual frequency of hybrids was found in a
population from Guinea Bissau; J. Pinto, personal communica-
tion]. However, among the 6 fixed sites that we sequenced, only
one (Crusoe-1) was localized on the chromosome arms, the others
were either on the unknown chromosome, or clearly embedded
within heterochromatin or other transposable elements. In
addition, Crusoe-1, which is fixed in the M form, reaches a
frequency of 0.35 in the S form, suggesting it had been a site of
high frequency in the initial population from which the M and S
forms both derive. The two other localized sites, Ozym-1 and
Crusoe-2, which were outside the known inversions and the
‘‘islands of speciation’’, were present at an intermediate
frequency in the M populations (Fig 6), but at very low
frequency in the S form. All these data are in agreement with
founder events (26, 28) and then global expansion in Africa.
The wide distribution of An. gambiae suggests the possibility of
population adaptations to local climatic conditions, resulting in
local differentiation between populations of a same molecular
form, as has indeed been observed for M populations in
Cameroon and Mali, in addition to the M and S molecular form
differentiation . Both the M and S forms exist in Western
Africa, while only the S form has been found in Eastern Africa,
which implies that the S form has greater climatic adaptability or
migratory capacities than the M form. Although these two forms
may coexist in the same area, they appear to be in the process of
incipient speciation throughout Africa. No differences have been
observed in Plasmodium infection rates between sympatric M and S
forms in Cameroon . Therefore, any attempt to construct a
genetically-modified, Plasmodium-resistant mosquito, with the
intention of replacing natural, infected populations, or any other
strategy of controlling An. gambiae, will have to take this incipient
speciation between the M and S molecular forms of the mosquito
Materials and Methods
A total of 446 An. gambiae mosquitoes (257 M and 189 S) were
sampled from 21 sites in Africa: Ladji (6u219 N, 2u279 E), Lema
(7u469 N, 2u149 E) in Benin, Valle ´e du Kou (VK7, 11u249 N, 4u249
W), Soum (12u359 N, 2u179 E) in Burkina Faso, Simbok (3u499 N,
11u289 E), Ipono (2u229 N, 9u499 E) in Cameroon, Bonia (10u529
N, 1u079 W), Mampong (5u249 N, 0u369 W), Odumasy (5u539 N,
0u019 W) in Ghana, Azureti (5u129 N, 3u469 W), M’be (7u149 N,
5u19 W), Niamoue (5u529 N, 4u499 W), Nieky (5u249 N, 4u169 W),
Yaokoffikro (7u119 N, 5u19 W) in Ivory Coast, Banizoumbou
(13u329 N, 2u409 E), Kosseye (13u319 N, 2u19 E), Zindarou (13u269
N, 2u559 E) in Niger, Beforona (18u589 S, 48u159 E) in
Madagascar, Bankoumana (12u509 N, 5u479 W) in Mali, Furvela
(23u439 S, 35u189 E) in Mozambique, and Zenet (5u169 S, 38u369
E) in Tanzania. The species and molecular form of the specimens
were identified using Fanello et al.’s protocol .
Figure 1. Geographic origin of the African An. gambiae populations. Sample sizes of the M and S molecular forms in each population are
indicated in gray and black, respectively. Az=Azureti, Ban=Bankoumana, Baniz=Banizoumbou, Bef=Beforona, Bon=Bonia, Fur=Furvela,
Ipo=Ipono, Kos=Kosseye, Lad=Ladji, Lem=Lema, Mam=Mampong, M’be=M’be, Nia=Niamoue, Nie=Nieky, Odu=Odumasy, Sim=Simbok,
Soum=Soum, VK7=Valle ´e du Kou, Yao=Yaokoffikro, Zen=Zenet, Zin=Zindarou. The populations were from B=Benin, BF=Burkina Faso,
C=Cameroon, G=Ghana, IC=Ivory Coast, MG=Madagascar, ML=Mali, MZ=Mozambique, N=Niger, T=Tanzania.
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org3 April 2008 | Volume 3 | Issue 4 | e1968
Total genomic DNA was isolated from individual mosquitoes
using a standard phenol-chloroform extraction procedure after
proteinase K digestion. The Transposon Display was performed
using a modified version of the protocol used by Zampicinini et al.
, as follows: 50 to 100 ng of genomic DNA was digested with
10 units of HhaI for 6 hours at 37uC; during the first round of
amplification, 3 mM of MgCl2 and 0.625 Units of Taq
Polymerase were used; during the second amplification run,
0.2 mM of adaptor primer, 0.05 mM of nested TE-specific primer
with HEX fluorescent labeling, 2.5 mM of MgCl2and 0.625 Units
of Taq Polymerase were used. The last steps of the nested-
amplification cycles lasted 45 sec, instead of 1 min. The sequences
of adaptors and primers are shown in Table S2. Negative controls
were performed using the adaptor-primer or the element specific-
The PCR products were diluted 5-fold, and 1 ml of the dilution
was loaded onto a MegaBace 1000 capillary sequencer (Amer-
sham BioSciences) with an ET900-ROX standard size marker
(Amersham BioSciences). Raw data were analyzed by Genet-
icProfiler software (Amersham BioSciences). To confirm whether
the amplified DNAs were identical to the expected TE product, 6–
8 fragments were cloned using the Topo TA cloning kit
(Invitrogen), following the Manufacturer’s instructions, and sent
to GenoScreen for sequencing. All analyzed fragments corre-
sponded to the expected TE.
Each band on the capillary gels was automatically ascribed a
molecular weight according to the DNA ladder, which was loaded
on each capillary. We assumed that the DNA bands with the same
molecular weight shared the same TE insertion. The individual
TE insertion patterns obtained from the Transposon Display were
thus recorded as a binary matrix of 0 and 1 denoting the absence
or presence of a given peak on the capillary gel, respectively. The
between-population genetic divergence, Wst, was calculated for
each pair of population samples for the three transposable
elements considered separately. This Wst, which allows for the
Figure 2. Example of individual TE profiles obtained by Transposon Display with Aara8, Ozymandias, and Crusoe. Each peak in the TE
profile corresponds to one TE insertion in the individual analyzed. Because the probability that two TE copies would be inserted independently at the
same site in two different individuals is negligible, fragments of the same size were assumed to be of identical descent, and each TE fragment of a
given size was considered to be a single insertion. The matrices of the presence/absence of peaks were used to estimate genetic distances and do the
PCoA analyses shown in Fig. 3, 4, and 5.
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org4 April 2008 | Volume 3 | Issue 4 | e1968
dominant nature of TE, is an analogue of the fixation index of
inter-population differentiation, FST[22,32]. Because the inter-
population index values calculated from samples consisting of less
than 5 individuals were not reliable, these values were not included
in the calculation of the mean Wst values between populations.
Graphical representations of the proximities between individuals
were obtained using a Principal Coordinate Analysis (PCoA), using
the R package ade4 . All individuals were included in these
analyses, because those from small samples were not expected to
bias the results, as they were not assigned a priori to any specific
population. For each population, we then drew the ellipses
centered on the gravity center of each scatterplot, with the size of
the two first axes equal to 1.5 times the standard deviation of the
coordinates of the projections on the axes. MANOVA between
molecular forms was performed using JMP Version 7 software
(SAS Institute Inc.), and the variance components were tested for
significance by nonparametric randomization tests with the null
hypothesis of no population structure.
The detection of population differentiation by the PCoA is
based on the sites that are either fixed or at high frequency in a
form and not in the other. Hence sites with very high insertion
polymorphism play no role in the differentiation.
Identification of transposable element insertion sites
Fragments obtained from the Transposon Display were
separated on a 6% denaturing polyacrylamide gel. Samples were
diluted with one volume of loading dye (95% formamide, 0.05%
xylene cyanol FF, and 0.05% bromophenol blue), heat denatured
at 95uC for 5 min, and immediately cooled on ice. Polyacryma-
mide gel was pre-run at 75 W for 30 min. Six ml of each sample
were run at 75 W for 4 h in 1xTBE. We used radioactive33P
labeling; the gel was transferred to Whatman 3 MM paper, and
vacuum dried at 65uC for 1 h; dried gels were exposed to X-ray
films overnight or for 48 h, depending on the signal intensity .
The fragments of interest were cut from the gels, the DNA was
eluted from the bands at 100uC for 15 min and resuspended in
150 ml of sterile water. The fragments were amplified according to
the second amplification run of the Transposon Display protocol,
and cloned using the Topo TA cloning kit (Invitrogen). About 5
clones for each fragment were sequenced by GenoScreen. The
genomic localizations of the sequenced DNAs were determined by
interrogation of the Anopheles gambiae genome database (Ensembl
AgamP3 assembly, release 46.3i). Among the 14 sequenced
fragments, only three presented a flanking sequence localized in
only one site on the chromosome arm. These three fragments
corresponded to two insertions of the DNA transposon Crusoe and
to one insertion of the LTR retrotransposon Ozymandias. Their
specificity to the M form was confirmed by PCR. Amplifications
were performed following the second amplification run of
Transposon Display, using primers Crusoe-1F 59-CCTATT-
Figure 3. Principal Coordinate Analyses (PCoA) of the Aara8
profiles obtained by the Transposon Display technique. The
ellipses, calculated for each population, were centered on the gravity
centre of each scatterplot, and the size of their axes was equal to 1.5
(the square root of the eigen values of the covariance matrix) times the
standard deviation of the coordinates of the projections on the first and
second axes. The percentage of variance explained by the first two axes
was 75.4%, and the F value of MANOVA test between individuals of
each molecular form was equal to 18.6 (p,0.001).
Figure 4. Principal Coordinate Analyses (PCoA) of the Ozyman-
dias profiles. The percentage of variance explained by the first two
axes was 55.1%, and the F value of MANOVA test between individuals
of each molecular form was equal to 4.3 (p,0.001).
Figure 5. Principal Coordinate Analyses (PCoA) of the Crusoe
profiles. The percentage of variance explained by the first two axes
was equal to 55.0% and the F value of MANOVA test between
individuals of each molecular form was equal to 4.1 (p,0.001).
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org5 April 2008 | Volume 3 | Issue 4 | e1968
Figure 6. Frequencies of the 20 insertion sites sequenced in the M (red squares), and S (green squares) molecular forms. Insertion
sites localized on the chromosomes: Crusoe-1-2, Ozym-1. Insertion sites localized on the unknown chromosome: Aara8-2-3-5. Insertion sites integrated
within other transposable elements or repeated sequences: Aara8-1-4-6, Ozym-2-5, Crusoe-3-4. Insertion sites not isolated: Aara8-7, Ozym-3-4-6-7,
Table 1. Number of insertion sites and average site numbers6SE of Aara8, Ozymandias, and Crusoe for the populations of the S
and M molecular forms.
Number of insertion sites Mean insertion sites number6SE
Specific to M
Specific to S
Total number of loci
with an insertion M molecular form S molecular form
Figure 7. Position on chromosomes 2 and 3 of the three TE insertions specific to the M molecular form (in box). The loci Ag2H325,
Ag2H417, Ag2H769, Ag3H555, Ag3H170, Ag3H750 from , kdr from , and Ion channel and LIM from , have been shown to differentiate the
two M and S forms in previous studies. The chromosomal inversions of the An. gambiae genome are indicated below the chromosome arms. The
GPRor39, GPRor38, and UNK1 loci, indicated by asterisks, have been shown to discriminate between the two forms only in Cameroon .
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org6April 2008 | Volume 3 | Issue 4 | e1968
GATTTGTCCGACACTG-39, Crusoe-1R 59-TCACTTCACGT-
CACTG-39, Crusoe-2R 59- TTTACCTGGC TTTTGGCAAT-39 and
Ozym-1F 59-TGCTATAAGCAATCCACCACA-39, Ozym-1R 59-
CTCAAAGTGTGCTTCCTCACC-39 for the Crusoe-1, Crusoe-2 and
Ozym-1 insertions, respectively.
Found at: doi:10.1371/journal.pone.0001968.s001 (1.58 MB DOC)
Found at: doi:10.1371/journal.pone.0001968.s002 (0.03 MB DOC)
We thank Abdoulaye Diabate for his help with collecting wild mosquitoes,
Emmanuelle Lerat and Cristina Vieira for their helpful comments, the
technical DTAMB platform at Lyon for providing access to the
MegaBACETMcapillary sequencer, Corinne Mhiri for her help with
sequencing of some specific TE insertions.
Conceived and designed the experiments: DF CB MD CE MB. Performed
the experiments: CE. Analyzed the data: CB CE. Contributed reagents/
materials/analysis tools: DF VR FS JD RD MD MB AK FC FT. Wrote
the paper: CB CE.
1. Chandre F, Manguin S, Brengues C, Dossou Yovo J, Darriet F, et al. (1999)
Current distribution of a pyrethroid resistance gene (kdr) in Anopheles gambiae
complex from west Africa and further evidence for reproductive isolation of the
Mopti form. Parassitologia 41: 319–322.
2. Amorosa LF Jr., Corbellini G, Coluzzi M (2005) Lessons learned from malaria:
Italy’s past and sub-Sahara’s future. Health Place 11: 67–73.
3. Boete C (2006) Genetically Modified Mosquitoes for Malaria Control. Austin,
Texas, U S A: Landes Biosciences. pp 174.
4. della Torre A, Costantini C, Besansky NJ, Caccone A, Petrarca V, et al. (2002)
Speciation within Anopheles gambiae–the glass is half full. Science 298: 115–117.
5. Fanello C, Petrarca V, della Torre A, Santolamazza F, Dolo G, et al. (2003) The
pyrethroid knock-down resistance gene in the Anopheles gambiae complex in Mali
and further indication of incipient speciation within An. gambiae s.s. Insect Mol
Biol 12: 241–245.
6. Wondji C, Simard F, Petrarca V, Etang J, Santolamazza F, et al. (2005) Species
and populations of the Anopheles gambiae complex in Cameroon with special
emphasis on chromosomal and molecular forms of Anopheles gambiae s.s. J Med
Entomol 42: 998–1005.
7. Favia G, Lanfrancotti A, Spanos L, Siden-Kiamos I, Louis C (2001) Molecular
characterization of ribosomal DNA polymorphisms discriminating among
chromosomal forms of Anopheles gambiae s.s. Insect Mol Biol 10: 19–23.
8. Wondji C, Simard F, Fontenille D (2002) Evidence for genetic differentiation
between the molecular forms M and S within the Forest chromosomal form of
Anopheles gambiae in an area of sympatry. Insect Mol Biol 11: 11–19.
9. Wang R, Zheng L, Toure YT, Dandekar T, Kafatos FC (2001) When genetic
distance matters: measuring genetic differentiation at microsatellite loci in
whole-genome scans of recent and incipient mosquito species. Proc Natl Acad
Sci U S A 98: 10769–10774.
10. Boulesteix M, Simard F, Antonio-Nkondjio C, Awono-Ambene HP,
Fontenille D, et al. (2007) Insertion polymorphism of transposable elements
and population structure of Anopheles gambiae M and S molecular forms in
Cameroon. Mol Ecol 16: 441–452.
11. Barnes MJ, Lobo NF, Coulibaly MB, Sagnon NF, Costantini C, et al. (2005)
SINE insertion polymorphism on the X chromosome differentiates Anopheles
gambiae molecular forms. Insect Mol Biol 14: 353–363.
12. della Torre A, Tu Z, Petrarca V (2005) On the distribution and genetic
differentiation of Anopheles gambiae s.s. molecular forms. Insect Biochem Mol Biol
13. Yawson AE, Weetman D, Wilson MD, Donnelly MJ (2007) Ecological zones
rather than molecular forms predict genetic differentiation in the malaria vector
Anopheles gambiae s.s. in Ghana. Genetics 175: 751–761.
14. Lehmann T, Licht M, Elissa N, Maega BT, Chimumbwa JM, et al. (2003)
Population Structure of Anopheles gambiae in Africa. J Hered 94: 133–147.
15. Stump AD, Fitzpatrick MC, Lobo NF, Traore S, Sagnon N, et al. (2005)
Centromere-proximal differentiation and speciation in Anopheles gambiae. Proc
Natl Acad Sci U S A 102: 15930–15935.
16. Turner TL, Hahn MW, Nuzhdin SV (2005) Genomic islands of speciation in
Anopheles gambiae. PLoS Biol 3: e285.
17. Turner TL, Hahn MW (2007) Locus- and population-specific selection and
differentiation between incipient species of Anopheles gambiae. Mol Biol Evol 24:
18. Slotman MA, Reimer LJ, Thiemann T, Dolo G, Fondjo E, et al. (2006) Reduced
recombination rate and genetic differentiation between the M and S forms of
Anopheles gambiae s.s. Genetics 174: 2081–2093.
19. Gentile G, Slotman M, Ketmaier V, Powell JR, Caccone A (2001) Attempts to
molecularly distinguish cryptic taxa in Anopheles gambiae s.s. Insect Mol Biol 10:
20. Van den Broeck D, Maes T, Sauer M, Zethof J, De Keukeleire P, et al. (1998)
Transposon Display identifies individual transposable elements in high copy
number lines. Plant J 13: 121–129.
21. Casa AM, Brouwer C, Nagel A, Wang L, Zhang Q, et al. (2000) The MITE
family heartbreaker (Hbr): molecular markers in maize. Proc Natl Acad Sci U S A
22. Peakall R, Smouse PE, Huff DR (1995) Evolutionary implications of allozyme
and RAPD variation in diploid populations of dioecious buffalograss Buchloe
dactyloides. Mol Ecol 4: 135–147.
23. Johnson L (2007) Transposon silencing: The extraordinary epigenetics of a
transposon trap. Heredity. doi:10.1038/sj.hdy.6801064.
24. Machado CA, Kliman RM, Markert JA, Hey J (2002) Inferring the history of
speciation from multilocus DNA sequence data: the case of Drosophila
pseudoobscura and close relatives. Mol Biol Evol 19: 472–488.
25. Wu C-I, Ting C-T (2004) Genes and speciation. Nature Rev Genet 5(2):
26. Garcia Guerreiro MP, Fontdevila A (2007) The evolutionary history of Drosophila
buzzatii. XXXVI. Molecular structural analysis of Osvaldo retrotransposon
insertions in colonizing populations unveils drift effects in founder events.
Genetics 175: 301–310.
27. Mukabayire O, Caridi J, Wang X, Toure ´ YT, Coluzzi M, Besansky NJ (2001)
Patterns of DNA sequence variation in chromosomally recognized taxa of
Anopheles gambiae: evidence from rDNA and single-copy loci. Insect Mol Biol 10:
28. Macpherson JM, Gonzalez J, Witten DM, Davis JC, Rosenberg NA, et al. (2008)
Nonadaptive explanations for signatures of partial selective sweeps in
Drosophila. Mol Biol Evol (in press).
29. Slotman MA, Tripet F, Cornel AJ, Meneses CR, Lee Y, et al. (2007) Evidence
for subdivision within the M molecular form of Anopheles gambiae. Mol Ecol 16:
30. Fanello C, Santolamazza F, della Torre A (2002) Simultaneous identification of
species and molecular forms of the Anopheles gambiae complex by PCR-RFLP.
Med Vet Entomol 16: 461–464.
31. Zampicinini G, Blinov A, Cervella P, Guryev V, Sella G (2004) Insertional
polymorphism of a non-LTR mobile element (NLRCth1) in European
populations of Chironomus riparius (Diptera, Chironomidae) as detected by
transposon insertion display. Genome 47: 1154–1163.
32. Excoffier L, Smouse PE, Quattro JM (1992) Analysis of molecular variance
inferred from metric distances among DNA haplotypes: application to human
mitochondrial DNA restriction data. Genetics 131: 479–491.
33. Chessel D, Dufour AB, Thioulouse J (2004) The ade4 package - I : One-table
methods. R news 4: 5–10.
34. Melayah D, Bonnivard E, Chalhoub B, Audeon C, Grandbastien MA (2001)
The mobility of the tobacco Tnt1 retrotransposon correlates with its
transcriptional activation by fungal factors. Plant J 28: 159–168.
Molecular Forms in An. gambiae
PLoS ONE | www.plosone.org7 April 2008 | Volume 3 | Issue 4 | e1968